calcium in living cells
TRANSCRIPT
Series Editors
Leslie WilsonDepartment of Molecular, Cellular and Developmental Biology
University of California
Santa Barbara, California
Paul MatsudairaDepartment of Biological Sciences
National University of Singapore
Singapore
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CONTRIBUTORS
Numbers in parentheses indicate the pages on which the authors’ contributions begin.
David L. Armstrong (183), Membrane Signaling Group, Laboratory of Neurobiology,National Institute of Environmental Health Sciences, NIH, Durham, North Carolina,USA
Darryl A. Auston (113), Center for Biomedical Engineering and Technology, andDepartment of Physiology, University of Maryland School of Medicine, Baltimore,Maryland, USA
S. Baudet (67), Ricerca Biosciences SAS, Saint Germain sur l’Arbresle, FranceD.M. Bers (67), Department of Pharmacology, University of California, Davis, Davis,California, USA
Donald M. Bers (1), Department of Pharmacology, University of California, DavisSchool of Medicine, Davis, California, USA
Francis Burton (225), School of Life Sciences, University of Glasgow, United KingdomChristian Erxleben (183), Membrane Signaling Group, Laboratory of Neurobiology,National Institute of Environmental Health Sciences, NIH, Durham, North Carolina,USA
L. Hove-Madsen (67), Cardiovascular Research Centre CSIC-ICCC, Hospital de laSanta Creu i Sant Pau, Barcelona, Spain
Joseph P.Y. Kao (113), Center for Biomedical Engineering and Technology, andDepartment of Physiology, University of Maryland School of Medicine, Baltimore,Maryland, USA
Eric Karplus (263), Science Wares Inc., Falmouth, Massachusetts, USAOle Johan Kemi (225), School of Life Sciences, University of Glasgow, UnitedKingdom
Gong Li (113), Center for Biomedical Engineering and Technology, and Department ofPhysiology, University of Maryland School of Medicine, Baltimore, Maryland, USA
Mark A. Messerli (91), BioCurrents Research Center, Cellular Dynamics Program,Marine Biological Laboratory, Woods Hole, Massachusetts, USA
Andrew L. Miller (263), Biochemistry and Cell Biology Section and State Key Labora-tory of Molecular Neuroscience, Division of Life Science, HKUST, Clear Water Bay,Kowloon, Hong Kong, PR China
Richard Nuccitelli (1), BioElectroMed Corp., Burlingame, California, USAChris W. Patton (1), Hopkins Marine Station, Stanford University, Pacific GroveCalifornia, USA
Taufiq Rahman (199), Department of Pharmacology, Tennis Court Road, Universityof Cambridge, Cambridge, United Kingdom
ix
x Contributors
Martyn Reynolds (225), Cairn Research Limited, Faversham, Kent, United KingdomKelly L. Rogers (263), The Walter and Eliza Hall Institute of Medical Research,
Parkville, AustraliaGodfrey Smith (225), School of Life Sciences, University of Glasgow, United KingdomPeter J. S. Smith (91), BioCurrents Research Center, Cellular Dynamics Program,
Marine Biological Laboratory, Woods Hole, Massachusetts, USAColin W. Taylor (199), Department of Pharmacology, Tennis Court Road, University
of Cambridge, Cambridge, United KingdomSarah E. Webb (263), Biochemistry and Cell Biology Section and State Key Laboratory
of Molecular Neuroscience, Division of Life Science, HKUST, Clear Water Bay,Kowloon, Hong Kong, PR China
Michael Whitaker (153), Institute of Cell and Molecular Biosciences, Medical School,Newcastle University, Framlington Place, Newcastle upon Tyne, United Kingdom
Jody A. White (183), Membrane Signaling Group, Laboratory of Neurobiology,National Institute of Environmental Health Sciences, NIH, Durham, North Carolina,USA
Robert Zucker (27), Molecular and Cell Biology Department, University of Californiaat Berkeley, Berkeley, California, USA
PREFACE
This volume of Methods in Cell Biology is a sequel to the oft-consulted Volume
40 of the series edited by Richard Nuccitelli that oVered a practical guide to the
study of calcium in living cells. Much in that volume remains relevant and this
volume oVers updates of chapters contributed to the original volume. But in the
decade and a half that have elapsed since the publication of Volume 40, as calcium
signaling has continued to find itself a ubiquitous element of cell regulation, new
technical advances have oVered themselves to the field and existing methods have
been refined.
This volume retains the bedrock of an understanding of calcium buVering and
the manipulation of intracellular free calcium concentration in cells: the subtleties
and peculiarities of an ion that acts at submicromolar concentrations and that is
very actively regulated by cellular buVers and pumps are covered extensively by the
early chapters on calcium buVers; a detailed treatment of dynamic changes in free
calcium achieved by the photosensitive release of calcium from buVers that under-go light-induced changes in calcium aYnity follows on.
Calcium-sensitive electrodes oVer the most quantitative approach to measuring
calcium concentrations within cells and in solutions. Two chapters in this volume
provide a deep understanding of both spatially homogeneous calcium sensing and
of the use of calcium-sensitive electrodes to measure standing fluxes and gradients
of calcium.
Calcium-sensitive fluorescent dyes present some advantages in measuring intra-
cellular free calcium over electrodes—what is lost in precision can be gained in
convenience and time resolution. The two chapters on calcium-sensitive fluores-
cent approaches cover low molecular mass indicators and the newer recombinant
techniques based on green fluorescent protein.
In many circumstances, particularly in studying neuronal calcium signaling in
individual neurones, patch clamp methods are king. Two chapters are devoted to
patch clamp analysis of calcium signaling. One of these concentrates on calcium
channels at the plasma membrane—an approach that remains key to understand-
ing neuronal signaling mechanisms; the other highlights the remarkable achieve-
ment of using patch clamp techniques to study both the aggregate and single
channel properties of calcium release channels in the membranes of intracellular
calcium stores.
The final two chapters of the volume explain the state-of-the-art in imaging
calcium signals. Confocal and multiphoton microscopy have much improved the
spatial and temporal resolution of the measurement of calcium signals, revealing,
among other things, how very localized calcium signals play a part in the versatile
xi
xii Preface
repertoire of this key second messenger. Low-intensity photon imaging using
aequorin has provided an approach best suited to the long-term recording of
calcium signals associated with cell division and pattern formation, situations in
which photobleaching and light-induced damage preclude the use of fluorescent
probes.
I thank all the authors of this volume for having made possible, as I see it, such a
valuable and detailed contribution to the methodological state-of-the-art in the
field. I also thank Zoe Kruze and Narmada Thangavelu of Elsevier for their help
and patience in bringing this volume into being.
Michael Whitaker
CHAPTER 1
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
A Practical Guide to the Preparation ofCa2þ BuVers
Donald M. Bers,* Chris W. Patton,† and Richard Nuccitelli‡
*Department of PharmacologyUniversity of California, DavisSchool of Medicine Davis, California, USA
†Hopkins Marine StationStanford University, Pacific GroveCalifornia, USA
‡BioElectroMed Corp.Burlingame, California, USA
A
OGY,All rig
bstract
VOL. 99 0091hts reserved. 1 DOI: 10.1016/S0091
-679X-679X
I. In
troduction II. R ationaleA.
Which Ca2þ BuVer Should You Use? B. EGTA: The Workhorse of Biological Ca2þ Chelators C. BAPTA Family of Ca2þ BuVersIII. M
ethods A. Basic Mathematical Relationships B. Temperature, Ionic Strength, and pH CorrectionsIV. M
aterials A. [Ca2þ] Measurement and Calibration Solutions B. Preparing BuVer Solution C. Software ProgramsV. D
iscussion and Summary R eferencesAbstract
Calcium (Ca2þ) is a critical regulator of an immense array of biological
processes, and the intracellular [Ca2þ] that regulates these processes is �10,000
lower than the extracellular [Ca2þ]. To study and understand these myriad Ca2þ-dependent functions requires control and measurement of [Ca2þ] in the nano- to
micromolar range (where contaminating Ca2þ is a significant problem). As with
/10 $35.00(10)99001-8
2 Donald M. Bers et al.
pH, it is often essential to use Ca2þ buVers to control free [Ca2þ] at the desired
biologically relevant concentrations. Fortunately, there are numerous available
Ca2þ buVers with diVerent aYnities that make this practical. However, there are
numerous caveats with respect to making these solutions appropriately with
known Ca2þ buVers. These include pH dependence, selectivity for Ca2þ (e.g., vs.
Mg2þ), ionic strength and temperature dependence, and complex multiple equili-
bria that occur in physiologically relevant solutions. Here we discuss some basic
principles of Ca2þ buVering with respect to some of these caveats and provide
practical tools (including freely downloadable computer programs) to help in the
making and calibration of Ca2þ-buVered solutions for a wide array of biological
applications.
I. Introduction
Cell biologists quickly learn how important it is to control the ionic composition
of the solutions used when studying cellular biochemistry, physiology, and phar-
macology. BuVering the pH of the solutions we use is so routine that one can
hardly imagine making a biological solution without the careful selection of the
appropriate pH buVer and measurement of pH in the resulting solution. Indeed,
there are an array of popular zwitterionic amino acid pH buVers introduced by
Good et al. (1966) that are in widespread use (e.g., HEPES) and which complement
the natural physiological pH buVers for these purposes. In contrast, there has been
less attention to buVering and measuring [Ca2þ] because extracellular [Ca2þ] levelsare typically in the millimolar range and such concentrations are easily measured
and prepared. However, intracellular [Ca2þ] ([Ca2þ]i) is quite another matter
because these levels are more typically in the 100 nM–10 mM range which is not
as easily prepared or measured. For example, your source of distilled water could
easily have trace Ca2þ contamination in the range of 1–10 mM. This range of
contaminant Ca2þ can also come from chemicals and biochemicals commonly
used to make solutions. Additionally, there is often a considerable amount of
endogenous Ca2þ in biological tissue or cell samples which is not easily removed
or controlled. Therefore, when we are interested in studying intracellular reactions,
Ca2þ buVering is extremely important.
In this chapter, we will present a practical guide to the preparation of Ca2þ
buVer solutions. Our goal is to emphasize the methods and important variables to
consider while making the procedure as simple as possible. We will also introduce
computer programs which may be of practical use to many workers in this field.
One is a spreadsheet useful in making and validating simple Ca2þ calibration
solutions. The others are more powerful and extensive programs for the calcula-
tion of [Ca2þ] (and other metals and chelators) in complex solutions with multiple
equilibria. These programs have been developed and described with maximum ease
of use in mind.
Table IMixed stability co
Ca2þ buVera
CDTA
EGTA
Quin 2
BAPTA
Fura-2
Dibromo-BAPTA
4,40-Difluoro-BAPTA
Nitr-5 photolyzed
5-Methyl-50-nitro-BA5-Mononitro-BAPTA
NTA
ADA
Citrate
5,50-Dinitro-BAPTA
aAbbreviations: C
N-N-N0-N0-tetraacetiADA, acetamidomin
bMeasured at pH
1. A Practical Guide to the Preparation of Ca2þ BuVers 3
II. Rationale
A. Which Ca2þ BuVer Should You Use?
When selecting the appropriate Ca2þ buVer for your application, the main
consideration is to choose one with a dissociation constant (Kd) close to the desired
free [Ca2þ]. The ability of a buVer to absorb or release ions and thus to hold the
solution at a given concentration of that ion is greatest at its Kd. Just as you should
not choose PIPES (pKa¼6.8) to buVer a solution at pH 7.8, choosing a Ca2þ buVerwith a Kd far from the desired [Ca2þ] set point is a mistake. As a rule of thumb, the
buVer’s Kd should not lie more than a factor of 10 from your desired [Ca2þ].In addition, the buVer should exhibit a much greater aYnity for Ca2þ than Mg2þ
since intracellular [Mg2þ] is typically 10,000-fold higher than [Ca2þ]i. Fortunately,about a dozen suitable buVers are available spanning the range from 10 nM to
100 mM (Table I). There are also a large number of fluorescent Ca2þ indicators
(see Chapter 5) that can also serve as Ca2þ buVers, giving one the opportunity to
both buVer and measure free [Ca2þ] with the same reagent. We will not focus on
nstants for useful Ca2þ buVers at 0.15 M ionic strength in order of Ca2þ aYnity
log K0Ca Kd
K0Ca (pH 7.4)/
K0Ca (pH 7.0) K0
Ca /K0Mg (pH 7.4) References(pH 7.4)
7.90 13 nM 2.7 120 Martell and Smith (1974, 1977),
Bers and MacLeod (1988)
7.18 67 nM 6.2 72,202 Martell and Smith (1974, 1977),
Bers and MacLeod (1988)
6.84 144 nM 1.15 25,114 Tsien (1980)
6.71 192 nM 1.14 158,244 Tsien (1980)
6.61 242 nM 1.14 72,373 Grynkiewicz et al. (1985)
5.74 1.83 mM 1.02 63,000 Tsien (1980)
5.77 1.7 mMb – – Pethig et al. (1989)
5.2 6.3 mMb – – Tsien and Zucker (1986)
PTA 4.66 22 mMb – – Pethig et al. (1989)
4.4 40 mMb – – Pethig et al. (1989)
3.87 134 mM 2.5 8 Martell and Smith (1974, 1977),
Bers and MacLeod (1988)
3.71 191 mM 1.24 32 Nakon (1979)
3.32 471 mM 1.03 1.3 Martell and Smith (1974, 1977),
Bers and MacLeod (1988)
2.15 7 mMb – – Pethig et al. (1989)
DTA, cyclohexilinedinitrilo-N-N-N0-N0-tetraacetic acid; EGTA, Ethylene glycol bis (b-aminoethylester)
c acid; BAPTA, 1,2-bis(o-aminophenoxy)ethane-N-N-N0-N0-tetraacetic acid; NTA, nitriloacetic acid;
odiacetic acid.
7 and 0.1 M ionic strength.
4 Donald M. Bers et al.
these fluorescent indicators here, but they can be substituted for the buVers de-
scribed (especially when the fundamental binding properties have been measured).
Ethylene glycol bis(b-aminoethylether)-N,N,N0,N0-tetraacetic acid (EGTA) is one
of the best-known Ca2þ buVers, and it can be a reliable buVer in the range of
10 nM–1 mM [Ca2þ] at the typical intracellular pH of 7.2. However, if your goal is
tomake buVers in the 1–10 mMrange, BAPTA (1,2-bis(o-aminophenoxy)ethane-N,
N,N0,N0-tetraacetic acid) or dibromo-BAPTA (Br2-BAPTA) would be better
choices.
B. EGTA: The Workhorse of Biological Ca2þ Chelators
By far the most popular Ca2þ buVer has been EGTA. This molecule has been
used extensively because its apparent dissociation constant (Kd) at pH 7 (0.4 mM) is
close to intracellular Ca2þ levels and it has a much higher aYnity for Ca2þ than for
Mg2þ (�100,000 times higher around neutral pH). However, the preparation of
Ca2þ buVers using EGTA is complicated by the strong pH dependence of its Ca2þ
aYnity (see Fig. 1 and Table I). Thus, while the free [Ca2þ] would be about 400 nM
when EGTA is half saturated with Ca2þ at pH 7, the free [Ca2þ] in this same
solution would decrease by nearly 10-fold to 60 nM by simply raising the pH to
7.4! Therefore, the pH of Ca2þ buVers made with EGTA must be very carefully
controlled, and the calculation of the appropriate amounts of EGTA and Ca2þ to
use must be made at the desired pH. The purity of the EGTA is also a variable that
can cause substantial errors, as large as 0.2 pCa units in the free [Ca2þ] (Bers, 1982;Miller and Smith, 1984).
6 7 84
5
6
7
8
9 EGTA
BAPTA
Br2-BAPTA
0.001
0.01
0.1
1
10
pH
Log
K� C
a (a
ppar
ent C
a2+ a
ffini
ty)
Fre
e [C
a2+] (mM
) fo
r1
Ca
: 2 li
gand
rat
io
Fig. 1 The pH dependence of apparent aYnities (K0Ca) for EGTA, BAPTA, and Br2-BAPTA at 20 �C
and 150 mM ionic strength.
1. A Practical Guide to the Preparation of Ca2þ BuVers 5
There are many papers explaining how to calculate the proper amounts of
EGTA and Ca2þ that must be combined to obtain a given free [Ca2þ] (some are
listed below). Due to the steep pH dependence and slightMg2þ sensitivity, both pH
and Mg2þ must be considered in the calculation and it is best accomplished by
computer. We provide a program for such calculations and describe it below.
Systematic errors in EGTA purity and pH can be a real practical problem (Bers,
1982), even with the best calculations for solution preparation. Thus, we also
recommendmeasuring the free [Ca2þ] whenever possible (see below andChapter 3).
C. BAPTA Family of Ca2þ BuVers
Roger Tsien developed an analogue of EGTA in which the methylene links
between oxygen and nitrogen atoms were replaced with benzene rings to yield a
compound called BAPTA (Tsien, 1980; Fig. 2). This compound exhibits a much
lower pH sensitivity and much higher rates of calcium association and dissocia-
tion. These characteristics are mainly due to the fact that BAPTA is almost
completely deprotonated at neutral pH. Moreover, modifications of BAPTA
have been made to provide Ca2þ buVers with a range of Kd values covering the
biologically significant range of 0.1 mM–10 mM (see Table I; Pethig et al., 1989).
However, one disadvantage compared with EGTA is that the BAPTA family of
buVers exhibits a greater ionic strength dependence (see Figs. 3–5). In particular,
increasing ionic strength from 100 to 300 mM decreases the apparent aYnity
constant, K0Ca for BAPTA or Br2-BAPTA by almost threefold, whereas the
COO–
COO–
COO–
COO– –OOC
–OOC
–OOC
–OOC
N
NN
X XX = H; BAPTA
X = Br; Br2-BAPTA
N
OO
O O
EGTA
Fig. 2 Structural formulas for the Ca2þ chelators EGTA (top) and BAPTA and Br2-BAPTA
(bottom).
1 8 15 22 29 361
1.5
2
2.5
3
Temperature (�C)
Data
Prediction
K� C
a (in
106
M−1
)K
� Ca
(in 1
06 M
−1)
0 0.05 0.1 0.15 0.2 0.25 0.30.7
0.8
0.9
1
1.1
1.2
1.3
1.4
1.5
Ionic strength (M)
Data
Prediction
A
B
Fig. 3 EGTA apparent Ca2þ aYnity (K0Ca) is influenced by temperature (A) and ionic strength (B).
The experimental data in A is from Harrison and Bers (1987) at pH 7.00 and 0.19 M ionic strength and
in B from Harafuji and Ogawa (1980) at pH 6.8 and 22 �C. Predicted values are based on the
temperature and ionic strength corrections described in the text.
6 Donald M. Bers et al.
EGTA aYnity is only reduced by about 30%. In contrast, raising temperature from
1 to 36 �C approximately doubles the apparent aYnity of all three of the Ca2þ
buVers shown in Figs. 3–5 (i.e., EGTA, BAPTA, and Br2-BAPTA).
III. Methods
A. Basic Mathematical Relationships
From the forgoing and the data shown in Figs. 1 and 3–5, it is clear that one
needs to know quantitatively how the buVers being used are altered by the typical
range of experimental conditions (e.g., pH, temperature, and ionic strength). While
1 8 15 22 29 362
2.5
3
3.5
4
4.5
5
Temperature (�C)
ΔH = 0
Data
Prediction
A
B
0.1 0.15 0.2 0.25 0.30
1
2
3
4
5
6
7
Ionic strength (M)
Data
Prediction
K� C
a (in
106
M−1
)K
� Ca
(in 1
06 M
−1)
Fig. 4 BAPTA apparent Ca2þ aYnity (K0Ca) is influenced by temperature (A) and ionic strength (B).
The experimental data is fromHarrison and Bers (1987) at pH 7.00 and 0.19 M ionic strength (A) and at
pH 7.00 and 22 �C (B). Predicted values are based on the temperature and ionic strength corrections
described in the text.
1. A Practical Guide to the Preparation of Ca2þ BuVers 7
we do not want to belabor the equations, it may be useful for some readers if we lay
out some of the basics. If you are not interested in the equations, you can ignore
this section and the next (and still use the programs as more of a black box). We
hope we have accounted for things as well as possible.
In the sections above, we used Kd to talk about Ca2þ aYnity. That Kd was the
apparent overall dissociation constant, which we will get back to below (see
Eq. (5)). It is more traditional to set out the mathematical expressions starting
with the simple definition of the Ca2þ association constant KCa
KCa ¼ ½CaR�Ca�½R�½ ð1Þ
1 8 15 22 29 362
3
4
5
6
Temperature (�C)
A
B
Data
Prediction
0.1 0.15 0.2 0.25 0.30
1
2
3
4
5
6
7
8
Ionic strength (M)
Data
Prediction
K� C
a (in
105
M−1
)K
� Ca
(in 1
05 M
−1)
Fig. 5 The eVect of temperature (A) and ionic strength (B) on the apparent Ca2þ aYnity (K0Ca) of Br2-
BAPTA. The experimental data is from Harrison and Bers (1987) at pH 7.00 and 0.19 M ionic strength
(A) and at pH 7.00 and 22 �C (B). Predicted values are based on the temperature and ionic strength
corrections described in the text.
8 Donald M. Bers et al.
where R is the Ca2þ buVer. This expression is not too useful directly, because we do
not know any of the variables on the right side. It is generally more useful to have
[Ca2þ] or bound Ca2þ ([CaR]) in terms of known quantities, like total Ca2þ ([Cat])
or total chelator ([Rt]). One of the complicating factors is also that Ca2þ buVerslike EGTA or BAPTA exist in multiple unbound forms in diVerent states of
protonation. Then for a tetravalent Ca2þ buVer like EGTA, the total of the non-
Ca2þ bound forms of the buVer is
Rt½ � � CaR� ¼ ½R� þ ½HR� þ ½H2R� þ ½H3R� þ ½H4R�� ð2Þ
1. A Practical Guide to the Preparation of Ca2þ BuVers 9
where we have omitted valency for simplicity. R (or R4�) is the form which binds
Ca2þ most avidly and it is convenient to transform Eq. (1) to one with an apparent
aYnity constant for Ca2þ (K0Ca) for a given pH
K0Ca ¼
½CaR�½Ca�½R� �
½R�½Rt� � ½CaR� ð3Þ
or using Eqs. (1) and (2)
K0Ca ¼ KCa
R½ �R½ � þ HR½ � þ H2R½ � þ H3R½ � þ H4R½ � ð4Þ
Then it is a simple matter to show that
K0Ca ¼
KCa
1þ Hþ½ � KH1½ � þ Hþ½ �2KH1KH2 þ Hþ½ �3KH1KH2KH3 þ Hþ½ �4KH1KH2KH3KH4
ð5Þwhere KH1–KH4 are the four acid association constants for the buVer. Now if we
know KCa, the pH, and KH1–KH4, we can calculate K0Ca. This K0
Ca is thus the
apparent aYnity for a given [H] where pH¼� log10 ([H]/gH), and gH is the activity
coeYcient for protons under the experimental conditions (see below). This K0Ca is
the reciprocal of the dissociation constant, Kd discussed in the previous section.
Eq. (3) can also be manipulated to yield
CaR½ �= Ca½ � ¼ K0Ca Rt½ � �K
0Ca CaR½ � ð6Þ
which is the linearization for Scatchard plots of Bound/Free ([CaR]/[Ca]) versus
Bound ([CaR], where slope¼–K0Ca and x-intercept¼ [Rt]). One can also solve for
[CaR] obtaining the familiar Michaelis–Menten form.
½CaR� ¼ ½Rt�1þ 1= K
0Ca Ca�Þ½� ð7Þ
Solving for free [Ca] is more complicated because we do not know [CaR] a priori,
but substituting [CaR]¼ [Cat]� [Ca] we can get a quadratic solution
Ca½ �2 þ ð½Rt� � ½Cat� þ 1=K0CaÞ½Ca� � Cat½ �=K 0
Ca ¼ 0 ð8ÞSimilar equations can be developed for Ca2þ binding to the protonated form (e.g.,
H-EGTA) which also binds Ca2þ with a lower aYnity (e.g., see Harrison and Bers,
1987). For example, when we include Ca2þ binding to the singly protonated form
of EGTA (or HR3�) the following term must be added to the apparent aYnity
expression on the right-hand side of Eq. (5)
KCa2
1=ð Hþ½ �KH1Þ þ 1þ Hþ½ �KH2 þ Hþ½ �2KH2KH3 þ ½Hþ�3KH2KH3KH4
ð9Þ
10 Donald M. Bers et al.
where KCa2 is the Ca2þ association constant for the chelator in the singly proto-
nated form, HR. This provides some basics of the relationships for a single
chelator. However, more complicated solutions have multiple equilibria (e.g.,
other cations that bind EGTA and other Ca binding moieties) which cannot be
readily solved simultaneously in an analytical manner.
It should, however, be noted that it is simpler to go from free [Ca2þ] to [Cat],
especially with no Ca2þ competitors. This is because all of the chelators which
might bind Ca2þ will be in equilibrium with the same free [Ca2þ]. Thus, one couldsimply use a series of equations like Eq. (7) for diVerent chelators if you know the
values on the right-hand side. Then you can simply add up free [Ca2þ] plus the[CaR] values from the chelators to obtain the [Cat]. If free [Ca2þ] is not known(or chosen) it requires multiple versions of equations like Eq. (8) to be solved
simultaneously. Thus, iterative computer programs are useful (see below).
B. Temperature, Ionic Strength, and pH Corrections
While the above explains the theoretical basis for calculating the pH eVect onK0
Ca, we should clarify how we normally correct for temperature, ionic strength,
and pH for the experimental conditions used. Again, those not interested in the
details can skip this section. Thus, the final apparent aYnity (or K0Ca) should
include correction for temperature and ionic strength as well as pH. Indeed, both
proton aYnity (KH1–KH4) and metal aYnity constants (e.g., KCa) should be ad-
justed for the experimental temperature and ionic strength before adjusting for pH
as above.
1. Temperature Corrections
The standard way to correct equilibrium constants for changes in temperature
depends on knowledge of the enthalpy (DH) of the reaction.
log10K0 ¼ log10K þ DH 1=T � 1=T
0� �
= 2:303� Rð Þ ð10Þ
where temperature, T is in �K, DH is in kcal/mol and R is 1.9872�10�3 kcal/
(mol�K). Unfortunately, the DH values are not known for all the constants we
might like. For example, for EGTA they are known for the first two acid associa-
tion constants (KH1 and KH2) and the higher aYnity Ca2þ constant (KCa1). This is
generally suYcient for calculations with EGTA (see Fig. 3A). However, no DHvalues have been reported for individual constants for BAPTA and Br2-BAPTA.
Harrison and Bers (1987) measured the temperature dependence of the apparent
K0Ca for BAPTA and Br2-BAPTA. We have fit that data, varying the value of the
DH for KCa. This is somewhat empirical because there is likely to also be tempera-
ture dependence of KH1–KH4. However, the data was well described using DHvalues (for KCa) of 4.7 and 5.53 kcal/mol for BAPTA and Br2-BAPTA, respectively
(see Figs. 4 and 5). Also, since BAPTA and Br2-BAPTA are almost completely
1. A Practical Guide to the Preparation of Ca2þ BuVers 11
unprotonated already at neutral pH (see Fig. 1), the adjustments toKH1 and KH2 are
less important than with EGTA.
However, it should be noted that one cannot simply use the DH values reported
by Harrison and Bers (1987) for the overall K0Ca of BAPTA and Br2-BAPTA (3.32
and 4.04 kcal/mol) as suggested byMarks andMaxfield (1991). That is because the
intrinsic eVect of increasing temperature on the K0Ca (with DH¼0) is to reduce the
K0Ca (due to the intrinsic temperature dependence of the ionic strength adjustment,
see Fig. 4A and below). Consequently, the apparent overall DH for K0Ca (3.32 for
BAPTA) is smaller than the actual DH for KCa required (our estimate is 4.7 kcal/
mol).
Additionally, Harrison and Bers (1987) found the K0Ca for Br2-BAPTA to be
somewhat higher than the value predicted by the initial values reported by Tsien
(1980). We find that using a slightly higher KCa (log KCa¼6.96 rather than 6.8)
allowed a considerably better fit to the array of experimental data shown in Fig. 5.
2. Ionic Strength Corrections
Ionic strength can also dramatically alter the K0Ca (see Figs. 3–5). We use the
procedure described by Smith and Miller (1985) with ionic equivalents (Ie) rather
than formal ionic strength (Ie¼0.5SCi|zi|, where Ci and zi are the concentration
and valence of the ith ion). We will use the terms equivalently here. Then the
expression used to adjust for ionic strength is
log10K0 ¼ log10K þ 2xyðlog10fj � log10f
0jÞ ð11Þ
where K0 is the constant after conversion, K is the constant before, x and y are the
valences of cation and anion involved in the reaction. The terms log10fj and log10fj0are adjustment terms related to the activity coeYcients for zero ionic strength and
desired ionic strength, respectively. To adjust for ionic strength:
log10fi ¼AIe
1=2
1þ Ie1=2
� bIe ð12Þ
where b is a constant (0.25). A is a constant which depends on temperature and the
dielectric constant of the medium (e)
A ¼ 1:8246� 106
eTð Þ3=2ð13Þ
whereT is the absolute temperature (�K) and e is the dielectric constant for water. Thedielectric constant is temperature dependent and can be found from tables, but the
following equation provides an excellent empirical description over the range 0–50 �C.
e ¼ 87:7251þ 0:3974762T þ 0:0008253T 2 ð14Þ
12 Donald M. Bers et al.
where T is in �C. Thus, there is some intrinsic temperature dependence in the ionic
strength adjustment itself (see Fig. 4A, broken line). These corrections provide a
reasonably good description of the influence of ionic strength on theK0Ca inFigs. 3–5.
a. Activity CoeYcient for ProtonsThe association constants as usually reported (e.g., in Martell and Smith, 1974,
1977) are often called stoichiometric (or concentration) constants. These terms are
sensible because they imply (correctly) that they are to be used with concentrations
or stoichiometric amounts in chemical equilibria (e.g., as in Eq. (1)). While we
routinely talk about ion concentrations in ‘‘concentration’’ or ‘‘stoichiometric’’
terms, the usual exception is pH (where pH¼� log Hydrogen ion activity or
10�pH¼aH¼gH[Hþ]).
Thus, one can simply convert pH to [Hþ] and go ahead using the ‘‘stoichiomet-
ric’’ constants at face value. That is, then everything is in concentration terms and
not activity. This is the way we have done it in our programs.
The alternative is to change the stoichiometric constants to ‘‘mixed’’ constants
(for proton interactions, or KH1–KH4 only). Then you can still use pH (or 10�pH
rather than 10�pH/gH) in your calculations. Thus, acid association constants (KH1–
KH4) should be divided by the value of gH. Then you can multiply the constant by
the proton activity (since they are always of the same order in the equations (see
Eq. (5))). That is to say that [Hþ]KH1¼ ([Hþ]gH)(KH1/gH), where [Hþ]gH¼10�pH.
This method seems a bit more awkward, but the result is the same.
The proton activity coeYcient, gH varies with both temperature and ionic
strength. The empirical relationship we devised to describe this relationship is the
following
gH ¼ 0:145045� exp �B� Ieð Þ þ 0:063546� exp �43:97704� Ieð Þ þ 0:695634
ð15Þwhere B¼0.522932�exp(0.0327016�T)þ4.015942 and Ie is ionic strength and T
is temperature (in �C). This gives very good estimates of gH from 0 to 40 �C and
from 0 to 0.5 M ionic strength. This expression was sent to Alex Fabiato for use in
his computer program (Fabiato, 1991). While there is a typographical error in text
(the first coeYcient was erroneously 1.45045), the correct expression is in the
program as it was distributed.
IV. Materials
A. [Ca2þ] Measurement and Calibration Solutions
1. Measuring [Ca2þ]
While we can calculate the free [Ca2þ] or [Cat] for our solutions with the
computer programs to be described below, there are still many potential
sources of error (e.g., contaminant Ca2þ, systematic errors in pH, impurities in
1. A Practical Guide to the Preparation of Ca2þ BuVers 13
chemicals, etc.). Thus, it is valuable to measure the free [Ca2þ] to check that the
solutions are as you expected (especially for complex solutions). Ca2þ sensitive
electrodes are a convenient way to do this (see Chapter 3). We normally use
Ca2þ minielectrodes (as described in Chapter 3) or commercial macroelectrodes.
Both can be connected to a standard pH meter, but it is best to have a meter
which can read in increments of 0.1 mV. We have had good luck with Orion brand
Ca2þ-electrodes and they can be stable for 6 months or so. However, they are
rarely as good as the home-made minielectrodes. These minielectrodes are very
easy to make and are sensitive to changes in free [Ca2þ] down to 1 nM or beyond.
They do not last as long as commercial macroelectrodes, but they are extremely
cheap to make (per electrode) and can be discarded if they get contaminated with
protein or are exposed to radioactive molecules. One can also use fluorescent
indicators, once suitably calibrated, in an analogous way. The only disadvantage
there is the more limited dynamic range of these Ca2þ indicators (10-fold
above and below the Kd) versus electrodes which can give linear responses over
the 10 nM–1 M range.
2. Spreadsheet for Calibration Calculations
Making up calibration solutions for Ca2þ-electrodes (or fluorescent indicators)is really a simpler version of the multiple equilibria problem which will be discussed
below (with respect to MaxChelator), because we really only need to consider the
Ca2þ-EGTA buVer system. This approach is based on the paper by Bers (1982).
This method has the following general steps:
1. Calculate how much total Ca2þ (or free [Ca2þ]) is required for the desired
solutions (using known constants, corrected as above). All solutions should have
the same dominant ionic constituents as the solutions to be measured (e.g.,
140 mM KCl, 10 mM HEPES).
2. Measure the free [Ca2þ] with a good quality Ca2þ electrode compared to free
[Ca2þ] standards without EGTA (at higher [Ca2þ] where [Ca2þ] is more easily
controlled).
3. Accepting (for the moment) that the values from the electrode are all correct,
allows the calculation of bound Ca2þ ([CaR]) from free [Ca2þ] and total [Ca2þ].4. Scatchard plot analysis allows the independent measurement of the apparent
K0Ca and total [EGTA] in your solutions and experimental conditions (even with
systematic errors). Note that the Scatchard plot is very sensitive and deviates from
linearity at very low [Ca2þ] where Ca2þ-electrodes can become sub-Nernstian in
response (see Figs. 6 and 7).
5. Using these ‘‘updated’’ values of total [EGTA] and K0Ca you can recalculate
the free [Ca2þ] in the solutions. Then you can either use the free [Ca2þ] predictedfrom the electrode directly or you can recalculate from the total [Ca2þ] and
Ca calibration For entry of pCaSolution conditions K �Ca calculation (see A32..G47) Regression analysis (see I29-K35)
7.2 pH B= 5.12535712 Intermed 4.954 [EGTA]tot (mM)0.15 M ionic equiv (0.5*sum |zi|Ci) Gamma H= 0.76295887 H activity coefficient 0.9980 r^2
23 � C Log [H] = −7.0825011 Range for linear regression for scatchard5 mM EGTA [H] (M) = 8.27E − 08 should be linear electrode/scatchard slope
500 ml bottle K �Ca
Assn= 6.363E + 06 Log K �Ca
= 6.80365 6.86839 = log K �Ca
from scatchardK
d= 1.57E-07 M or 0.1572mM K �
Ca Discn M= 1.572E − 07 1.35E−07 = K �
Ca dissociation from scatchard
" (nM)= 157.2 135.4 nMInitial Ca-free Ca-total ml 100 mM V-Ca Ca-free Ca-free Ca-bound B/F Regresn InterpCa (nM) (mM) CaCl
2 (mV) (M) (nM) (mM) line B/F mediate (nM) pCa1 8.5000 3.162 0.099 0.493 −152 5.38E − 09 5.383 0.099 18319.405 35857.379 −4.86E − 03 2.75 8.5612 8.0000 10 0.299 1.496 −142.6 1.14E − 08 11.355 0.299 26341.654 34376.672 −4.65E − 03 8.70 8.0603 7.5000 31.623 0.838 4.188 −131.5 2.74E − 08 27.411 0.838 30553.384 30400.143 −4.12E − 03 27.55 7.5604 7.0000 100 1.944 9.722 −117.1 8.60E − 08 85.997 1.944 22608.359 22226.140 −3.01E − 03 87.48 7.0585 6.5000 316.228 3.340 16.701 −101.4 2.99E − 07 299.131 3.340 11165.474 11917.990 −1.61E − 03 280.25 6.5526 6.0000 1000 4.322 21.609 −85 1.10E − 06 1099.966 4.321 3928.010 4674.610 −6.32E − 04 924.58 6.0347 5.5000 3162.278 4.766 23.831 −70.4 3.51E − 06 3506.127 4.763 1358.415 1409.423 −1.87E − 04 3381.53 5.4718 5.0000 10000 4.932 24.662 −51.9 1.52E − 05 15232.059 4.917 322.822 268.545 −2.13E − 05 17,312.44 4.7629 4.5000 31622.777 5.007 25.034 −38.1 4.56E − 05 45563.879 4.961 108.884 −55.893 5.30E − 05 63,647.15 4.196
10 3.0000 1,000,000 5.999 29.995 0 9.38E − 04 938455.736 5.061 5.392 −790.218 1.05E − 03 1,046,090.19 2.98011 3.0000 1,000,000 5.999 29.995 0 9.38E − 04 938455.736 5.061 5.392 −790.218 1.05E − 03 1,046,090.19 2.98012 3.0000 1,000,000 5.999 29.995 0 9.38E − 04 938455.736 5.061 5.392 −790.218 1.05E − 03 1,046,090.19 2.980
2.0 10 mM 10 mM 28.2 From regression3.0 1mM 1mM 0.8 [EGTA]tot K-Ca-EGTA4.0 100mM 100mM −29.6 4.953608052 mM 7.38567143 × 10^6/M
Avg slope= 28.9 99.07% % pure 6.868389983 = log KSlope (mV)= 29 Regression I/J14-: 21 Regression I/J13-: 21
mV offset at 1 mM Ca= 0.8 Slope B/F intercept Slope B/F intercept-7385.67143 36585.72 −6516.46 32752.75
Temperature and ionic strength correction SE of coeff 135.41 548.75 435.32 1663.88Std cond Ionic str Final Final R^2 0.9980 571.40 0.9697 2324.14
Temp 20 incl T eff 23� C F stat 2,975 6 224 7I-Eq 0.100 0.150 0.150 M Delta H Valence Reg sum Sq 9.71E + 08 1.96E + 06 1.21E + 09 3.78E + 07Stoich Log K Log K � Log K � K � (M) kcal/mol 2*x*yconstK1 9.47 9.3576 9.3138 2.060E + 09 −5.8 8K2 8.85 8.7657 8.7219 5.271E + 08 −5.8 6K3 2.66 2.6038 2.6038 4.016E + 02 0 4K4 2 1.9719 1.9719 9.374E + 01 0 2KCa 10.97 10.7453 10.6840 4.831E + 10 −8.1 16KCa2 5.3 5.1315 5.1315 1.353E + 05 0 12Log f 0.109225 0.12327007Temp 293 296A 0.507424 0.51006648Epsil 80.1057 79.0197311
Recalculated
−200
−160
−120
−80
−40
0
40
23456789
Ele
ctro
de r
esp
(mV
)
pCa
Electrode calibration
Initial pCa
Recalculated
No EGTA
−1.E + 04
0.E + 00
1.E + 04
2.E + 04
3.E + 04
4.E + 04
0.0 2.0 4.0 6.0
Bou
nd/fr
ee
Bound (mM)
Scatchard plot
Data
Regression
Fig. 6 Excel spreadsheet used to prepare Ca2þ calibration buVers using a Ca2þ electrode. This version is used when you want to start
with the pCa of the calibration solutions as input and determine howmuch total Ca2þ is needed to achieve the desired free [Ca2þ]. It alsoallows updating of the apparentK0
Ca and free [Ca2þ] in the calibration solutions. This and related spreadsheets can be freely downloaded
(see text for details).
0 1 2 3 4 50
40,000
30,000
20,000
10,000
Regression line
Bou
nd/F
ree
(mM
/mM
)
Bound Ca2+-EGTA (mM)
3 4 5 6 7 8 9
−150
−100
−50
0
pCa
Ele
ctro
de r
espo
nse
(mV
)
Original pCaElectrode pCaRecalc. pCa
A B
Fig. 7 Scatchard plot (A) and electrode calibration curves (B) for the spreadsheet shown in Fig. 6. The
Scatchard plot allows estimation of the total [EGTA] (x-intercept) and the apparent association
constant, K0Ca (-slope). The Scatchard plot is very sensitive to the detection limit of the Ca2þ electrode.
The leftmost two points in A are the lowest free [Ca2þ] in the calibration curve in B (and are not included
in the regression). The three calibration curves shown are for the original (or planned pCa), the pCa
predicted solely by the electrode and the pCa after recalculation, using the values determined in the
Scatchard plot along with the total Ca2þ added to the buVers. In this instance, there was good agreement
between the three curves, but this is not always the case (see Bers, 1982).
1. A Practical Guide to the Preparation of Ca2þ BuVers 15
updated constants. The latter is necessary for the lowest free [Ca2þ] where the
electrode response is becoming nonlinear (�pCa 9).
We use a spreadsheet (Excel) to greatly simplify all of these steps (see Fig. 6).
There are three basic versions of this spreadsheet: one for starting with free
[Ca2þ] as the input (DMB-CAF-2010.xls), one for pCa as input (DMB-PCA-
2010.xls), and one for total Ca2þ as input (DMB-CAT-2010.xls). These can be
freely downloaded from the MaxChelator site as described below. We will walk
you through the use of this spreadsheet in making a series of free [Ca2þ]standards here.
The fields for the input of data are shaded dark gray. For the pCa version of the
spreadsheet in Fig. 6 you proceed as follows (the others versions are completely
analogous):
1. Enter your solution conditions (upper left, pH, ionic strength, temperature,
total [EGTA], and bottle size you will use). The K0Ca values are then automatically
adjusted for the selected temperature, pH, and ionic strength (lower left box).
2. Enter the desired pCa values. The free [Ca2þ], total [Ca2þ], and ml of 100 mM
Ca2þ stock are automatically calculated (next three columns) using the adjusted
K0Ca.
16 Donald M. Bers et al.
3. Enter the mV readings from a Ca2þ electrode (including values for Ca2þ
standards lacking EGTA at 100 mM, 1 mM, and 10 mM [Ca2þ] and the electrode
reading at 1 mM free [Ca2þ] as the ‘‘oVset’’). This is the fourth column (V-Ca) and
you can choose the electrode slope (rather than assume the average). The free
[Ca2þ], Ca2þ-bound to EGTA, and the bound/free (B/F) are then automatically
calculated (based on the electrode response and total [Ca2þ]).4. Those calculated values (light shaded box, yellow in downloaded file) will be
subject to linear regression Scatchard analysis (automatically). The Scatchard plot
and Electrode calibration curves (Fig. 6, bottom) can be inspected to check
linearity. If values within the regression window are not on the linear range, they
can throw oV the analysis. The top and bottom two [Ca2þ] are excluded from the
regression to allow calculations of [Ca2þ] for solutions outside that range.5. Finally, the free [Ca2þ] and pCa are automatically recalculated using the
measured K0Ca and total [EGTA] (from the auto-analysis) as well as the total
Ca2þ values (last two columns). The Electrode calibration curve and Scatchard
plot allow you to get an overview of the results. (Fig. 6).
We routinely use this for calibration solutions for both Ca2þ-electrodes and
fluorescent indicators. In addition to improving the reliability of Ca2þ calibration
solutions, one of the convenient aspects of this spreadsheet is that you can see all
the details of what is going on. For example, you can see that the EGTA is almost
completely saturated as you get up to 10 mM free [Ca2þ]. In this range we usually
believe the electrode, rather than our ability to pipette within 1% of the required
volume. On the other hand, as you approach the detection limit of the electrode
(e.g., �pCa 9), we use the recalculated pCa values. The measured versus predicted
K0Ca, EGTA purity and [Ca2þ] can also be useful in identifying potential systematic
errors or changes in your procedures.
B. Preparing BuVer Solution
1. Basic Steps in Solution Preparation
There are no hard and fast rules or special tricks to make these buVers, butspecial care in weighing and pipetting, and common sense can help avoid some
potential problems. The water should be well purified to minimize contamination
with Ca2þ and other metals. We usually use water that is first distilled and then run
through a water purification system containing at least one ion exchange column
(e.g., Nanopure, from Barnstead). This provides water with resistivity of
>15 MOhm-cm. Starting with good water like this is important for removal of
other metal contaminants as well as Ca2þ. There can also be contaminating Ca2þ
and metals in the salts and chemicals used to make solutions. In the end, it is
typical to find 1–3 mM free [Ca2þ] in nominally Ca2þ-free solutions. This can be
checked with a Ca2þ-electrode.
1. A Practical Guide to the Preparation of Ca2þ BuVers 17
Some people include 1–2 mM TPEN, a heavy metal chelator in Ca2þ-buVersolutions. This can chelate submicromolar amounts of heavy metals, which may
or may not be chelated by the dominant Ca2þ buVer. This may not be important in
routine applications, but may ensure that the Ca2þ-sensitive process under study
will not be altered by trace amounts of other metals. All solutions should be made
and stored in clean plastic ware (careful washing and extensive rinsing in deionized
water is required). Glass containers should be avoided. EGTA can leach Ca2þ out
of glass leading to gradual increase in free [Ca2þ] in the solutions. We have often
been able to store Ca2þ calibration solutions for more than 6 months in polypro-
pylene bottles (provided that there is no organic substrate to foster bacterial
growth).
An accurate [Ca2þ] standard is important for making Ca2þ buVers. It is diYcult
to make accurate [Ca2þ] using CaCl2 � 2H2O typically used to make physiological
solutions. This is because the hydration state varies making stoichiometric weigh-
ing imprecise. CaCO3 can be more accurately weighed, but has the disadvantage
that you must then drive oV the CO2 with prolonged heating and HCl, unless
HCO3 is desired in the solutions (which is a weak Ca2þ buVer itself). A convenient
alternative is to buy a CaCl2 standard solution and we use a 100 mM CaCl2solution from Orion (BDH also sells an excellent 1 M CaCl2 standard). To save
money, one can titrate a larger volume of CaCl2 to the same free [Ca2þ] as the
Orion standard using a Ca2þ-electrode.It is also important to prepare accurate stock solutions of Ca2þ chelators. EGTA
from diVerent commercial sources diVer somewhat in purity (Bers, 1982; Miller
and Smith, 1984), but manufacturers provide purity estimates that help (we find
that purity typically ranges from 95 to 100% of the stated purity). BAPTA has also
been reported to contain 20% water by weight (Harrison and Bers, 1987), but can
be dried at 150 �C until the weight is constant to assure removal of water. If one
measures the total buVer concentration (as described in section above) this prob-
lem can be largely obviated. We typically measure the purity of each lot of EGTA
or BAPTA that we use, taking this approach. Then we often keep track on the
bottle itself, so that we can confirm the value upon subsequent tests with the same
batch. EGTA (in the free acid form) is also not very soluble because of the acid pH.
For neutral pH solutions, it is practical to dissolve EGTA with KOH in a 1:2
stoichiometry, since at neutral pH two of the four protons on EGTA are disso-
ciated (vs. all four for BAPTA).
When Ca2þ is added to EGTA solutions, 2 mol of Hþ are released for each mole
of Ca2þ bound. Thus, the pH should always be adjusted as the Ca2þ is being added
or afterward. The strong pH dependence of the K0Ca of EGTA (Fig. 1) emphasizes
the importance of this point. We typically measure [Ca2þ] and pH simultaneously
just before the solutions are brought up to final volume (for approximate pH
adjustment) and after, for final pH adjustment (as close to the third decimal
place as possible) and [Ca2þ] measurement. The solutions are also checked again
later to assure consistency. The rigorous attention to pH adjustment will obviously
be less crucial for the BAPTA buVers.
18 Donald M. Bers et al.
It may well be asked, why not just use BAPTA rather than EGTA? The main
reason is expense, BAPTA is about 30 times more expensive. The other reason is
that EGTA is the ‘‘Devil we know’’ and indeed we do know much about its
chemistry (e.g., metal binding constants, DH values). For applications with small
volumes of solution though, it may be quite reasonable to replace EGTA with
BAPTA.
The ionic strength contribution of the pH buVer should also be included in the
ionic strength calculation (Ie¼0.5SCi|zi|). This requires calculation of the fraction
of buVer in ionized form (i.e., not protonated).
2. Potential Complications
Not all of the desired constants have been determined for the metals and
chelators of interest. This places some limitations on how accurately one can
predict the free [Ca2þ] of a given complex solution or determine how much total
Ca2þ is required to achieve a desired free [Ca2þ]. The same is true for other species
of interest (e.g., Mg2þ, Mg2þ-ATP). Some Ca2þ buVers also can interact with Ca2þ
in multiple stoichiometries (e.g., the low aYnity Ca2þ buVer, NTA (nitrilotriacetic
acid) can form Ca2þ-NTA2 complexes). There can also be systematic errors in pH
measurements (Illingworth, 1981) or purity of reagents. Purity can be estimated as
described above.
The pH problem is actually quite common, especially with combination pH
electrodes. To put it simply, the reference junction of some electrodes (particularly
with ceramic junctions) can develop junction potentials which are sensitive to ionic
strength. This problem can be exacerbated when the ionic strength of the experi-
mental solutions diVers greatly from the pH standards (typically low ionic strength
phosphate pH standard buVers). A systematic error in solution pH of about 0.2 pH
units is not at all uncommon. As is clear from Fig. 1, this could translate into a 0.4
error in log K0Ca and produce a two- to threefold diVerence in free [Ca2þ] even
where EGTA is at its best in terms of buVer capacity.While measuring the free [Ca2þ] with an electrode can be extremely valuable, it is
not foolproof either. Ca2þ electrodes are not perfectly selective for Ca2þ (see
Chapter 3). For example, the selectivity of these electrodes for Ca2þ over Mg2þ
is about 30,000–100,000 (Schefer et al., 1986). This roughly corresponds to the
diVerence in intracellular concentrations. Thus, a 100 nM Ca2þ solution with
1 mM Mg2þ would look to the electrode like a 110–130 nM Ca2þ solution. For
the Ca2þ electrodes described in Chapter 3 (using the ETH 129 chelator), the
interference by Na or K is less. For 140 mM Na or K in a 100 nM Ca2þ solution,
the apparent [Ca2þ] would be only about 101 nM.
Some Ca2þ buVers can also interfere with Ca2þ electrodes. Citrate, DPA (dipi-
colinic acid), and ADA (acetamidoiminodiacetic acid), three low aYnity Ca2þ
buVers were found to interfere with Ca2þ electrode measurements, while NTA
did not (Bers et al., 1991). Interestingly, citrate, DPA, and ADA (which modified
electrode behavior) also modified Ca2þ channel characteristics, but NTA did not.
1. A Practical Guide to the Preparation of Ca2þ BuVers 19
When Ca2þ electrodes cannot be practically used, one may still be able to use
optical indicators such as the fluorescent indicators fura-2, indo-1, Fluo-4, Fluo-
5N for selected [Ca2þ] ranges, or the metallochromic dyes antipyralazo III, mur-
exide, or tetramethylmurexide for higher free [Ca2þ] (Kd�200 mM, 3.6, and
2.8 mM, respectively, Ohnishi, 1978, 1979; Scarpa et al., 1978). Of course, these
indicators require calibration too.
A general potential complication with Ca2þ buVers is that they may alter the very
processes one is interested in studying with Ca2þ buVers. For example, EGTA and
other Ca2þ-chelators have been documented to increase the Ca2þ sensitivity of the
plasmalemmal and SR Ca2þ-ATPase pumps and also of Naþ/Ca2þ exchange
(Berman, 1982; Sarkadi et al., 1979; Schatzmann, 1973; Trosper and Philipson,
1984). For example, 48 mMEGTAdecreased the apparentKCa ofNaþ/Ca2þ exchange
in cardiac sarcolemmal vesicles from 20 to 5 mMCa2þ (Trosper and Philipson, 1984).
These points above are not meant to discourage one from using Ca2þ buVers,but simply to point out some of the potential problems that one might encounter.
Being aware of what might occur can help troubleshoot, when things do not make
sense. Clearly, the use of Ca2þ buVer solutions is essential for the understanding ofCa2þ-dependent phenomena. Our aim here is to provide helpful information.
C. Software Programs
While the above spreadsheet is useful for very simple Ca-EGTA or Ca-BAPTA
solutions used for calibrations, it is not suYcient for more complex buVers that onetypically uses experimentally (which include Mg2þ in addition to Ca2þ and multiple
anionic species like ATP that bind Ca2þ and Mg2þ). Several computer programs
have been described (Bers et al., 1994; Brooks and Storey, 1992; Fabiato, 1988;
McGuigan et al., 1991; Schoenmakers et al.., 1992; Taylor et al., 1992), but we will
focus on, MaxChelator developed by one of the authors (CWP Bers et al., 1994). We
have seen above that care is needed in using Ca2þ electrodes. This is equally true for
any software used to determine free metal concentrations in the presence of chela-
tors. In both cases, careful measurement of environmental conditions is needed:
temperature, pH, and ionic strength, as well as attention to the quality and accuracy
of measurement of all reagents. In addition, software is aVected by the choice of
stability constants, quality of the code, and the particular algorithms used, and of
course, the understanding of those using the software (being dependent on personal
knowledge and the ease of use of the software).
Fabiato and Fabiato (1979) broke ground for average users by publishing their
paper on using a hand held programmable calculator to determine free [Ca2þ] or[Mg2þ] in the presence of EGTA. Before then complicated and user unfriendly
software running on main frames and mini computers was all that was available.
Use of Ca2þ electrodes was also just starting and not easy for most labs to
implement. The Fabiato code opened this door, but was somewhat limited.
Richard Steinhardt’s lab used the Fabiato paper to write a version for the Apple
2e, and one of us (CWP) further developed this to a program known as the
20 Donald M. Bers et al.
MaxChelator series of programs, first introduced in the 1994 version of this
chapter.
There was no internet 16 years ago when the first edition of this chapter was
presented. The compilation of useful stability and thermodynamic constants (e.
g., Martell and Smith, 1974, 1977) has not grown with the explosion of biological
use of Ca2þ buVers and novel Ca2þ indicators (although resources are available
at the National Institute of Standards and Technology (NIST) web site http://
www.nist.gov/srd/nist46.htm). For most of these new compounds, accurate sta-
bility constants have not been determined. Further, there is some disagreement
over which constants and algorithms are best. However, as implied above, it is
valuable to be able to calculate appropriate stoichiometric concentrations of, for
example, Ca2þ, Mg2þ, EGTA, and ATP to use in your solutions to obtain the
desired free [Ca2þ] and [Mg2þ] and [Mg2þ-ATP]. On the other hand, there is no
substitute for actually measuring the concentration when possible to
avoid imperfections in the calculations and also systematic errors (McGuigan
et al., 2007).
Two commonly used programs are Chelator by Theo Schoenmakers
(Schoenmakers et al., 1992) and the MaxChelator series by one of the authors
(CWP). Chelator is written for DOS and has not been updated since 1992 (making
it less broadly useful in 2010) as fewer computers and users run DOS programs and
the user interface is dated. The MaxChelator series expanded into Windows (both
16 and 32 bit), andmore recently into the web via Javascript to be more OS neutral.
This website has downloadable versions of the MaxChelator suite, Chelator, and
several other related tools (including the Bers’ Spreadsheets as in Fig. 6): http://
maxchelator.stanford.edu/downloads.htm
1. Ideal Software Criteria
1. First and foremost is the software has to give the correct answer or at least
close enough that it does not aVect the experimental conclusions (and allows
measurement verification).
2. Must be easy to use. Users should not be confused as to where to enter
information or what information to enter.
3. Adaptable. There should be an easy way to enter diVerent constants and
possibly even allow for diVerent methods of doing some of the calculations.
4. Source code available so the knowledge is not lost with the programmer/
researchers.
No current software handles all these requirements well.
1. A Practical Guide to the Preparation of Ca2þ BuVers 21
2. Accuracy
We think that the constants and algorithms for the calculations in these pro-
grams are appropriate, but the ambiguities in available fundamental constants,
some nuances in their application and the systematic experimental errors discussed
above conspire such that solution making by recipe is imperfect. Experience and
direct [Ca2þ] measurement are the best ways to limit inaccuracies in the long run.
Indeed, blind acceptance of the calculations, and a presumption that there are no
systematic errors (in either the solution making, pH, temperature, or in the
calculations) enhances the likelihood for inaccuracies.
3. Ease of Use and Adaptability
Early versions of these programs were DOS based and developed within various
labs, with user interfaces not consistent with present day expectations. Patton’s
MaxChelator has attempted to maintain a user friendly interface that has evolved
during the past 15 years. One can input either desired free concentrations of Ca2þ,Mg2þ, Mg-ATP to obtain the total concentrations required or vice versa, and there
are simple intuitive screens for these inputs. We are not aware of a commercial
program that does these calculations. To ensure adaptability in this future, code
should be available as open source to maximize access for future improvements
(including by others).
Both Chelator and MaxChelator allow for additional chelators and sets of
constants to be created or changed (a useful feature), but do not allow for their
inner workings or equations to be changed. If programs were open sourced then
the inner algorithms could be changed to try out diVerent ideas. Software could be
‘‘tweaked’’ and refined to hopefully overcome its limitations. Another issue for the
future is whether there will be suYcient interest in the continual evolution of these
software suites.
4. Other Things to be Aware of When Doing This Work
In line with the aforementioned concerns, Patton et al. (2004) mentioned several
precautions. First, pH control is critical (within 0.01 pH unit) especially for EGTA.
Moreover, when metals bind to chelators, Hþ is released aVecting pH, and that
increases the importance of appropriate pH buVer choice. Second, chelators
cannot reduce free metal concentration to zero. An equilibrium is set up, and
[Ca2þ] and [Mg2þ] (like [Hþ]) are always finite. If proteins or other moieties in your
system have higher aYnity for the metal than your chelator, it can complicate
chelator eVectiveness. Contamination with Ca2þ is almost always present. Third,
select the right chelator for metal concentrations of interest. Just like pH buVerswhich work in a range of �0.5 pH units, chelators work in a range of �0.5pKd
(�0.3–3Kd). Using too high Kd allows contaminant Ca2þ to strongly influence
[Ca2þ] at the low levels, while too low Kd will result in saturation and loss of
22 Donald M. Bers et al.
buVering near the higher end. Note that the lower aYnity buVers typically have
higher oV-rates and thus equilibrate faster and damp rapid [Ca2þ] spikes more
eVectively.
5. Why Use Software and Where to Get MaxChelator?
Despite all the caveats, using software to calculate free [Ca2þ], [Mg2þ], and [Mg-
ATP] is necessary to have a reasonable chance of getting the solutions right. And
we think that MaxChelator is a useful tool in this regard. More information and
downloads (free) are available at http://maxchelator.stanford.edu/. Whenever
practical, it is also highly desirable to measure the [Ca2þ] using either electrodes
or fluorescent Ca2þ indicators to confirm the predictions and check for reproduc-
ibility. These measurements are less practical for other metals (even Mg2þ) or
anions, for which electrodes and fluorescent indicators are less available.
6. MaxChelator for Windows
The earliest MaxChelator eVort was a DOS program which was then moved to
Windows (Winmaxc), and the latest version is posted at the above website. The
current Windows version allows visualization in two or three dimensions, some of
the key factors that aVect the result. The source code is hundreds of pages long andis complied under the Delphi (Visual Pascal) environment (not posted). The files of
constants are editable using a text editor and any number of files of constants can
be maintained. However, the algorithms used to calculate the eVects of tempera-
ture and ionic concentration are hidden, limiting the flexibility of this version. On
the other hand, it is straightforward to use and multiple metals and chelators can
be easily used together (e.g., Ca2þ, Mg2þ, Ba2þ, BAPTA, Br2BAPTA, and ATP).
7. Javascript Web Versions
Not everyone wants to use windows software, so the algorithms have been
ported to Javascript which runs on all platforms that have a browser with Java-
script enabled (with syntax similar to C programming language). One limitation is
that the math libraries for interpreted Javascript are not as accurate as those for
compiled programs (and rounding errors can create limitations, especially with
simultaneous use of multiple metals and chelators). Some people also disable
Javascript because of the fear of malware.
An advantage of Javascript, besides running on most computers, is that the
source code is readily accessible and can be saved, edited, and then run on any
machine. If the result is an improvement, it can be shared. Another advantage is
the simple user interface. Everything is in front of you all the time, and it is very
easy and intuitive to change pH, temperature, ionic concentration, or metal/
chelator concentrations. Several variants are available for either online calcula-
tions or download. Some are simple binary Ca-EGTA or Mg-ATP calculators like
1. A Practical Guide to the Preparation of Ca2þ BuVers 23
the one in the screenshot below. One simply chooses the calculation type at the top,
enter the temperature, pH, and ionic strength (line 2) and the two known
Ca-EGTA concentrations. Not only are the traditional find free Ca2þ and find
total Ca2þ calculations performed, but also the occasionally useful find total
EGTA given the free and total (or free and bound) Ca2þ levels can be performed.
There are also the slightly more complex versions for Ca–Mg-ATP-EGTA equili-
bria. Finally, there is the more comprehensive version (Web MaxC) that allows any
combination of cations (Al3þ, Ba2þ, Ca2þ, Cd2þ, Cu2þ, Fe2þ, Mg2þ, Sr2þ, Zn2þ)and 12 diVerent chelators (including EGTA, BAPTA, Br2BAPTA, EDTA, ATP,
ADP, and citrate), but the simplicity and functionality are the same as the simple Ca-
EGTA version above. There are also versions posted that use the Schoenmakers
constants and conditions and other versions will be posted as they are written. These
programs can thus be helpful in designing solutions with particular free ion con-
centrations, but should be used with understanding of the limitations.
V. Discussion and Summary
It is important to be able to prepare solutions with buVered [Ca2þ], and often
these solutions are complicated by multiple equilibria, and theoretical and practi-
cal limitations. Here we have discussed some of the basic principles that are
involved, several key factors that complicate the process and provide some practi-
cal tools and advice to increase the probability that one can make the desired
solution. However, neither the calculations nor the solution preparation nor
measurement are foolproof. One must be alert to some of the potential caveats,
and make independent measurements when possible.
Often it is useful to make a very careful set of calibration standards at a selected
ionic strength, temperature, and pH, using simpler solutions (e.g., containing
simple Ca-EGTA buVers) for standardization of either a Ca2þ electrode or fluo-
rescent Ca2þ indicator (as in Fig. 6). Note also that there are [Ca2þ] solution sets
24 Donald M. Bers et al.
sold commercially for this purpose, but they may not mimic your preferred con-
ditions (and we have not used them). Once your electrode or fluorescent indicator
is calibrated, you can use it to measure [Ca2þ] in more complex solutions, where
solution predictions are less reliable. These more complex solutions could be a
series of solutions of diVerent [Ca2þ] or [Mg-ATP], for example, to activate
skinned muscle fiber contraction, expose to permeabilized cells, dialyze into cells
via patch pipettes or use directly in biochemical assays in vitro. This is certainly a
rational and practical approach. One practical caveat is that the aYnity of most
fluorescent Ca2þ indicators changes (usually decreases two- to fourfold) in the
cellular environment versus in protein-free solutions (Harkins et al. 1993; Hove-
Madsen and Bers, 1991; Konishi et al., 1988; Uto et al., 1991) and this seems to be
due to the interaction of the indicators with cellular proteins (which can be
mimicked in vitro). So precise control and measurement of [Ca2þ]i in cells are
both very diYcult to fully achieve. On the other hand, the importance of [Ca2þ]makes it important to measure and try to control [Ca2þ] as best one can. Aware-
ness of the limitations may seem daunting, but should not dissuade one from these
valuable experiments. Even relative [Ca2þ] changes and imperfect control or
measurement of [Ca2þ] are of value in understanding these processes.
Acknowledgements
This work was supported by a grant from the National Institutes of Health (HL30077).
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CHAPTER 2
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
Photorelease Techniques for Raising orLowering Intracellular Ca2þ
Robert ZuckerMolecular and Cell Biology DepartmentUniversity of California at BerkeleyBerkeley, California, USA
OGY,All rig
A
Vhts
bstract
OL. 99 0091reserved. 27 DOI: 10.1016/S0091-
-679X679X
I.
I ntroduction II. N itr CompoundsA.
Chemical Properties B. Calculating [Ca2þ]i Changes in CellsIII.
D M-Nitrophen A. Chemical Properties B. Calculating Changes in [Ca2þ]iIV.
D iazo Compounds A. Chemical Properties B. Calculating EVects of PhotolysisV.
I ntroduction into Cells VI. L ight Sources V II. C alibration V III. P urity and Toxicity IX. B iological ApplicationsA.
Ion Channel Modulation B. Muscle Contraction C. Synaptic Function D. Other ApplicationsX.
C onclusions R eferences/10 $35.00(10)99002-X
28 Robert Zucker
Abstract
The quantitative manipulation of intracellular calcium concentration ([Ca2þ]i) is avaluable instrument in the modern cell biologists’ toolbox for unraveling the many
cell processes controlled by calcium. I summarize here the major classes of photo-
sensitive calcium chelators used to elevate or reduce [Ca2þ]i, with an emphasis on
their physicochemical properties and methods of calculating magnitudes and kinet-
ics of eVects on [Ca2þ]i of flashes and steady light, in order to encourage the choice of
the best substance for particular applications. The selection and calibration of
appropriate light sources, and procedures for introducing the chelators into cells,
spatially restricting [Ca2þ]i changes, and measuring the profiles of [Ca2þ]i changesimposed by photolysis, are also described. The final section describes a selection of
biological applications.
I. Introduction
Photolabile Ca2þ chelators, sometimes called caged Ca2þ chelators, are used to
control [Ca2þ]i in cells rapidly and quantitatively. A beam of light is aimed at cells
filled with a photosensitive substance that changes its aYnity for binding Ca2þ.Several such compounds have been invented that allow the eVective manipulation
of [Ca2þ]i in cells. These compounds oVer tremendous advantages over the alter-
native methods of microinjecting Ca2þ salts, pharmacologically releasing Ca2þ
from intracellular stores, or increasing cell membrane permeability to Ca2þ using
ionophores, detergents, electroporation, fusion with micelles, or activation of
voltage-dependent channels, in terms of specificity of action, repeatability and
reliability of eVect, maintenance of cellular integrity, definition of spatial extent,
and rapidity of eVect, all combined with the ability to maintain the [Ca2þ]i changefor suYcient time to measure its biochemical or physiological consequences. Only
photosensitive chelators allow the concentration of Ca2þ in the cytoplasm of intact
cells to be changed rapidly by a predefined amount over a selected region or over
the whole cell. Since loading can precede photolysis by a substantial amount of
time, cells can recover from the adverse eVects of the loading procedure before the
experiments begin. The ideal photosensitive Ca2þ chelator does not exist, but
would have the following properties.
1. The compound could be introduced easily into cell, by microinjection or by
loading a membrane-permeating derivative that would be altered enzymatically to
an impermeant version trapped in cells.
2. The compound could be loadedwithCa2þ to such a level that the unphotolyzed
formwould buVer the [Ca2þ]i to near the normal resting level, so its introduction into
cells would not perturb the resting Ca2þ level. Additionally, by adjusting the Ca2þ
loading or selecting chelator variants, the initial resting Ca2þ level could be set to
somewhat higher or lower than the normal resting concentration.
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 29
3. The chelator should be chemically and photolytically stable.
4. Photolysis by a bright flash of light should allow rapid changes in the free
Ca2þ level; this characteristic requires rapid photochemical and subsequent dark
reactions of the chelator.
5. Photolysis should be achievable with biologically appropriate wavelengths,
which requires a high quantum eYciency and absorbance at wavelengths that
readily penetrate cytoplasm but cause little biological damage, that is, that are
not highly ionizing. For the chelator to be protected from photolysis by light
needed to view the preparation would also be useful.
6. The photoproducts, or postphotolysis buVer mixture, should continue to
buVer Ca2þ, and so hold it at the new level in the face of homeostatic pressure
from membrane pumps and transport processes.
7. Neither the unphotolyzed chelator nor its photoproducts should be toxic, but
rather should be inert with respect to all ongoing cellular molecular and physio-
logical processes. Three classes of compounds, the nitr series, DM-nitrophen, and
the diazo series, share enough of these properties to have generated intense interest
and widespread popularity, and form the subjects of this review.
Numerous more general reviews of photolabile or caged compounds, which
contain some information on photolabile Ca2þ chelators, have appeared (Adams
and Tsien, 1993; Gurney, 1993; Kao and Adams, 1993; Kaplan and Somlyo, 1989;
McCray and Trentham, 1989; Ogden, 1988; Parker, 1992; Walker, 1991). Reviews
focused more on photosensitive Ca2þ chelators may also be consulted (Ashley
et al., 1991a; Ellis-Davies, 2003; Gurney, 1991; Kaplan, 1990).
II. Nitr Compounds
A. Chemical Properties
The first useful class of photosensitive Ca2þ chelators was developed by Roger
Tsien. This nitr class of compounds relies on the substitution of a photosensitive
nitrobenzyl group on one or both of the aromatic rings of the Ca2þ chelator 1,2-bis
(o-aminophenoxy)ethane-N,N,N0,N0-tetracetic acid (BAPTA) (Adams and Tsien,
1993; Adams et al., 1988; Kao and Adams, 1993; Tsien and Zucker, 1986). Light
absorption results in the abstraction of the benzylic hydrogen atom by the excited
nitro group and oxidation of the alcohol group to a ketone. The resulting nitro-
sobenzoyl group is strongly electron withdrawing, reducing the electron density
around the metal-coordinating nitrogens and reducing the aYnity of the tetracar-
boxylate chelator for Ca2þ. In the first member of this series, nitr-2, methanol is
formed as a by-product of photolysis, but in subsequent members (nitr-5, nitr-7,
and nitr-8) only water is produced. Photolysis of nitr-2 is also slow (200 ms time
constant). For the other nitr chelators, the dominant photolysis pathway is much
faster (nitr-7, 1.8 ms; nitr-5, 0.27 ms; and nitr-8, not reported). For these reasons,
30 Robert Zucker
nitr-2 is no longer used. For the three remaining nitr compounds, photolysis is
most eYcient at the absorbance maximum for the nitrobenzhydrol group, about
360 nm, although light between 330 and 380 nm is nearly as eVective. The quantumeYciency of the Ca2þ-bound form is about 1/25 (nitr-5, 0.035 and nitr-7, 0.042) and
is somewhat less in the Ca2þ-free form (0.012 and 0.011). The absorbance at this
wavelength is 5500 M�1 cm�1 (decadic molar extinction coeYcient) for nitr-5 and
nitr-7, and 11,000 M�1 cm�1 for nitr-8. The structures of the nitr series of com-
pounds are given in Fig. 1; and the photochemical reaction of the most popular
member of this group, nitr-5, is shown in Fig. 2. The physico-chemical properties
of these and other photosensitive chelators are summarized in Table I.
These chelators share the advantages of the parent BAPTA chelator: high
specificity for Ca2þ over Hþ and Mg2þ (Mg2þ aYnities, 5–8 mM), lack of
dependence of Ca2þ aYnity on pH near pH 7, and fast buVering kinetics. One
limitation is that the drop in aYnity in the nitr compounds after photolysis is
relatively modest, about 40-fold for nitr-5 and nitr-7. The Ca2þ aYnity of nitr-5
drops from 0.15 to 6 mM at 120-mM ionic strength after complete photolysis.
These aYnities must be reduced at higher ionic strength, roughly in proportion to
the tonicity (Tsien and Zucker, 1986). By incorporating a cis-cyclopentane ring
into the bridge between the chelating ether oxygens of BAPTA, nitr-7 was created
COO−
COO− COO− COO− COO− COO− COO−
O
O O
O O
OO
O
O
O
O
O
O
OHOH
OH OH
O2N
HO
OMe
MeHH
H H H
N N
NO2NO2
NO2 NO2
nitr-5 nitr-7
nitr-9nitr-8
O O O
N
N
N
COO− COO−COO− COO−
N
COO−
Me
N
COO−COO−
Fig. 1 Structures of the nitr series of photolabile chelators, which release calcium on exposure to light.
NO2 NO+
Photolyzed nitr-5(low Ca2+ affinity)
H2O
CH3
N N
CH3H
nitr-5(high Ca2+ affinity)
HO
OO O
O
O
O OO
Ca2+
Ca2+
−O2C
−O2C −O2C
−O2C
O
N
CO2− CO2
−
CO2− CO2
−
N
hn
Fig. 2 Reaction scheme for the photorelease of nitr-5.
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 31
with significantly higher Ca2þ aYnities (54 nM, decreasing to 3 mM after photol-
ysis at 120 mM ionic strength). To increase the change in Ca2þ-binding aYnity on
photolysis, nitr-8 was created with a 2-nitrobenzyl group on each aromatic ring of
BAPTA. Photolysis of each group reduces aYnity only about 40-fold, as for nitr-
5 and nitr-7, but photolysis of both nitrobenzyl groups reduces aYnity nearly
3000-fold, to 1.37 mM, with a quantum eYciency of 0.026. Finally, nitr-9 is a
dicarboxylate 2-nitrobenzhydrol with a low Ca2þ aYnity that is unaVected by
photolysis; this compound can be used to control for nonspecific eVects of the
photoproducts.
Initially, nitr-5 was the substance most often applied in biological experiments,
largely because it was the first photolabile chelator to have most of the qualities of
the ideal substance. The limited aYnity for Ca2þ of this substance in the unpho-
tolyzed form requires that it be lightly loaded with Ca2þ when introduced into
cells; otherwise, the resting [Ca2þ]i will be too high. However, the compound in a
lightly loaded state contains little Ca2þ to be released on photolysis. Nitr-7 alle-
viates this problem with an aYnity closer to that of normal resting [Ca2þ]i, but itssynthesis is more diYcult and its photochemical kinetics are significantly slower.
Both compounds permit less than two orders of magnitude increase in [Ca2þ]i,generally to only the low micromolar range, and then only with very bright flashes
or prolonged exposures to steady light to achieve complete photolysis. Nitr-8 per-
mits a much larger change in [Ca2þ]i. Photolysis kinetics for this compound have
not yet been reported. Neither nitr-8 nor the control compound nitr-9 is presently
commercially available; nitr-5 and nitr-7 are supplied by CalBiochem (La Jolla,
California).
Table IProperties of photosensitive Ca chelators
Compound
(availability)
tPhot(ms)
lmax
(nm) Q.E.Ca Q.E.free
Before
photolysis
After
photolysis
Before
photolysis
After
photolysisCa-bind-
ing on-
rate
KD-Ca
(mM)
KD-Mg
(mM)
KD-Ca
(mM)
KD-Mg
(mM)
e10-Ca(M�1 cm�1)
e10-free(M�1 cm�1)
e10-Ca(M�1 cm�1)
e10-free(M�1 cm�1)
Nitr-5 0.27 365 0.035 0.012 0.145 8.5 6.3 8 5450 5750 13,800 27,300 0.5
Nitr-7 1.8 365 0.042 0.011 0.054 5.4 3.0 5 5780 5540 24,700 10,000 0.2
Nitr-8 – 365 0.026b – 0.5 – 1370 – 11,000 1100 �50,000 �20,000 –
Nitr-9 – 365 �0.02 �0.02 �1000 �10 �1000 �10 �5500 �5500 �15,000 �25,000 –
Azid-1 <2.0 342 1.0 0.9 0.23 8 120 8 33,000 27,000 11,500 5550 0.8
DM-
nitrophen
0.015;1.9 370 0.18 0.18 0.007c 0.0017c 4200;89 2.5 4330 4020 3150 3150 0.02
NP-EGTA 0.002 345 0.23 0.23 0.08c 9 1000 9 975 975 1900 1900 0.017
DMNPE-4 – 347 0.09 0.09 0.048d 7 1000 7 5140 5140? 5140? 5140? 0.01?
NDBF-
EGTA
0.01;0.52 330 0.7 0.7 0.1 15 1000 15 18,400 18,400? 18,400? 18,400? –
Diazo-2 0.134 370 0.057a 0.030a 2.2 5.5 0.073 3.4 2080 22,200 700 2080 0.8
Diazo-4 – 370 0.030a,b 0.030a,b 89 – 0.055 2.6 4600 46,000 <500 <500 0.8
Diazo-3 0.24 375 0.048 0.048 >1000 20 >1000 20 2100 22,800 700 2100 0.8
tPhot, photolysis time constant; lmax, absorbance maximum of Ca-loaded compound; most eVective photolysis wavelength; Q.E.Ca,free, quantum eYciency,
Ca-bound (free); KD-Ca,Mg, Ca (Mg) dissociation constant (1/aYnity) at 0.1–0.15 M ionic strength; and e10-Ca,free, decadic absorbance extinction coeYcient of
Ca-bound (free) compound.a10% of absorbed photons produce a nonphotolyzable photoproduct similar in absorbance and aYnities to unphotolyzed diazo.bFor photolysis of each site.cAt pH 7.2; doubles for each 0.3 pH unit reduction.dAt pH 7.2; increases 2.5� for each 0.2 pH unit reduction.
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 33
The latest addition to the nitr-like class of compounds based on BAPTA is
Azid-1 (Adams et al., 1997). This compound was derived from the high-aYnity
fluorescent indicator derivative of BAPTA, fura-2, by addition of an azido substit-
uent to fura’s benzofuran-3 position. Unlike fura-2, neither this compound nor its
photoproducts are fluorescent; and unlike the other nitr compounds and the
dimethoxynitrophenyl class of Ca2þ chelators (see below), it relies on the photo-
sensitivity of an aromatic azide rather than a nitrobenzyl group. UV absorption
peaking at 372 nm (342 nm for the Ca2þ-bound form) probably leads to formation
of a nitrene which steals hydrogen from water to produce an amidine, which with
another hydrogen converts to a nitrenium that rapidly combines with water to
form an amidinium that reacts with OH� to produce the final low-aYnity electron-
withdrawing benzofurane-3-one photoproduct plus ammonia. Thus, photolysis
absorbs one net proton and produces one molecule of ammonia for each molecule
of azid-1 photolyzed, which can lead to an elevation of pHi in weakly buVered cells.
This disadvantage is counterbalanced by substantial advantages. Photolysis of
both Ca2þ-bound and Ca2þ-free forms of zaid-1 is phenomenally eYcient
(Q.E.�1), and azid-1 is very UV-dark, absorbing at 33,000 M�1 cm�1 when
Ca2þ-bound (or 27,000 M�1 cm�1 when free); these factors combine to make it
250–300 times more sensitive to light than nitr-5! Moreover, its Ca2þ-aYnity drops
from 230 nM to 120 mM, on photolysis, a change that is 12 times the change in nitr-
5 aYnity on photolysis. Like the nitr compounds, it hardly binds Mg2þ at all
(KD¼8 mM), and its Ca2þ-binding (�109 M�1 s�1) and photolysis rates (t<2 ms)are equally rapid. In most respects, azid-1 comes closest to the ideal-caged Ca2þ
compound. Unfortunately, its synthesis is quite diYcult, and it has never been
commercially available; at present, apparently none exists at all.
B. Calculating [Ca2þ]i Changes in Cells
If nitr-5 or azid-1 is photolyzed partially by a flash of light, the reduction in Ca2þ
aYnity of a portion of the chelator occurs within �0.3 ms. During this period of
photolysis, low-aYnity buVer is being formed and high-aYnity buVer is vanishingwhile the total amount of Ca2þ remains unchanged. As the buVer concentrationschange, Ca2þ ions reequilibrate among the new buVer concentrations by shifting
from the newly formed low-aYnity photoproduct to the remaining unphotolyzed
high-aYnity caging chelator. Since the on-rate of binding is close to the diVusionlimit (as calculated from Adams et al., 1988; see also Ashley et al., 1991b), this
equilibration occurs much faster than photolysis, and Ca2þ remains in quasi-
equilibrium throughout the photolysis period. The [Ca2þ]i in a cell rises smoothly
in a step-like fashion over a period of 0.3 ms from the low level determined by the
initial concentrations of total Ca2þ, and unphotolyzed chelator to a higher level
determined by the final concentrations of all the chelator species after partial
photolysis. At least in the case of nitr-5, [Ca2þ]i remains under the control of the
low- and high-aYnity species, so the elevated Ca2þ is removed only gradually by
extrusion and uptake into organelles. Thus, nitr-5 and azid-1 are well suited to
34 Robert Zucker
producing a modest but quantifiable step-like rise in response to a partially
photolyzing light flash, or a gradually increasing [Ca2þ]i during exposure to steady
light. Subsequent flashes cause further increments in [Ca2þ]i. These increments
actually increase because, with each successive flash, the remaining unphotolyzed
chelator is loaded more heavily with Ca2þ. Eventually, unphotolyzed nitr-5 or
azid-1 is fully Ca2þ-bound, and subsequent flashes elevate Ca2þ by smaller incre-
ments as the amount of unphotolyzed chelator drops.
If a calibrated light source is used that photolyzes a known fraction of nitr in the
light path, or in cells filled with chelator and exposed either fully or partially to
light, then the mixture of unphotolyzed nitr and photoproducts may be calculated
with each flash (Lando and Zucker, 1994; Lea and Ashley, 1990). The diVerentquantum eYciencies of free and Ca2þ-bound chelators must be taken into account.
Simultaneous solution of the buVer equations for photolyzed and unphotolyzed
chelators and native Ca2þ buVers predicts the [Ca2þ]i. For suYciently high nitr-5
concentration (above 5 mM), the native buVers have little eVect and usually may
be ignored in the calculation. Further, since [Ca2þ]i depends on the proportion of
chelator loaded with Ca2þ, the exact chelator concentration in the cell makes little
diVerence, at least in small cells or cell processes.
If the cell is large, the light intensity will drop as it passes from the front to the
rear of the cell. Knowing the absorbance of cytoplasm and chelator species at
360 nm, and the chelator concentration before a flash, the light intensity and
photolysis rate at any point in the cytoplasm may be calculated. A complication
in this calculation is that nitr-5 photoproducts have very high absorbance (Ca2þ-free photoproduct, 24,000 M�1 cm�1 and Ca2þ-bound photoproduct,
10,000 M�1 cm� l) (Adams et al., 1988). As photolysis proceeds, the cell darkens
and photolysis eYciency is reduced by self-screening. For azid-1 the situation is
reversed: its photoproducts have much lower absorbance (11,500 and
5000 M�1 cm�1) for Ca2þ-bound and free species or 1/3 and 1/4 of the respective
unphotolyzed forms. Thus light penetrates more deeply as photolysis proceeds.
Regardless, with estimation of the spatial distribution of light intensity from Beer’s
Law, the spatial concentrations of photolyzed and unphotolyzed chelator can be
computed; from this calculation follows the distribution of the rise in [Ca2þ]i. Thesubsequent spatial equilibration of [Ca2þ]i can be calculated by solving diVusionequations, often in only one dimension, using the initial [Ca2þ]i and chelator
distributions as the boundary conditions. EVects of endogenous buVers, uptake,and extrusion mechanisms on the rise in [Ca2þ]i can be incorporated into the
calculations. Simulations of the temporal and spatial distribution of [Ca2þ]i havebeen devised (Zucker, 1989) and applied to experimental data on physiological
eVects of [Ca2þ]i; the predicted changes in [Ca2þ]i have been confirmed with Ca2þ-sensitive dyes (Lando and Zucker, 1989). Simplified and approximate models using
the volume-average light intensity to calculate volume-average photolysis rate and
average [Ca2þ]i changes often suYce when the spatial distribution of [Ca2þ]i is notimportant, for example, in small cells or processes or when estimating the change in
[Ca2þ]i in a cell after diVusional equilibration has occurred.
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 35
III. DM-Nitrophen
A. Chemical Properties
Graham Ellis-Davies followed a diVerent strategy for releasing Ca2þ—by
attaching a 2-nitrobenzyl group to one of the chelating amines of ethylenediami-
netetraacetic acid (EDTA) to form the photosensitive chelator dimethoxynitro-
phenyl-EDTA or DM-nitrophen (Ellis-Davies and Kaplan, 1988; Kaplan and
Ellis-Davies, 1988). Photolysis by UV light in the wavelength range 330–380 nm
cleaves the DM-nitrophen with a quantum eYciency of 0.18 through multiple
intermediate pathways (McCray et al., 1992) to form iminodiacetic acid and a
Hþ-absorbing 2-nitrosoacetophenone derivative, with 65% of the photoproducts
formed with a time constant of 15 ms and the rest with t¼1.9 ms (Faas et al., 2005,
2007). A simplified reaction is shown in Fig. 3. Although DM-nitrophen binds
Ca2þ with an aYnity of 7 nM at pH 7.2, Ca2þ-bound chelator forms a photoprod-
uct-binding Ca2þ with a 4-mM aYnity, while the free form (and Mg2þ-boundforms, see below) photolyze to a 90 mM-KD photoproduct at an ionic strength of
150 mM. These values are from the most recent of the continually evolving models
of DM-nitropen photolysis (Ayer and Zucker, 1999; Bollmann and Sakmann,
2005; Faas et al., 2005, 2007; Kaplan and Ellis-Davies, 1988; Neher and Zucker,
1993). Thus, complete photolysis of Ca2þ-DM-nitrophen can elevate Ca2þ over
50,000-fold, much more than photolysis of the nitr compounds or azid-1. This
significant advantage is counterbalanced to some extent by the facts that the
photoproducts buVer Ca2þ so weakly that the final [Ca2þ]i will be determined
largely by native cytoplasmic buVers, and that the Ca2þ liberated by photolysis of
DM-nitrophen will be removed more readily by extrusion and uptake pumps.
CO2−
CO2−
CO2−
CO2− CO2
−
CO2−
−O2C
−O2CCa2+
Ca2+
NO2 NO
ON
DM-nitrophen
(high Ca2+ affinity)
DM-nitrophen photolysis products
(low Ca2+ affinity)
OCH3
OCH3 OCH3OCH3
N N
NH
+hn
Fig. 3 Structure of and reaction scheme for DM-nitrophen, which releases calcium on exposure to
light.
36 Robert Zucker
The absorbance of Ca2þ-saturated and free DM-nitrophen is 4330 and
4020 M�1 cm�1, respectively, and 0.18.
A serious complication of DM-nitrophen is that it shares the cation-binding
properties of its parent molecule EDTA. In particular, Hþ and Mg2þ compete for
Ca2þ at the hexacoordinate-binding site. The aYnity of DM-nitrophen for Mg2þ
at pH 7.2 is 1.7 mM, whereas the photoproducts bind Mg2þ with aYnities of about
2 mM. Further, both the Ca2þ- and Mg2þ-aYnities of DM-nitrophen are highly
pH-dependent (Grell et al., 1989), doubling for each 0.3 units of pH increase. Thus,
in the presence of typical [Mg2þ]i levels of 1–3 mM, DM-nitrophen that is not
already bound to Ca2þ will be largely in the Mg2þ-bound form. Further, excess
DM-nitrophen will suck Mg2þ oV ATP, which binds it substantially more weakly,
compromising the ability of ATP to serve as an energy source or as a substrate for
ATPases. Finally, photolysis of DM-nitrophen will lead to a jump in [Mg2þ]i aswell as [Ca2þ]i, and to a rise in pH. Unless controlled by native or exogenous pH
buVers, this pH change can alter the Ca2þ and Mg2þ aYnities of the remaining
DM-nitrophen. In the absence of Ca2þ-loading, DM-nitrophen even may be used
as a caged Mg2þ chelator (Ellis-Davies, 2006). Attributing physiological responses
to a [Ca2þ]i jump, therefore, requires control experiments in which DM-nitrophen
is not charged with Ca2þ. DM-nitrophen currently is sold by CalBiochem.
To circumvent the problems arising fromMg2þ competing for the Ca2þ-bindingsite of DM-nitrophen, a second generation derivative of ethylene glycol bis(b-aminoethylether)-N,N,N0,N0-tetraacetic acid (EGTA, which binds Mg2þ only
very weakly) coupled to a light-sensitive ortho-nitrophenyl group was developed
(Ellis-Davies and Kaplan, 1994). This compound, nitrophenyl-EGTA or NP-
EGTA, is very rapidly cleaved (t¼2 ms) (Ellis-Davies, 2003) to Hþ-absorbingimidodiacetic acid photoproducts with eVective Ca2þ-KD of 1 mM, 12,500-fold
higher (lower aYnity) than that of the unphotolyzed cage (80 nM) at pH 7.2, with
pH-dependence similar to that of EGTA, EDTA, and DM-nitrophen. Unlike DM-
nitrophen, Mg2þ binding to NP-EGTA is negligible (9 mM before and after
photolysis). Quantum eYciency (0.23) is similar to that of DM-nitrophen, and
higher than for the nitr compounds, but less than that of azid-1. However,
photolysis eYciency is seriously limited by its low absorbance (975 M�1 cm�1),
only 1/6—1/4 those of the nitr compounds and DM-nitrophen, and less than 3% of
azid-1’s absorbance.
More recently, a dimethoxy-ortho-nitrophenyl derivative of EGTA (DMNPE-4)
was introduced (Ellis-Davies and Barsotti, 2006), with somewhat higher Ca2þ
aYnity (48 nM), dropping with time constants of 10 and 17 ms to 1 mM on
photolysis, low Mg2þ-aYnity (7 mM), and under half the quantum eYciency
(0.09) but over five times the absorbance (5140 M�1cm�1), thus twice the photoly-
sis eYciency of NP-EGTA. An additional very slow phase releasing 30% of caged
Ca2þ with t�667 ms was observed.
Ellis-Davies’ lab has also produced a new generation of EGTA-based chelators
using the novel photosensitive chromophore nitrodibenzofuran or NDBF-EGTA
(Momotake et al., 2006). This compound binds Ca2þ with KD¼100 nM at pH 7.2,
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 37
presumably with an on-rate similar to that of the other EGTA derivatives. Ca2þ
aYnity drops sharply to �1 mM on photolysis with time constants of 14 and
520 ms. Quantum eYciency (0.7) and absorbance (18,400 M�1 cm�1) are extremely
high, as is the change in Ca2þ aYnity (10,000-fold at pH 7.2), making this a very
attractive candidate for future-caged Ca2þ research.
B. Calculating Changes in [Ca2þ]i
Calculating [Ca2þ]i changes on photolysis of NP-EGTA and its congeners issimilar to that for the nitr compounds (if the pH dependence of binding constants
is ignored), sinceMg2þ binding is not an issue. Since the chelators’ Ca2þ aYnities is
similar to resting cytoplasmic [Ca2þ]i levels, filling cells with a half-Ca2þ-loadedchelator will not disturb [Ca2þ]i but can release substantial amounts of Ca2þ
(�1 mM) which will be reduced about 100-fold by the cell’s endogenous buVers.However, except for NDBF-EGTA, the low absorbance usually limits flash pho-
tolysis to at most about 20%.
Quantifying changes in [Ca2þ]i caused by photolysis is much more diYcult for
DM-nitrophen. The initial level of [Ca2þ]i before photolysis depends upon the total
concentrations of Mg2þ, Ca2þ, DM-nitrophen, ATP, and native Ca2þ buVers,because at least two buVers (DM-nitrophen and endogenous buVers) compete
for Ca2þ, two buVers (ATP and DM-nitrophen) compete for Mg2þ, and, afterpartial photolysis, both cations also bind to the two photoproducts. Calculating
equilibrium Ca2þ levels involves simultaneous solution of at least six nonlinear
buVer equations (Delaney and Zucker, 1990), which is a tedious chore at best.
Also, the various dissociation constants depend on ionic strength, and have been
measured only at 150 mM. The high aYnity of DM-nitrophen for Ca2þ might
appear to dominate the buVering of Ca2þ in cytoplasm, but this idea is misleading.
A solution of DM-nitrophen that is 50% saturated with Ca2þ will hold the free
[Ca2þ]i at 7 nM at pH 7.2; this action will be independent of the total DM-
nitrophen concentration. However, 5 mM DM-nitrophen with 2.5 mM Ca2þ and
5 mM Mg2þ will buVer free [Ca2þ]i to about 2 mM; now doubling all concentra-
tions results in a final [Ca2þ]i of around 5 mM. Since the total [Mg2þ]i available, asfree or weakly bound to ATP, is several millimolar, partially Ca2þ-loaded DM-
nitrophen may bring the resting Ca2þ level to a surprisingly high level. Because the
solution is still buVered, this [Ca2þ] may be reduced only gradually by pumps and
uptake, but eventually Ca2þwill be pumped oV the DM-nitrophen until the [Ca2þ]iis restored to its normal level. Then photolysis may lead to only tiny jumps in
[Ca2þ]i. However, if a large amount of Ca2þ-loaded-DM-nitrophen is introduced
into a cell relative to the total [Mg2þ]i, Ca2þ can be buVered to low levels while
photolysis can release a large amount. In fact, if enough DM-nitrophen is intro-
duced into cells with no added Ca2þ, it may gradually absorb Ca2þ from cytoplasm
and intracellular stores and photolysis can produce a substantial jump in [Ca2þ]i.Therefore, both resting and the postphotolysis levels of Ca2þ may vary over very
wide ranges, depending on [DM-nitrophen]i, [Mg2þ]i, and cellular [Ca2þ]i control
38 Robert Zucker
processes, all of which are diYcult to estimate or control. Thus, quantification of
changes in [Ca2þ]i is not easy to achieve.
The situation may be simplified by perfusing cells with Ca2þ-DM-nitrophen
solutions while dialyzing out Mg2þ and mobiles endogenous buVers (Neher and
Zucker, 1993; Thomas et al., 1993). Of course, this procedure will not work in
studies of cell processes requiring Mg2þ-ATP or if perfusion through whole-cell
patch pipettes is not possible.
Another consequence of Mg2þ binding by DM-nitrophen is that cytoplasmic
Mg2þ may displace Ca2þ from DM-nitrophen early in the injection or perfusion
procedure, leading to a transient rise in [Ca2þ]i before suYcient DM-nitrophen is
introduced into the cell (Neher and Zucker, 1993; Parsons et al., 1996; Thomas
et al., 1993). Such a ‘‘loading transient’’ was accurately predicted from models of
changes of the concentrations of total [Ca2þ]i, [Mg2þ]i, ATP, native buVer, andDM-nitrophen during filling from a whole-cell patch electrode (R. S. Zucker,
unpublished). Since this process may have important physiological consequences,
controlling it is important. The process may be eliminated largely by separating the
Ca2þ-DM-nitrophen-filling solution in the pipette from the cytoplasm by an
intermediate column of neutral solution [such as dilute EGTA or BAPTA] in the
tip of the pipette, which allows most of the Mg2þ to escape from the cell before the
DM-nitrophen begins to enter. Then most of the loading transient occurs within
the tip of the pipette.
One method of better controlling the change in [Ca2þ]i in DM-nitrophen experi-
ments is to fill cells with a mixture of Ca2þ-DM-nitrophen and another weak Ca2þ
buVer such as N-hydroxyethylethylenediaminetriacetic acid (HEEDTA) or l,3-
diaminopropan-2-ol-tetraacetic acid (DPTA) (Neher and Zucker, 1993). These
tetracarboxylate Ca2þ chelators have Ca2þ aYnities in the micromolar or tens of
micromolar range. If cells are filled with such a mixture without Mg2þ, the initialCa2þ level can be set by saturating the DM-nitrophen and adding appropriate
Ca2þ to the other buVer. Then photolysis of DM-nitrophen releases its Ca2þ onto
the other buVer; the final Ca2þ can be calculated from the final buVer mixture in
the same fashion as for the nitr compounds. Since all the constituent aYnities are
highly pH dependent, a large amount of pH buVer (e.g., 100 mM) should be
included in the perfusion solution, and the pH of the final solution adjusted
carefully.
The kinetic behavior of DM-nitrophen and the NP-EGTAs is much more
complex than their equilibrium reactions. Photolysis proceeds rapidly (ts¼0.2
and 2 ms for DM-nitrophen, 2 ms for NP-EGTA), but the on-rate of Ca2þ binding
is much slower, about 20 mM�1 ms�1 (Ellis-Davies, 2003; Faas et al., 2005, 2007).
This characteristic has particularly interesting consequences for partial photolysis
of partially Ca2þ-loaded chelator. A flash of light will release some Ca2þ, whichinitially will be totally free. If the remaining unphotolyzed and unbound chelator
concentration exceeds that of the released Ca2þ, this Ca2þ will rebind, displacing
Hþ within milliseconds and producing a brief [Ca2þ]i ‘‘spike’’ (Ellis-Davies et al.,
1996; Grell et al., 1989; Kaplan, 1990;McCray et al., 1992), followed by a near-step
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 39
fall in pH. IfMg2þ is also present, a secondary relaxation of [Ca2þ]i follows becauseof the slower displacement of Mg2þ from DM-nitrophen (Ayer and Zucker, 1999;
Delaney and Zucker, 1990; Escobar et al., 1995, 1997). Moreover, if a steady UV
source is used to photolyze DM-nitrophen, rebinding continually lags release,
leading to a low (micromolar range) free [Ca2þ]i while the illumination persists.
When the light is extinguished, the [Ca2þ]i drops rapidly to a low level under control
of the remaining chelator (Zucker, 1993). In the case of DM-nitrophen bound to
Mg2þ, achievement of equilibrium is somewhat slower (tens of milliseconds). Thus,
a reversible ‘‘pulse’’ of [Ca2þ]i is generated, the amplitude of which depends on light
intensity and the duration of which is controlled by the length of the illumination.
This situation remains so until the remaining unphotolyzed cage becomes fully
saturated with Ca2þ, whereupon [Ca2þ]i escapes from the control of the chelator,
imposing a practical limit on the product of [Ca2þ]i and duration of about
0.75 mM s for DM-nitrophen. Similar kinetic considerations apply when Ca2þ is
passed by photolysis from a Ca2þ cage to another buVer such as BAPTA, EGTA,
HEEDTA, orDPTA. Judicious selection of buVers and buVer ratiosmay be used to
shape this Ca2þ ‘‘spike’’ to match a hypothetical naturally occurring [Ca2þ]i(t)waveform and test its physiological consequences (Bollmann and Sakmann,
2005). If this behavior is considered undesirable, it may be avoided by using only
fully Ca2þ-saturated DM-nitrophen, due to its extremely high Ca2þ-aYnity, for
which rebinding to unphotolyzed chelator is impossible. Thus, the kinetic complex-
ity of the nitrophen class of chelators can be turned to experimental advantage,
greatly magnifying the flexibility of experimental [Ca2þ]i control.
IV. Diazo Compounds
A. Chemical Properties
In some experiments, being able to lower the [Ca2þ]i rapidly, rather than raise it,
is desirable. For this purpose, caged chelators were developed. Initial attempts
involved attachment of a variety of photosensitive protecting groups to mask one
of the carboxyl groups of BAPTA, thus reducing its Ca2þ aYnity until restored by
photolysis. Such compounds displayed low quantum eYciency (Adams et al., 1989;
Ferenczi et al., 1989) and their development has not been pursued. A more suc-
cessful approach (Adams et al., 1989) involved substituting one (diazo-2) or both
(diazo-4) of the aromatic rings of BAPTA with an electron-withdrawing diazoke-
tone that reduces Ca2þ aYnity, much like the photoproducts of the nitr com-
pounds. Figure 4 shows the structures of the diazo series of chelators. Photolysis
converts the substituent to an electron-donating carboxymethyl group while
releasing a proton; the Ca2þ aYnity of the photoproduct is thereby increased.
The reaction is illustrated in Fig. 5.
Diazo-2 absorbs one photon with quantum eYciency 0.03 to increase aYnity, in
433 ms, from 2.2 mM to 73 nM at 120-mM ionic strength (or to 150 nM at 250 mM
COO−
N N N N N
Mediazo-2 diazo-3 diazo-4
O O OMe
O
O O
OOO
N+ N+ N+ +N
N−N− N− −N
COO− COO− COO− COO− COO− COO− COO− COO− COO−
Fig. 4 Structures of the diazo series of photolabile chelators, which take up calcium on exposure to
light.
O
OCH
CH3
N2
−O2C
−O2C −O2C−O2C
−O2C−O2C
O O O O
H+
O
O
C+HC N2
N
CO2−
CO2−
N N N N N
hn
CO2−
CO2−
CO2−
CO2−
CO2−
CH3 CH3
H2O
H2C +
Ca2+
Diazo-2(low Ca2+ affinity)
Photolyzed diazo-2(high Ca2+ affinity)
Fig. 5 Reaction scheme for the photolysis of diazo-2.
40 Robert Zucker
ionic strength). The absorbance maximum of the photosensitive group is
22,200 M� l cm�1 at 370 nm, and drops to negligible levels at this wavelength
after photolysis. A small remaining absorbance reflects formation of a side product
of unenhanced aYnity and unchanged molar extinction coeYcient in 10% of the
instances of eVective photon absorption. This ‘‘inactivated’’ diazo still binds Ca2þ
(with some reduction in absorbance), but is incapable of further photolysis. The
Ca2þ-bound form of diazo-2 has about one-tenth the absorbance of the free form,
dropping to negligible levels after photolysis, with quantum eYciency of 0.057 and
a time constant of 134 ms. Binding of Ca2þ to photolyzed diazo-2 is fast, with an
on-rate of 8�108 M�1 s�1. Mg2þ binding is weak, dropping from 5.5 to 3.4 mM
after photolysis, and pH interference is small with this class of compound.
One limitation of diazo-2 is that the unphotolyzed chelator has suYcient Ca2þ
aYnity that its incorporation into cytoplasm is likely to reduce resting levels to
some degree, and certainly will have some eVect on [Ca2þ]i rises that occur
physiologically. To obviate this problem, diazo-4 was developed with two
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 41
photolyzable diazoketones. Absorption of one photon increases the Ca2þ aYnity
from 89 to 2.2 mM (with a 10% probability of producing a side-product with one
inactivated group). Absorption of two photons (with a probability assumed to
equal the square of the probability of one group absorbing one photon, and with a
measured quantum eYciency of 0.015) results in further increase of the aYnity to
55 nM, a total increase of 1600-fold. This large increase in aYnity is, to some
extent, oVset by the small fraction of diazo-4 that can be doubly photolyzed
readily. Thus, a flash of light produces a variety of species: unphotolyzed, singly
photolyzed, doubly photolyzed, singly inactivated, doubly inactivated, and singly
photolyzed-singly inactivated, with a variety of transition probabilities among
species (Fryer and Zucker, 1993). Unphotolyzed diazo-4 is highly absorbent
(46,000 M�1 cm�1 at 371 nm for the free form; about 4600 M�1 cm�1 for the
Ca2þ-bound form). The singly photolyzed species have absorbances of half these
values and doubly photolyzed diazo-4 has negligible absorbance at this wave-
length. Inactivation causes little change in absorbance.
A third member of this series, diazo-3, has a diazoketone attached to half the
cation-coordinating structure of BAPTA, and has negligible Ca2þ aYnity.
On photolysis, diazo-3 produces the photochemical intermediates of diazo-2 plus
a proton, and may be used to control for these eVects of photolysis of the diazo
series. At one time, diazo-2 and diazo-3 (but not diazo-4) were commercially
available (Molecular Probes, Eugene, and Oregon), but these stocks appear to
have been exhausted.
B. Calculating EVects of Photolysis
As for the nitr compounds, equilibration is faster than photolysis, so a flash of
light leads to a smooth step transition in the concentration of Ca2þ chelator
species. If the percentage of photolysis caused by a light flash is known, the
proportions of photolyzed and inactivated diazo-2, or of the six species of diazo-
4, can be calculated. Usually, diazo is injected without any added Ca2þ, so the
eVect of photoreleased buVers is to reduce the [Ca2þ]i from its resting value. This
change can be calculated only if the total Ca2þ bound to the native buVer in
cytoplasm as well as the characteristics of that buVer are known. These character-istics often can be inferred from available measurements on cytoplasmic Ca2þ
buVer power and the normal resting [Ca2þ]i level. The more usual application of
these substances is to reduce the eVect of a physiologically imposed rise in [Ca2þ]i.In many cases, the magnitude of the source of this Ca2þ is known, as in the case of
a Ca2þ influx measured as a Ca2þ current under voltage clamp or the influx
through single channels estimated from single channel conductances. Also, the
magnitude of the total Ca2þ increase in a response can be estimated frommeasured
increases in [Ca2þ]; and estimates of cytoplasmic buVering. With this information,
the expected eVect of newly formed diazo photoproducts on a physiological rise in
[Ca2þ]i can be calculated by solving diVusion equations that are appropriate for the
distribution of Ca2þ sources before and after changing the composition of the
42 Robert Zucker
mixture of buVers in the cytoplasm. Examples of such solutions of the diVusionequation exist for spherical diVusion inward from the cell surface (Nowycky and
Pinter, 1993; Sala and Hernandez-Cruz, 1990), cylindrical diVusion inward from
membranes of nerve processes (Stockbridge and Moore, 1984; Zucker and
Stockbridge, 1983), diVusion from a point source (Fryer and Zucker, 1993;
Stern, 1992), and diVusion from arrays of point sources (Fogelson and Zucker,
1985; Matveev et al., 2002, 2004, 2006, 2009; Pan and Zucker, 2009; Simon and
Llinas, 1985; Tang et al., 2000; Yamada and Zucker, 1992). For large cells, the
spatial nonuniformity of light intensity and photolysis rate also must be consid-
ered, taking into account the absorbances of all the species of diazo and the
changes in their concentration with photolysis. Like azid-1, the self-screening
imposed by diazo chelators is reduced with photolysis, so successive flashes (or
prolonged illumination) are progressively more eVective.
V. Introduction into Cells
Photolabile chelators are introduced into cells by pressure injection from micro-
pipettes, perfusion from whole-cell patch pipettes, or permeabilization of the cell
membrane. Iontophoresis is also suitable for diazo compounds, since this proce-
dure inserts only the Ca2þ-free form. For the caged Ca2þ substances, this method
of introduction requires that the chelator load itself with Ca2þ by absorbing it from
cytoplasm or intracellular stores. Filling cells from a patch pipette has the special
property that, if the photolysis light is confined to the cell and excludes all but the
tip of the pipette inside the cell, the pipette barrel acts as an infinite reservoir of
unphotolyzed chelator. Then the initial conditions of solutions in the pipette can be
restored within minutes after photolysis of the chelator in the cell. The nitr and
diazo compounds are soluble at concentrations over 100 mM and DM-nitrophen
is soluble at 75 mM, so levels in cytoplasm exceeding 10 mM can be achieved
relatively easily, even by microinjection, making the exogenous chelator com-
pound the dominant buVer.Nitr and diazo chelators also have been produced as membrane-permeant acet-
oxymethyl (AM) esters (Kao et al., 1989). Exposure of intact cells to medium
containing these esters (available from CalBiochem and Molecular Probes, respec-
tively) might result in the loading of cells with nearly millimolar concentrations, if
suYcient activity of intracellular esterase is present to liberate the membrane-
impermeant chelator. However, nitr-5 or nitr-7 introduced in this manner is not
bound to Ca2þ, so it must sequester Ca2þ from cytoplasm, from intracellular stores,
or after Ca2þ influx is enhanced, for example, by depolarizing excitable cells. The
final concentration, level of Ca2þ loading, and localization of the chelator are
uncertain, so this method of incorporation does not lend itself to quantification
of eVects of photolysis unless cells are coloaded with a Ca2þ indicator.
During loading and other preparatory procedures, the photolabile chelators may
be protected fromphotolysis with low passUV-blocking filters in the light path of the
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 43
tungsten or quartz halide beams used for viewing. For more detail on these filling
procedures, see Gurney (1991). Other methods of loading cells, used primarily with
other sorts of caged compounds, are discussed by Adams and Tsien (1993).
VI. Light Sources
Photolysis of caged Ca2þ chelators requires a bright source of near UV light.
If speed is unimportant, an ordinary mercury or xenon arc lamp may be used.
Mercury lamps have a convenient emission line at 366 nm. Exposure can be
controlled with a shutter, using MgF-coated Teflon blades for particularly bright
sources. Lamps of 100–150 W power with collimating quartz lenses provide suY-cient energy to photolyze�25% of caged Ca2þ compounds in�2 s. Bulbs of larger
power only generate bigger arcs, with more energy in a larger spot of similar
intensity. With additional focusing, photolysis can be achieved in one-tenth the
time or even less. These light sources are the appropriate choice in applications
using reversible [Ca2þ]i elevation with DM-nitrophen.
Fast events require the use of a laser or xenon arc flashlamp. The xenon lamps
are less expensive and cumbersome; convenient commercial systems are available
from Chadwick-Helmuth (El Monte, California), Rapp Optoelektronik (Hamburg,
Germany), TILL Photonics (Grafelting, Germany), and Cairn Research (Faver-
sham, UK). These flashlamps discharge up to 200–300 J electrical energy across
the bulb to provide a pulse of �1 ms duration with up to 300 mJ energy in the
330�380-nm band. The Chadwick-Helmuth unit includes only a power supply and
lamp socket, so a housing with focusing optics must be constructed (see Rapp and
Guth, 1988). Focusing can be accomplished with a UV-optimized elliptical reflec-
tor or with quartz refractive optics. The reflector can be designed to capture more
light (i.e., have a larger eVective numerical aperture), but reflectors have greater
physical distortion than well-made lenses. In practice, the reflector generates a
larger spot with more total energy, but somewhat less intensity, than refractive
methods. One advantage of reflectors is that they are not subject to chromatic
aberration—focusing is independent of wavelength—so the UV will be focused in
the same spot as visual light. This is not true of refractive lenses. To focus and aim
them accurately at the sample, a UV filter must be used to block visual light and the
beam must be focused on a fluorescent surface for visualization. Both types of
housing are available from Rapp Optoelektronik. Using either system, photolysis
rates approaching 80–90% in one flash are achievable. This rate may be reduced by
imposing neutral density filters or reducing discharge energy, but the relationship
between electrical and light energy is not linear and should be measured with a
photometer. Flashlamps can be reactivated only after their storage capacitors have
recharged, setting the minimal interval between successive maximal flashes at
several seconds or more.
F1ashlamps are prone to generating a number of artifacts. The discharge causes
electrical artifacts that can burn out semiconductors and op amps, and reset or
44 Robert Zucker
clear digital memory in other nearby equipment. Careful electrostatic shielding,
wrapping inductors with paramagnetic metal, power source isolation, and using
isolation circuits in trigger pulse connections to other equipment prevent most
problems, which are also diminished in pulsed mercury lamps (Denk, 1997). The
discharge generates a mechanical thump at the coil used to shape the current pulse
through the bulb; this thump can dislodge electrodes from cells or otherwise
damage the sample. Mechanical isolation of the oVending coil solves the problem.
Lamp discharge also produces an air pressure pulse that can cause movement
artifacts at electrodes, which can be seen to oscillate violently for a fraction of a
second when videotaped during a flash. This movement can damage cells severely,
especially those impaled with multiple electrodes. Small cells sealed to the end of a
patch pipette often fare better against such mistreatment. To reduce this source of
injury, the light can be filtered to eliminate all but the near UV. Commercial Schott
filters (UG-I, UG-Il), coated to reflect infrared (IR) light, serve well for this
purpose, but can cut the 330�380-nm energy to 30% or less. Liquid filters to
remove IR and far UV also have been described (Tsien and Zucker, 1986).
Removing IR reduces temperature changes, which otherwise can exceed 1 �C per
flash, whereas removing far UV prevents the damaging eVects of ionizing radia-
tion. Chlorided silver pellets and wires often used in electrophysiological recording
constitute a final source of artifact. These components must be shielded from the
light source or they will generate large photochemical signals.
To simply aim and focus the light beam directly onto the preparation is easiest.
If isolating the lamp from the preparation is necessary, the light beam may be
transmitted by a fiber optic or liquid light guide, with some loss of intensity. If a
microscope is being used already, the photolysis beam may be directed through the
epifluorescence port of the microscope. The lamp itself, or a light guide, may be
mounted onto this port. Microscope objectives having high numerical aperture
and good UV transmission will focus the light quite eVectively onto a small area,
which can be delimited further by a field stop aperture. With the right choice of
objectives and proper optical coupling of the lamp to the light guide and the guide
to the microscope port, light intensities 25 times greater than those obtained by
simply aiming the focused steady lamp or flashlamp can be achieved—suYcient to
half-photolyze DM-nitrophen in 25 ms of steady bright light. TILL Photonics
make a xenon arc spectrophotometer (the Polychrome) with eYcient optical
coupling to several commercial epifluorescence microscopes. Half reflective mir-
rors can be used to combine the photolysis beam with other light sources, such as
those used for [Ca2þ]i measurement. However, as the optical arrangement becomes
more complex, photolysis intensity inevitably decreases.
The newest development in light sources is the high intensity light-emitting diode
(Bernardinelli et al., 2005). This rapidly evolving and inexpensive technology can
already produce 365-nm UV light at 50 mW/cm2 (with LEDs made by Prizmatix,
Modi’in Ilite, Israel, e.g.), or about20%of the intensityof a collimatedxenonarc lamp.
It is often important to restrict photolysis to one region of a cell (Wang and
Augustine, 1995). With epi-illumination, this may be done with a field stop
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 45
diaphragm, or by conveying the photolysis beam through a tapered quarty fiber
optic filter to the cell surface (Eberius and Schild, 2001; Godwin et al., 1997).
Lasers provide an alternative source of light with the advantages of a coherent
collimated beam that is focused much more easily to a very small spot. Pulsed
lasers such as the frequency-doubled ruby laser or the XeF excimer laser provide at
least 200 mJ energy at 347 or 351 nm in 50 and 10 ns, with possible repetition rates
of 1 and 80 Hz, respectively. Liquid coumarin-dye lasers, with up to 100 mJ
tunable energy in the UV and pulse duration, are also available. Inexpensive
nitrogen lasers providing lower pulse energies (0.25 mJ) in 5-ns pulses at 337 nm
also have been developed (Engert et al., 1996) and, with appropriate focusing,
might be useful. To date, lasers have found their widest application in studies of
muscle contraction. More information on these laser options is contained in
discussions by Goldman et al. (1984) and McCray and Trentham (1989).
An adaptation of laser photolysis is the two-photon absorption technique (Denk
et al., 1990). A colliding-pulse mode-locked Ti:Sapphire laser generating 100-fs
pulses of 630-nm light at 80 MHz is focused through a confocal scanning micro-
scope. Photolysis of UV-sensitive caged compounds requires simultaneous absorp-
tion of two red photons, so photolysis occurs only in the focal plane of the scanning
beam. This behavior restricts photolysis to about 1 mm3 in three dimensions, but for
most compounds the photolysis rate is so slow, due to their extremely limited two-
photon cross sections, that several minutes of exposure are required with currently
available equipment. The best results were achieved with azid-1 and NDBF-EGTA
(Brown et al., 1999; DelPrincipe et al., 1999; Momotake et al., 2006), as expected
from their high single photon absorbances. Azid-1 could be fully photolyzed in the
two-photon focal volume with a 10-ms pulse train of 7 mW average power, with a
retention time of the released Ca2þ in this volume of about 150 ms. This technique isexpensive and specialized, and is still under development, but may have practical
applications in revealing the precise localization within cells or subcellular orga-
nelles of fixed targets of Ca2þ action or of highly localized Ca2þ buVers.Near-UV light alone seems to have little eVect on most biological tissues, with
the obvious exception of photoreceptors and the less obvious case of smooth
muscle (Gurney et al., 1987). Control experiments on the eVects of light on
unloaded cells, and on the normal physiological response under study, can be
used to ascertain the absence of photic eVects.
VII. Calibration
When designing a new optical system or trying a new caged compound, being
able to estimate the rate of photolysis of the apparatus used is important. This
information is necessary to adjust the light intensity or duration for the desired
degree of photolysis, and to insure that photolysis is occurring at all.
In principle, the fraction (F) of a substance photolyzed by a light exposure of
energy J can be computed from the formula e–(J–J0)¼ (I�F)/0.1, where J0 is the
46 Robert Zucker
energy needed to photolyze 90% of the substance and is given by J0 ¼hcA/Qel,where h is Planck’s constant, c is the speed of light, A is Avogadro’s number, Q is
the quantum eYciency, e is the decadic molar extinction coeYcient, and l is the
wavelength of the light. In practice, however, this equation is rarely useful for the
following reasons.
1. Measuring the energy of the incident light on a cell accurately is diYcult,
especially for light of broad bandwidth with varying intensity at diVerentwavelengths.
2. The quantum eYciency, although provided for all the photolabile chelators, is
not such awell-defined quantity. The value depends critically on how it is measured,
which is not always reported. In particular, the eVective quantum eYciency for a
pulse of light ofmoderate duration (e.g., from a flashlamp) is often greater than that
of either weak steady illumination or a very brief pulse (e.g., from a laser), because
of the possibility of multiple photon absorptions of higher eYciency by photochem-
ical intermediates. This phenomenon has been noted to play a particularly strong
role in nitr-5 photolysis (McCray and Trentham, 1989). Thus, apparent diVerencesin quantum eYciencies between diVerent classes of chelators may be mainly the
results of diVerent measurement procedures.
3. Finally, the quantum eYciency is a function of wavelength, which is rarely
given.
A more practical and commonly adopted approach is mixing a partially Ca2þ-loaded photolabile chelator with a Ca2þ indicator in a solution with appropriate
ionic strength and pH buVering, and measuring the [Ca2þ] change in a small
volume of this solution, the net absorbance of which is suYciently small to
minimize inner filtering of the photolyzing radiation. Suitable indicators include
fura-2, indo-1 (Grynkiewicz et al., 1985), furaptra (Konishi et al., 1991), f1uo-3,
rhod-2 (Minta and Tsien, 1989), Calcium GreenTM, OrangeTM, and CrimsonTM
(Eberhard and Erne, 1991), arsenazo III (Scarpa et al., 1978), and fura-red
(Kurebayashi et al., 1993). The choice depends largely on available equipment.
Fura-2, indo-1, and furaptra are dual-excitation or-emission wavelength fluores-
cent dyes, allowing more accurate ratiometric measurement of [Ca2þ], but theyrequire excitation at wavelengths that photolyze the photolabile Ca2þ chelators
and are subject to bleaching by the photolysis light. The former problem may be
minimized by using low intensity measuring light with a high sensitivity detection
system. Furaptra is especially useful for DM-nitrophen, because of its lower Ca2þ
aYnity. Fluo-3 and rhod-2 were designed specifically for use with photolabile
chelators (Kao et al., 1989), being excited at wavelengths diVerent from those
used to photolyze the chelators, but they are not ratiometric dyes and are diYcult
to calibrate accurately. Calcium Green, Orange, and Crimson suVer the same
limitation, but they are often used because of their fast kinetics and bright intensity,
allowing the accurate tracking of fast changes in [Ca2þ]i. Arsenazo and antipyr-
alazo are metallochromic dyes that change absorbance on binding Ca2þ,
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 47
fortunately at wavelengths diVerent from those at which the photolabile chelators
show any significant absorbance. However, these dyes are also diYcult to calibrate
for absolute levels of [Ca2þ], although changes in [Ca2þ] may be determined fairly
accurately. Fura-red is a ratiometric dye excited by visible light, so it might have
some application in calibrating photolysis. A problem common to all the fluores-
cent indicators is that their fluorescent properties may be altered by the presence of
photolabile chelators, which generally are used at millimolar levels whereas the
indicators are present at 100 mM or less. The photolabile chelators often produce
contaminating fluorescence, which also may be Ca2þ-dependent and may partially
quench the fluorescence of the indicators (Hadley et al., 1993; Zucker, 1992). Thus,
the indicators must be calibrated in the presence of photolabile chelator at three
well-controlled [Ca2þ] levels, preferably before and after exposure to the photolysis
flash, before they can be used to measure the eVects of photolysis on [Ca2þ] (Neher
and Zucker, 1993). The low and high [Ca2þ] calibrating solutions may be made
with excess Ca2þ or another buVer such as EGTA or BAPTA, but the intermediate
[Ca2þ] solution is more diYcult to generate, since photolysis of the chelator will
release some Ca2þ and change the [Ca2þ]i and pH in this solution unless it contains
a very high concentration of controlling chelator and pH buVer.The calibration procedure is generally the same for any combination of chelator
and indicator. A small sample of the mixture is placed in a 1-mm length of micro-
cuvette with a 20-mm pathlength (Vitro Dynamics, Rockaway, New Jersey) under
mineral oil to prevent evaporation. This cuvette is exposed repeatedly to the
photolysis beam or to flashes, which should illuminate the whole cuvette uniformly,
and the [Ca2þ] after each flash or exposure is measured using a microscope-based
fluorescence or absorbance photometer. A small droplet of solution under mineral
oil alone would work, and may be necessary if the photolysis beam is directed
through the microscope and illuminates a very small area, but sometimes the
fluorescent properties of the indicators are aVected by the mineral oil. This eVectwould be detected in the procedure for calibrating the chelator-indicator mixture,
but is best avoided using the microcuvettes, in which contact with oil is only at the
edges, the fluorescence or absorbance change of which need not be measured.
In some applications, such as whole-cell patch clamping of cultured cells, using
the cell as a calibration chamber can be easier than any other procedure.
The expected changes in [Ca2þ] depend on the chelator used. The nitr and diazo
chelators should lead to a stepwise rise or fall in [Ca2þ] after each exposure; the
results can be fitted to models of the chelators and their photoproducts, using their
aYnities and the relative quantum eYciencies of free and bound chelators (Fryer
and Zucker, 1993; Lando and Zucker, 1989). The percentage photolysis of the
chelator in response to each light exposure is the only free parameter, and is varied
until the model fits the results. In the case of the high-aYnity DM-nitrophen, little
rise in [Ca2þ] will occur until the total amount of remaining unphotolyzed chelator
equals the total amount of Ca2þ in the solution, whereupon the [Ca2þ] will increasesuddenly. Equations relating initial and final concentrations of DM-nitrophen,
48 Robert Zucker
total [Ca2þ], and photolysis rate (Zucker, 1993) then may be used to calculate
photolysis rate per flash or per second of steady light exposure.
Most photosensitive compounds also undergo substantial absorbance changes
after photolysis. These changes can be monitored during repeated exposure to the
light source without a Ca2þ indicator; the number of flashes or the duration of light
exposure required to reach a given percentage photolysis then can be determined.
Realizing that photolysis proceeds exponentially to completion (Zucker, 1993),
these data can be used to determine the photolysis rate directly. Ideally, both
methods should be used to check for consistent results. A final method for
determining photolysis rate is using high pressure liquid chromatography
(HPLC) to separate and quantify parent chelators and photoproducts in the
reaction solution after partial photolysis (Walker, 1991).
VIII. Purity and Toxicity
When experiments do not work as planned, the first suspected source of error is
the integrity of the photolabile chelator. DiVerent procedures have proved most
useful for testing the diVerent classes of compounds. The nitr and diazo com-
pounds undergo large absorbance changes on binding calcium and photolysis.
A 100 mM solution (nominally) of the chelator is mixed with 50 mM Ca2þ in
100 mM chelexed HEPES solution (pH 7.2), and 0.3 ml is scanned in a 1-mm
pathlength spectrometer. Then 1 ml 1 MK2EGTA is added to bring the [Ca2þ] to 0,
and the sample is scanned again. Finally, 1 ml 5 M CaCl2 is added to provide excess
Ca2þ, and a third scan is recorded. The first scan should be midway between the
other two. If the first scan is closer to the excess Ca2þ scan, it is indicative of a lower
than expected concentration of the chelator, probably because of an impurity.
Alternatively, Ca2þ may have been present with the chelator, which may be
checked by running a scan on the chelator with no added Ca2þ and comparing
the result with a scan with added EGTA; they should be identical. Ca2þ free and
Ca2þ-saturated chelator solutions also are scanned before and after exposure to
UV light suYcient to cause complete photolysis; the spectra are compared with
published figures (Adams et al., 1988, 1989; Kaplan and Ellis-Davies, 1988) to
determine whether the sample was partially photolyzed at the outset. The Ca2þ
aYnities of unphotolyzed and photolyzed chelators can be checked by measuring
the [Ca2þ] of 50%-loaded chelators with a Ca2þ-selective electrode.The absorbance of DM-nitrophen and some related chelators is almost Ca2þ
independent, so these procedures are not eVective. A solution of DM-nitrophen
nominally of 2 mM concentration is titrated with concentrated CaCl2 until the
[Ca2þ] measured with an ion-selective electrode suddenly increases; this change
indicates the actual concentration of the chelator and gives an estimate of purity.
The aYnity of the photolysis products can be measured as for the other chelators;
spectra before and after photolysis indicate whether the sample was already
partially photolyzed.
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 49
Purities of 80–90% are typical for commercial samples of all the chelators, but
occasional batches of 60% purity or less have been seen; these also sometimes show
high degrees of toxicity. Whether such low purity is the result of poor synthesis or
storage is unclear. Nitr compounds decompose detectably after only 1 day at room
temperature, and exposure to ambient fluorescent lighting for 1 day causes detect-
able photolysis. Chelators should be shipped on dry ice and stored at�80 �C in the
dark; even under these conditions they do not last forever. Repeated thawing and
freezing also degrades the compounds.
Some of the photolabile Ca2þ chelators display a degree of biological toxicity in
some preparations. Commercial samples of nitr-5 have been seen to lyse sea urchin
eggs (R. S. Zucker and L. F. JaVe, unpublished results) and leech blastomeres
(K. R. Delaney and B. Nelson, unpublished results) within minutes. Zucker and
Haydon (1988) found that nitr-5 blocked transmitter release within 10 min of
perfusion in snail neurons, whereas DM-nitrophen has no similar eVect(P. Haydon, unpublished results). These eVects are not caused by the photoproducts,since photolysis is not necessary for the problems to occur. DM-nitrophen has been
observed to reduce secretion in chromaYn cells; higher chelator concentrations,
photolyzed to give the same final [Ca2þ]i level, caused less secretion (C. Heinemann
and E.Neher, unpublished results). The eVect was overcome partially by inclusion of
glutathione in the perfusion solution, as reported for the photoproducts of other
2-nitrobenzhydrol-based caged compounds (Kaplan et al., 1978). These signs
of toxicity have been observed sporadically; whether they are properties of the
chelators themselves or of impurities in the samples used is unclear. The chelators
have been applied successfully to a wide range of preparations without obvious
deleterious results, although subtle eVects may have been missed.
IX. Biological Applications
A brief synopsis of the earliest biological applications of the caged Ca2þ chela-
tors follows along with a much more selective sampling of the more recent and
extensive literature. This is included in this chapter because many of the original
papers include a wealth of detail about methodology and interpretation of Ca2þ
photorelease technology.
A. Ion Channel Modulation
1. Potassium and Nonspecific Cation Channels
The first and still one of the major applications of photosensitive Ca2þ chelators
is analysis of Ca2þ-dependent ion channels in excitable cells. In 1987, Gurney et al.
first used nitr-2,-5, and-7 to activate Ca2þ-dependent Kþ current in rat sympathetic
neurons. These researchers found that a single Ca2þ ion binds to the channel with
rapid kinetics and 350 nM aYnity.
50 Robert Zucker
The next application of the nitr chelators was in an analysis of Ca2þ-activatedcurrents in Aplysia neurons (Lando and Zucker, 1989). We found that Ca2þ-activated Kþ and nonspecific cation currents in bursting neurons were linearly
dependent on [Ca2þ]i jumps in the micromolar range, as measured by arsenazo
spectrophotometry and modeling studies. Both currents relaxed at similar rates
after photolysis of nitr-5 or nitr-7, reflecting diVusional equilibration of [Ca2þ]inear the front membrane surface facing the light source. Potassium current relaxed
more quickly than nonspecific cation current, after activation by Ca2þ entry during
a depolarizing pulse, because of the additional voltage sensitivity of the Kþ
channels. This diVerence was responsible for the more rapid decay of hyperpolar-
izing afterpotentials than of depolarizing afterpotentials.
The role of Ca2þ-activated Kþ current in shaping plateau potentials in gastric
smooth muscle was explored by Carl et al. (1990). In fibers loaded with nitr-5/AM,
Ca2þ photorelease accelerated repolarization during plateau potentials and
delayed the time to subsequent plateau potentials, suggesting a role for changes
in [Ca2þ]i and Ca2þ-activated Kþ current in slow wave generation.
Another current modulated by [Ca2þ]i is the so-called M current, a muscarine-
blocked Kþ current in frog sympathetic neurons. Although inhibition is mediated
by G-protein coupling of the receptor to phospholipase C, resting M current is
enhanced by modest elevation of [Ca2þ]i (some tens of nanomolar) and reduced by
greater elevation of [Ca2þ]i, which also suppresses the response to muscarine
(Marrion et al., 1991). As for ventricular ICa (see below), several sites of modula-
tion of M current by [Ca2þ]i apparently exist. In these experiments, [Ca2þ]i waselevated by photorelease from nitr-5 and simultaneously measured with fura-2.
Step changes in [Ca2þ]i imposed by diazo-2 photolysis and monitored with bis-
fura-2 fluorescence changes have also been used to characterize the modulation of
cGMP-gated ion channels by [Ca2þ]i (Rebrik et al., 2000).
The after-hyperpolarization that follows spikes in rat hippocampal pyramidal
neurons is caused by a class of Ca2þ-dependent Kþ channels called IAHP channels.
This after-hyperpolarization and the current underlying it rise slowly to a peak 0.5 s
after the endof abriefburstof spikes.Ca2þphotorelease fromnitr-5 orDM-nitrophen
activates this currentwithout delay (Lancaster andZucker, 1994), and the currentmay
be terminated rapidly by photolysis of diazo-4 (but see conflicting results of Sah and
Clements, 1999), suggesting that the delay in its activation following action potentials
is caused by a diVusion delay between points of Ca2þ entry and the IAHP channels.
The Ca2þ sensitivity of the mechanoelectrical transduction current in chick
cochlear hair cells was studied using nitr-5 introduced by hydrolysis of the AM
form (Kimitsuki and Ohmori, 1992). Elevation of [Ca2þ]i to 0.5 mM (measured
with fluo-3) diminished responses to displacement of the hair bundle, and acceler-
ated adaptation during displacement when Ca2þ entry occurred. Preventing Ca2þ
influx blocked adaptation. Evidently, adaptation of this current was the result of
an action of Ca2þ ions entering through the transduction channels.
In guinea pig hepatocytes, noradrenaline evokes a rise in Kþ conductance after a
seconds-long delay. Photorelease of Ca2þ from nitr-5 and use of caged inositol
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 51
1,4,5-trisphosphate (caged-IP3) show that this delay arises from steps prior to or
during generation of IP3 (Ogden et al., 1990), which releases Ca2þ from intracellu-
lar stores to activate Kþ current.
2. Calcium Channels
The first application of DM-nitrophen was in a study of Ca2þ channels in chick
dorsal root ganglion neurons (Morad et al., 1988). With divalent charge carriers,
inactivation by photorelease of intracellular Ca2þ occurred within 7 ms, whereas
with monovalent charge carriers a nearly instantaneous block occurred, especially
when Ca2þ was released extracellularly. A similar rapid block of monovalent
current through Ca2þ channels was observed in response to photorelease of
extracellular Ca2þ in frog ventricular cells (Nabauer et al., 1989). DiVerent Ca2þ-binding sites may be exposed if altered conformational states are induced in the
channels by the presence of diVerent permeant ions.
The regulation of Ca2þ current (ICa) in frog atrial cells by [Ca2þ]i also has been
studied with nitr-5 (Charnet et al., 1991; Gurney et al., 1989). Rapid elevation of
[Ca2þ]i potentiated high-voltage-activated or L-type ICa and slowed its deactiva-
tion rate when Ba2þ was the charge carrier, after a delay of several seconds.
Inclusion of BAPTA in the patch pipette solution blocked the eVect of nitr-5
photolysis. The similarity of eVect of Ca2þ and cAMP and their mutual occlusion
suggest a common phosphorylation mechanism.
Regulation of ICa in guinea pig ventricular cells appears to be more complex
(Bates and Gurney, 1993; Hadley and Lederer, 1991). A fast phase of inactivation
reflects a direct action on Ca2þ channel permeation, since ICa inactivation caused
by photorelease of Ca2þ from nitr-5 is independent of the phosphorylation state of
the channels and does not alter gating currents. A late potentiation is also present,
the magnitude of which depends on the flash intensity delivered during a depolar-
izing pulse, but not on the initial [Ca2þ]i level, the degree of loading of nitr-5, or thepresence of BAPTA in the patch pipette. This result suggests that, during a
depolarization, nitr-5 becomes locally loaded by Ca2þ entering through Ca2þ
channels, and that the Ca2þ-binding site regulating potentiation is near the channel
mouth. Larger [Ca2þ]i jumps elicited by photolysis of DM-nitrophen evoke greater
ICa inactivation, but no potentiation, perhaps because of the more transient rise in
[Ca2þ]i when DM-nitrophen is photolyzed.
DM-nitrophen loaded with magnesium in the absence of Ca2þwas used to study
the Mg2þ-nucleotide regulation of L-type ICa in guinea pig cardiac cells (Backx
et al., 1991; O’Rourke et al., 1992). In the presence of ATP, a rise in [Mg2þ]i to50–200 mM led to a near doubling of the magnitude of ICa. Release of caged ATP
also increased ICa. Therefore, the eVect on Ca2þ channels was caused by a rise in
Mg2þ-ATP. Nonhydrolyzable ATP analogs worked as well as ATP, soMg2þ-ATP
seems to modulate Ca2þ channels directly.
We microinjected Aplysia neurons with nitr-5, DM-nitrophen, or diazo-4 to
characterize Ca2þ-dependent inactivation of Ca2þ current (Fryer and Zucker,
52 Robert Zucker
1993). Elevation of [Ca2þ]i to a few micromolar with nitr-5 caused little inactiva-
tion, but photolysis of DM-nitrophen rapidly inactivated half the ICa, presumably
that in the half of the cell facing the light source. Thus, inactivation requires high
[Ca2þ]i levels and occurs rapidly in all channels, even if they are closed. Experi-
ments with diazo-4 showed that an increase in buVering power reduced the rate of
inactivation of ICa modestly. DiVusion-buVer reaction simulations suggest that
Ca2þ acts at a site within 25 nm of the channel mouth (see also Johnson and
Byerly, 1993).
B. Muscle Contraction
One of the earliest applications of photolabile Ca2þ chelators was initiating
muscle contraction in frog cardiac ventricular cells by photorelease of extracellular
Ca2þ from DM-nitrophen (Nabauer et al., 1989). The strength of contraction
elicited by a stepwise rise in [Ca2þ]e showed a membrane potential dependence
that was indicative of entry through voltage-dependent Ca2þ channels rather than
of transport by Naþ–Ca2þ exchange.
Several laboratories have used caged Ca2þ chelators to study Ca2þdependentCa2þ release from the sarcoplasmic reticulum in rat ventricular myocytes.
Valdeolmillos et al. (1989) loaded cells with the AM form of nitr-5, Kentish et al.
(1990) subjected saponin-skinned fibers to solutions containing Ca2þ-loaded nitr-5,and Nabauer and Morad (1990) perfused single myocytes with DM-nitrophen
loaded with Ca2þ. Photolysis elicited a contraction blocked by ryanodine or
caVeine, procedures that prevent release of Ca2þ from the sarcoplasmic reticulum,
implicating Ca2þ-induced Ca2þ release, which could be confined to a portion of a
fiber by localized photolysis (O’Neill et al., 1990).
Gyorke and Fill (1993) used Ca2þ-DM-nitrophen to show that the cardiac
ryanodine receptors adapt to maintained [Ca2þ]i elevation, remaining sensitive to
larger [Ca2þ]i changes and responding by releasing still more Ca2þ. In smooth
muscle from guinea pig portal vein, the IP3-dependent release of Ca2þ was itself
dependent upon [Ca2þ]i (Iino and Endo, 1992). Ca2þ photoreleased from DM-
nitrophen and measured with fluo-3 accelerated Ca2þ release from a ryanodine-
insensitive, IP3-activated store. The possibility that adaptation reflected slow
unbinding of Ca2þ from the channels following a flash-induced Ca2þ ‘‘spike’’
was refuted by demonstrating a rapid deactivation of channel function to a sudden
drop in [Ca2þ]i imposed by diazo-2 (Velez et al., 1997).
Ca2þ-loaded nitr-5 was used in skinned frog and scallop muscle fibers to show
that the rate-limiting step in contraction is not the time-course of the rise in [Ca2þ]ibut rather the response time of the contractile machinery (Ashley et al., 1991b; Lea
and Ashley, 1990). Using isolated myofibrillar bundles from barnacle muscle, Lea
and Ashley (1990) showed that nitr-5 photolysis elevating [Ca2þ]i by 0.2–1.0 mMCa2þ not only activated contraction directly and rapidly but also evoked a slower
phase of contraction that was dependent on Ca2þ-induced Ca2þ release from the
sarcoplasmic reticulum. Analysis of [Ca2þ]i steps imposed by DM-nitrophen or
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 53
diazo-2 revealed kinetics of muscle contraction and relaxation steps following the
binding and unbinding of Ca2þ to troponin C (Ashley et al., 1993).
The first biological application of the caged chelator diazo-2 was in the study of
muscle relaxation. Mulligan and Ashley (1989) showed that rapid reduction in
[Ca2þ]i in skinned frog semitendinosus muscle resulted in a relaxation similar to
that occurring normally in intact muscle, indicating that mechanochemical events
subsequent to the fall in [Ca2þ] were rate limiting. However, Lannergren and Arner
(1992) reported some speeding of isometric relaxation after photolysis of diazo-2,
loaded in the AM form into frog lumbrical fibers. Lowered pH slowed relaxation
to a step reduction in [Ca2þ]i (Palmer et al., 1991), perhaps accounting for a
contribution of low pH to the sluggish relaxation of fatigued muscle. In contrast
to frog muscle, photorelease of Ca2þ chelator caused a much faster relaxation in
skinned scallop muscle than in intact fibers (Palmer et al., 1990), suggesting that, in
these cells, relaxation is rate limited primarily by [Ca2þ]i homeostatic processes.
C. Synaptic Function
Action potentials evoke transmitter release in neurons by admitting Ca2þ
through Ca2þ channels. Because of the usual coupling between depolarization
and Ca2þ entry, assessing the possibility of an additional direct action of mem-
brane potential on the secretory apparatus has been diYcult. Photolytic release of
presynaptic Ca2þ by nitr-5 perfused into presynaptic snail neurons cultured in
Ca2þ-free media was combined with voltage clamp of the presynaptic membrane
potential to distinguish the roles of [Ca2þ]i and potential in neurosecretion (Zucker
and Haydon, 1988), revealing no direct eVect of membrane potential on transmit-
ter release.
Hochner et al. (1989) injected Ca2þ-loaded nitr-5 into crayfish motor neuron
preterminal axons, and used a low-[Ca2þ] medium to block normal synaptic
transmission. They found that action potentials transiently accelerated transmitter
release evoked by modest photolysis of nitr-5. However, Mulkey and Zucker
(1991) used fura-2 to show that the extracellular solutions used by Hochner et al.
(1989) failed to block Ca2þ influx through voltage-dependent Ca2þ channels.
When external Ca2þ chelators or channel blockers eliminated influx completely,
spikes failed to have any influence on transmitter release, even when it was
activated strongly by photolysis of intracellularly injected Ca2þ-loaded DM-
nitrophen.
Delaney and Zucker (1990) confirmed at the squid giant synapse that in a Ca2þ-free medium, action potentials have no eVect on transmitter release triggered by a
rise in [Ca2þ]i upon photolysis of presynaptically injected DM-nitrophen. Flash
photolysis of DM-nitrophen produced a transient postsynaptic response resem-
bling normal excitatory postsynaptic potentials. The intense phase of transmitter
release was probably caused by the brief spike in [Ca2þ]i following partial photol-
ysis of partially Ca2þ-loaded DM-nitrophen. This response began a fraction of a
millisecond after the rise in [Ca2þ]i, a delay similar to the usual synaptic delay
54 Robert Zucker
following Ca2þ influx during an action potential; both delays had the same
temperature dependence. Thus photolysis of DM-nitrophen caused a [Ca2þ]i tran-sient resembling that occurring normally at transmitter release sites in the vicinity
of Ca2þ channels that open briefly during an action potential. After the secretory
burst, a moderate phase of transmitter release persisted for 15 ms, corresponding
to a relaxation in [Ca2þ]i measured with fura-2 that probably reflected slow Ca2þ
displacement of Mg2þ bound to unphotolyzed DM-nitrophen.
Similar responses to partial flash photolysis of lightly Ca2þ-loaded DM-nitro-
phen were observed at crayfish neuromuscular junctions (Lando and Zucker,
1994). Transmitter release evoked by slow photolysis of Ca2þ-DM-nitrophen
using steady illumination also has been studied at this junction (Mulkey and
Zucker, 1993). The rate of quantal transmitter release, measured as the frequency
of miniature excitatory junctional potentials (MEJPs), was increased �1000-fold
during the illumination. Brief illumination (0.3–2 s) evoked a rise in MEJP fre-
quency that dropped abruptly back to normal when the light was extinguished, as
would be expected from the reversible rise in [Ca2þ]i that should be evoked by such
illumination, which leaves most of the DM-nitrophen unphotolyzed (Zucker,
1993). Longer light exposures caused an increase inMEJP frequency that outlasted
the light signal, as would be expected from the rise in resting [Ca2þ]i after photoly-sis of most DM-nitrophen. These experiments illustrate the utility of steady pho-
tolysis of partially Ca2þ-loaded DM-nitrophen in generating reversible changes in
[Ca2þ]i in cells.
At cultured snail synapses, FMRFamide inhibits asynchronous transmitter
release elicited by [Ca2þ]i elevated by photolysis of presynaptic nitr-5 (Man-Son-
Hing et al., 1989), and blocks synchronous release to partial flash photolysis of
partially Ca2þ-loaded DM-nitrophen (Haydon et al., 1991). As at crayfish and
squid synapses, these flash-evoked postsynaptic responses resembled the spike-
evoked responses and were triggered by the spike in [Ca2þ]i that results when DM-
nitrophen is used in this fashion.
At leech serotonergic synapses, a presynaptic Ca2þ uptake process may be
activated by photolysis of presynaptic DM-nitrophen; blocking it with zimelidine
or by external Naþ removal eliminated the presynaptic transport current and
prolonged the postsynaptic response, uncovering a contribution of this process
to the termination of transmitter release (Bruns et al., 1993).
DM-nitrophen has been used extensively to probe the steps involved in exocyto-
sis in endocrine cells. Measuring [Ca2þ]i changes with furaptra, we and others
(Heinemann et al., 1994; Neher and Zucker, 1993; Thomas et al., 1993) observed—
in bovine chromaYn cells and rat melanotrophs—three kinetic secretory phases in
response to [Ca2þ]i steps to �100 mM, reflecting release from diVerent vesicle
pools. Prior exposure to a modest [Ca2þ]i rise primed phasic responses to a
subsequent step in [Ca2þ]i, indicating that [Ca2þ]i not only triggers exocytosis but
also mobilizes vesicles into a docked or releasable status. After exocytosis, another
[Ca2þ]i stimulus often evoked a rapid reduction in membrane capacitance signaling
a [Ca2þ]i-dependent compensatory endocytosis.
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 55
The release of synaptic transmitter by action potentials is often enhanced for about
a second after short bouts of presynaptic activity (synaptic facilitation), and formuch
longer after sustained activity lasting minutes (posttetanic potentiation, PTP). We
have used photolytic release of presynaptic Ca2þ from DM-nitrophen to induce
facilitation without electrical activity (Kamiya and Zucker, 1994). Photolysis of
diazo-2 or diazo-4 to terminate the [Ca2þ]i increase lingering briefly after a short
spike train abolished facilitation immediately at crayfish neuromuscular junctions.
PTP induced by longer stimulation, both in this preparation and inAplysia neuronal
synapses (Fischer et al., 1997), was abolished more slowly by rapid reduction of the
prolonged residual [Ca2þ]i resulting frommitochondrial overload (Tang and Zucker,
1997). Thus, facilitation and PTP arise from residual [Ca2þ]i acting on distinct
molecular targets diVerent from the secretory trigger, which is also activated by
Ca2þ. We compared responses to [Ca2þ]i steps on flash photolyzing DMNPE-4 at
weakly transmitting but strongly facilitating neuromuscular junctions to responses at
strongly transmitting but depressible junctions (Millar et al., 2005), and concluded
that the diVerence in response kinetics was best explained by a diVerence in the state
of Ca2þ-dependent priming, such that strongly transmitting synapses were already
preprimed at rest by a priming target tuned to have a higher Ca2þ-sensitivity. Thisled, in turn, to the development of a comprehensive model of synaptic transmission,
facilitation, and depression that comprised three Ca2þ-dependent processes—vesicle
mobilization to docking sites, priming of docked vesicles, and activation of mem-
brane fusion (Pan and Zucker, 2009).
We (Lando and Zucker, 1989) and Heidelberger et al. (1994) were the first to use
DM-nitrophen photolysis to characterize the Ca2þ-cooperativity of secretion at
neuromuscular junctions and retinal bipolar neurons; subsequently, we (Ohnuma
et al., 2001) used NP-EGTA and Kasai et al. (1999) used DM-nitrophen to show
diVerences in the Ca2þ-dependence and sensitivity of peptidergic or aminergic
large dense core vesicle fusion and cholinergic small clear vesicle fusion at central
molluscan synapses and in PC12 cells. Hsu et al. (1996) reported that transmitter
release at squid giant synapses decayed to step [Ca2þ]i increases produced by
NP-EGTA photolysis; subsequent higher steps evoked more release, indicating
transmitter stores had not been depleted, suggesting either an adaptation of the
release process, as the authors proposed, or possibly vesicle heterogeneity in
sensitivity to release, or the operation of mobilization or priming processes
enabling release of previously undocked or unprimed vesicles at higher [Ca2þ]i.Caged Ca2þ photolysis has been used extensively in the last decade in many
elegant experiments, especially from the laboratories of Erwin Neher and Bert
Sakmann, to kinetically characterize in detail the secretory trigger for neurosecre-
tion, primarily at the giant synapse of the calyx of Held (Bollmann and Sakmann,
2005; Bollmann et al., 2000; Felmy et al., 2003a,b; Hosoi et al., 2007; Sakaba et al.,
2005; Schneggenburger and Neher, 2000; Wadel et al., 2007; Wang et al., 2004;
Young and Neher, 2009). Ca2þ uncaging from DM-nitrophen has been used to
probe the kinetics and cooperativity of Ca2þ binding to the secretory trigger,
kinetic consequences of SNARE protein and synaptotagmin mutation, eVects of
56 Robert Zucker
Ca2þ on synaptic facilitation, the dependence of release kinetics on the time-course
of local [Ca2þ]i changes, heterogeneity of vesicle Ca2þ-sensitivity and release
kinetics, the role of Ca2þ in mobilizing vesicles to replenished pools depleted in
synaptic depression, and to compare secretion evoked by global [Ca2þ]i manipula-
tion in uncaging to local [Ca2þ]i influx though voltage-dependent channels to
address the question of distance of the secretory target from Ca2þ channels and
its changes in development. Kinetic studies of the Ca2þ-dependence of secretion
using DM-nitrophen have also been conduced on cochlear hair cells (Beutner et al.,
2001) and photoreceptors (Duncan et al., 2010).
Zoran et al. (1991) used Ca2þ photorelease to study synapse maturation. Spike-
evoked transmitter release begins only several hours after cultured snail neurons
contact a postsynaptic target. DM-nitrophen photolysis showed this developmen-
tal change to result from the delayed increase in Ca2þ-sensitivity of the secretory
machinery.
Long-term potentiation and depression (LTP and LTD) in mammalian cortical
synapses are involved in cognitive processes such as memory consolidation and
spatial learning. We found that a brief but strong postsynaptic [Ca2þ]i elevationwas suYcient to induce LTP in rat hippocampal CA1 synapses onto the injected
neuron, while a more prolonged but modest [Ca2þ]i elevation specifically induced
LTD, and a brief but modest Ca2þ rise could elicit either (Malenka et al., 1988;
Neveu and Zucker, 1996a,b; Yang et al., 1999). By terminating the [Ca2þ]i risefollowing a brief aVerent tetanus by photoactivating the Ca2þ chelator diazo-4, we
showed that postsynaptic [Ca2þ]i must remain elevated for several seconds before it
can induce (Malenka et al., 1992). We also found that long-lasting changes in
synaptic transmission at CA3 hippocampal pyramidal cells can be produced by
postsynaptic [Ca2þ]i elevations induced by DM-nitrophen, NP-EGTA, or
DMNPE-4 photolysis (Wang et al., 2004).
A diVerent form of LTD in cerebellar Purkinje neurons that plays a role in
motor skill learning, parallel fiber synapses are depressed when their activity
coincides with postsynaptic firing, especially when the latter is triggered by climb-
ing fiber input. Lev-Ram et al. (1997) showed that photolytic release of caged Ca2þ
from nitr-7 could replace postsynaptic spiking and that photolytic release of either
caged NO or caged cGMP could replace parallel fiber activity; simultaneous
uncaging of Ca2þ and either NO or cGMP could induce LTD without any
electrical stimulation at all. Kasono and Hirano (1994) showed that a modest
release of Ca2þ from nitr-5 depressed responses to glutamate application to a
dendrite only when the stimuli were temporally paired. Using DMNPE-4,
Tanaka et al. (2007) found that a suYciently high and prolonged Ca2þ elevation
alone could induce LTD, and that the threshold for LTD induction was history-
dependent.
Depolarization-induced suppression of inhibition (DSI) is another form of
cortical synaptic plasticity, which is mediated by activation of postsynaptic endo-
cannabinoid synthesis by activity-induced [Ca2þ]i elevation and subsequent retro-
grade regulation of inhibitory transmitter release. We found identical DSI
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 57
sensitivities to uniform postsynaptic [Ca2þ]i elevation by NP-EGTA photolysis vs.
the volume-average of the highly nonuniform [Ca2þ]i elevation on opening voltage-
sensitive Ca2þ channels by depolarizations (Wang and Zucker, 2001), implying
that the enzymatic targets of postsynaptic Ca2þ entering through Ca2þ channels in
activating DSI are not tightly colocalized with the channels—a situation exactly
opposite of the case for Ca2þ activation of classical transmitter release.
Long-lasting synaptic regulation also occurs at developing neuromuscular junc-
tions, where repeated activation of one of two motor neuron inputs results in a
postsynaptic Ca2þ-dependent compensatory or homeostatic reduction in presyn-
aptic transmitter release to action potentials at terminals facing the activated
receptors. Using focal DM-nitrophen or nitr-5 photolysis to mimic the localized
postsynaptic [Ca2þ]i elevation seen to accompany the stimulus normally used to
induce this selective persistent depression, we were able to induce a similar syn-
apse-specific modification (Cash et al., 1996a). Subsequently, synapses made by the
modified motor neuron onto other muscle fibers also became depressed by the
spread of an unidentified presynaptic intracellular signal (Cash et al., 1996b).
D. Other Applications
The tight regulation of cytoplasmic [Ca2þ]i is essential for ensuring that Ca2þ can
act reliably and eYciently as a localized second messenger of a huge variety of
cellular processes. Endogenous buVers play a defining role in this process, and an
appreciation of the functions of these buVers and their characteristics (aYnities,
binding kinetics, mobility, and localization) is crucial to our understanding how
Ca2þ performs its central cellular functions. Use of photosensitive Ca2þ chelators
has become an important tool in the estimation of cytoplasmic buVer characteristics,and much eVort has gone into developing procedures and protocols for defining
them with some precision. Some of the best examples of this sort of analysis come
from the laboratories of Stephen Bolsover, Istvan Mody and Julio Vergara, and
ErwinNeher, whose papers should be consulted for the analytical details (Faas et al.,
2007; Fleet et al., 1998; Nagerl et al., 2000; Naraghi et al., 1998; Xu et al., 1997).
In addition to these major areas of application of caged Ca2þ chelators, this
method of [Ca2þ]i manipulation has been used to address an increasingly diverse
range of biological problems. Nitr and diazo compounds were inserted by AM
loading into fibroblasts that were activated by mitogenic stimulation to produce
[Ca2þ]i oscillations monitored using f1uo-3 (Harootunian et al., 1988). Photore-
lease of Ca2þ from nitr-5 enhanced and accelerated the oscillations, whereas
release of caged chelator by photolysis of diazo-2 inhibited them. Nitr-7 photolysis
caused not only an immediate rise in [Ca2þ]i liberated from the photolyzed chela-
tor, but also elicited a later rise in [Ca2þ]i (Harootunian et al., 1991). This eVect wasshown, pharmacologically, to be caused by IP3-sensitive stores, suggesting that an
interaction between [Ca2þ]i and these stores underlies the [Ca2þ]i oscillations.Photorelease of Ca2þ from DM-nitrophen has been used to study the binding
kinetics of Ca2þ to the Ca2þ-ATPase of sarcoplasmic reticulum vesicles (DeLong
58 Robert Zucker
et al., 1990). The relaxation of the [Ca2þ] step, measured by arsenazo spectropho-
tometry after photolysis, revealed the kinetics of binding to the ATPase. Changes
in the Fourier transform infrared spectrum consequent to photorelease of Ca2þ
from nitr-5 provided information on structural changes in the ATPase after
binding Ca2þ (Buchet et al., 1991, 1992). In a final application to the study of
enzyme conformational changes, photolysis of Mg2þ-loaded DM-nitrophen was
used to form Mg2þ-ATP rapidly to activate Naþ/Kþ exchange, the state of which
was monitored by fluorescence of aminostyrylpyridinium dyes (Forbush and
Klodos, 1991). Rate-limiting steps were measured at 45 s�1 by this method.
Ca2þ has been implicated in the control of filopodial activity in the responses of
growth cones of developing neurons to environmental cues. Pioneer neurons lay
out peripheral aVerent pathways in developing grasshoppers. We loaded pioneer
neurons by de-esterification of the AM esters of DM-nitrophen and calcium green
(Lau et al., 1999) and showed that elevation of local [Ca2þ]i in a growth cone to
� 1 mM for just 10 s was suYcient to activate subsequent filopodial prolongation
and induce the formation of new filopodia at spots with high actin concentration
(labeled with rhodamine-phalloidin).
In other applications, Gilroy et al. (1991) and Fricker et al. (1991) microinjected
Ca2þ-loaded nitr-5 into guard cells of lily leaves and showed that photorelease of
about 600 nM intracellular Ca2þ (measured with fluo-3) initiated stomatal pore
closure. Kao et al. (1990) loaded Swiss 3T3 fibroblasts with nitr-5/AM and showed
that photolysis that elevated [Ca2þ]i by hundreds of nanomolar (measured by
fluo-3) triggered nuclear envelope breakdown, an early step in mitosis, while
having little eVect on the metaphase to anaphase transition. Control experiments
using nitr-9 showed no eVect of reactive photochemical intermediates or products.
Groigno and Whitaker (1998) initiated chromosome disjunction and segregation
in embryonic sea urchin cells by Ca2þ photorelease from NP-EGTA or by photol-
ysis of caged IP3. Ca2þ buffers prevented chromatid separation but not the later
stages of anaphase, indicating a specific role for Ca2þ in early anaphase chromo-
some disjunction. Tisa and Adler (1992) used electroporation to introduce Ca2þ-loaded nitr-5 or DM-nitrophen into Escherichia coli bacteria, and showed that
elevation of [Ca2þ]i enhanced tumbling behavior characteristic of chemotaxis
whereas photorelease of caged chelator from diazo-2 decreased tumbling. Photo-
lysis of diazo-3, which reduces pH without aVecting [Ca2þ]i, caused only a small
increase in tumbling. Mutants with methyl-accepting chemotaxis receptor proteins
still responded to Ca2þ, whereas mutants of specific Che proteins did not, indicat-
ing that the action of these proteins lay downstream of the Ca2þ signal.
X. Conclusions
Interest in photolabile Ca2þ chelators has been intense. Their range of applica-
tion has broadened well beyond the original nerve, muscle, and fibroblast prepara-
tions. They remain one of the most valuable tools for the precise definition of
2. Photorelease Techniques for Raising or Lowering Intracellular Ca2þ 59
calcium’s roles and mechanisms of action in cell biology. They have attracted and
challenged some of the best minds in physiology, resulting in great conceptual and
skillful sophistication in the rapid evolution of this technology, which shows little
sign of abating.
Acknowledgments
I thank Steve Adams for valuable discussion and Joseph Kao for drawings of chelator structures.
The research done in my laboratory in this area was supported primarily by National Institutes of
Health Grant NS 15114.
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CHAPTER 3
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
Making and Using Calcium-SelectiveMini- and Microelectrodes
L. Hove-Madsen,* S. Baudet,† and D. M. Bers‡
*Cardiovascular Research Centre CSIC-ICCCHospital de la Santa Creu i Sant PauBarcelona, Spain
†Ricerca Biosciences SASSaint Germain sur l’Arbresle, France
‡Department of PharmacologyUniversity of California, Davis,Davis, California, USA
A
OGY,All rig
bstract
VOL. 99 0091hts reserved. 67 DOI: 10.1016/S0091
-679X-679X
I. In
troduction A. Main Characteristics of Ca2þ-Selective ElectrodesII. R
ationale III. M ethodsA.
Preparation of Minielectrodes B. Application of Minielectrodes C. Preparation of Ca2þ-Selective MEs D. Application of Ca2þ-Selective MEsIV. D
iscussion R eferencesAbstract
Detection and measurement of intracellular calcium concentration ([Ca2þ]i)have relied on various methods, the popularity of which depends on their ease of
use and applicability to diVerent cell types. Historically, Ca2þ-selective electrodeshave been used concomitantly with absorption indicators such as arsenazo-III,
but their interest has been eclipsed by the introduction of a large number of
fluorescent calcium probes with calcium sensitivities varying from the nanomolar
/10 $35.00(10)99003-1
68 L. Hove-Madsen et al.
to the micromolar range such as fura-2, indo-1, fluo-4, and many others. In this
chapter, we emphasize the utility of Ca2þ-selective electrodes and show that their
use is complementary to use of fluorescent indicators; indeed, each method has
advantages and disadvantages. We first describe the preparation and application
of Ca2þ-selective minielectrodes based on the Ca2þ ligand ETH 129 (Schefer
et al., 1986) that have a larger dynamic range and faster response time than most
commercially available calcium electrodes. The second part of the chapter is
dedicated to ETH 129-based Ca2þ-selective microelectrodes (MEs), and their
application in the determination of [Ca2þ]i in cardiac cells. Since numerous
reviews and books have been dedicated to the theoretical aspects of ion-selective
ME principles and technology, this chapter is not intended for investigators who
have no experience with MEs.
I. Introduction
A. Main Characteristics of Ca2þ-Selective Electrodes
The key advantage of the Ca2þ-selective electrodes is the wide dynamic range
of their response (e.g., from pCa 9 to 1), as compared, for example, to
fluorescent and metallochromic Ca2þ indicators that typically have a dynamic
range of four or less pCa units (Fig. 1). There has been developed a plethora of
useful fluorescent calcium probes with calcium sensitivities varying from the
nanomolar to the micromolar range such as fura-2, indo-1, fura red, fluo-4,
furaptra, fluo-5N, and others (Grynkiewicz et al., 1985; Harkins et al., 1993;
Lipp et al., 1996; Picht et al., 2006; Shannon and Bers, 1997). These are widely
used and are extremely important tools for study of Ca2þ, but Ca2þ-selectiveelectrodes are a valuable complementary tool. For more basic reference to
electrode technology and electrophysiology, we suggest monographs by
Ammann (1986), Purves (1981), and Thomas (1982). An electronic introduc-
tion to ion-selective electrodes can be found at www.nico2000.net/Book/
Guide1.html.
Their response is based on a semiempirical equation (Nicolski–Eisenman equa-
tion) derived from the Nernst equation:
Ex ¼ Eo þ RT=ZxF lnðax þKpotxy azx=zyy Þ ð1Þ
where Ex is the ion-selective electrode potential, Eo is a constant, R, T, Z, and F
have their usual meaning, ax is the activity of the ion that is measured (activity
(a) is related to concentration (C) by the relation: a¼gC where g is the activity
coeYcient) and Kxypot is the selectivity coeYcient. This expression is strictly
valid for activities only, but if the activity coeYcients do not change, they can
be used with free concentrations too. This is often for convenience, since
solutions and chemical equilibria are more often described in these concentra-
tions terms. So, if x is Ca2þ and y is Naþ (the most common interfering cation
100
Indo-1
Minielectrode
Microelectrode
pCa
8
Pot
entia
l (m
V)
75
50
25
0
−25
−50
−757 6 5 4 3
Flu
ores
cenc
e ra
tio
3
2
1
0
Fig. 1 Dynamic range of Ca2þ-selective electrodes and indo-1. The electrode potential of Ca2þ-selective mini and MEs is shown together with the fluorescence ratio (400/470) for indo-1. Measure-
ments were performed in a KCl buVer containing 140 mM KCl, 10 mM HEPES, 10 mM NaCl, and
1 mM EGTA. Notice that indo-1 is suitable for measurements between pCa 7.5 and 5, while the
dynamic range for the Ca2þ electrode is wider, ranging from pCa 9 to 1 for minielectrodes and 7.5 to
1 for MEs.
3. Calcium Selective Mini- and Microelectrodes 69
for the Ca2þ-selective ligand), the relationship becomes, for 30 �C and changing
to log 10:
ECa ¼ Eo þ 30 log Ca2þ� �þKpot
NaCa Naþ½ �2� �
ð2Þ
Thus for the case of no interfering ion and when extracellular Ca2þ¼ [Ca2þ]ref,then the potential diVerence (DE) between two solutions of diVerent [Ca2þ] reducesto the Nernst equation:
DE ¼ 30 logð½Ca2þ�=½Ca2þ�refÞ ð3ÞThe response of an ion-selective electrode to changes in free [Ca2þ] is much slower
than the fluorescent, bioluminescent, and metallochromic Ca2þ indicators. Thus,
Ca2þ electrodes are ideal for measurements of slow changes over wide ranges of
[Ca2þ], but not appropriate for very rapid changes of free [Ca2þ] although the Ca2þ
electrodes can respond in the millisecond range at higher [Ca2þ] (Bers, 1983). Theresponse time can also be improved by using Ca2þ selective electrodes with a
concentric inner micropipette that reduces the longitudinal resistance of the
Ca2þ-selective resin and thereby decreases the electrical time constant (Fedirko
et al., 2006; Ujec et al., 1979). These limitations, together with the physical size of
Ca2þ-selective electrodes put them at disadvantage with fluorescent and lumines-
cent calcium indicators for measurements of dynamic changes in calcium levels in
70 L. Hove-Madsen et al.
cellular environments. Moreover, about the time that indo-1 and fura-2 were
synthesized (Grynkiewicz et al., 1985), a Ca2þ-ligand of improved selectivity
(ETH 129) was introduced (Schefer et al., 1986), which made it more realistic to
measure low Ca2þ levels typical of intracellular environments, since the electrode
response was Nernstian until pCa 8–9 (depending on the ionic background).
However, the availability and popularity of the fluorescent indicators did to a
large extent eclipse and limit further use and characterization of ETH 129-based
electrodes.
II. Rationale
We will not present an extensive review of the advantages and disadvantages of
Ca2þ-selective electrodes compared to fluorescent indicators, but the purpose of
this chapter is rather to show how the advantages of both approaches can be
combined for specific purposes. For example, a classical problem with Ca2þ
indicators is to convert the fluorescent signal to actual [Ca2þ]. In fact, it is
commonly acknowledged that in vitro calibration curves of fluorescent calcium
indicators are not applicable to in vivo or in situ conditions, because the indicators
bind to intracellular constituents, which modifies their excitation/emission spec-
trum and decreases the apparent aYnity of Ca2þ for the probe (Blatter and Wier,
1992; Harkins et al., 1993; Hove-Madsen and Bers, 1992; Konishi et al., 1988).
In vivo calibration curves are even more diYcult to obtain and are highly depen-
dent on the cell type under study. Ca2þ-selective electrodes are still one of the most
straightforward ways to measure and quantify Ca2þ because (1) their behavior is
not appreciably altered by the intracellular milieu (ions and proteins); (2) they can
be easily included in an electrophysiological setup and do not require extensive
and expensive apparatus; (3) their linear (Nernstian) response simplifies the con-
version of the voltage signal to [Ca2þ]i; (4) their behavior allows determination
of wide ranges of pCa (see above); and (5) the Ca2þ ligand itself does not change
(or buVer) [Ca2þ]i.Thus, Ca2þ-selective electrodes continue to be a good choice for the preparation
of calibration solutions for Ca2þ determinations and for measurements of dissoci-
ation constants for Ca2þ-binding compounds such as ethylene glycol bis(b-amino
ethyl ether)-N,N,N0,N0-tetraacetic acid (EGTA), indo-1, 1,2-bis(o-aminophenoxy)
ethane-N,N,N0,N0-tetraacetic acid (BAPTA), and oxalate under diVerent experi-mental conditions (Bers, 1982; Harrison and Bers, 1987, 1989; Hove-Madsen and
Bers, 1992, 1993a; Hove-Madsen et al., 1998) Indeed, commercially available Ca2þ
electrodes are largely directed towards determination of the calcium concentration
in solutions or biological fluids, but Ca2þ electrodes can also be used to measure
cellular Ca2þ buVering and changes in the free [Ca2þ] in cell suspensions (Hove-
Madsen and Bers, 1993a, 1993b; Hove-Madsen et al., 1998) or combined with
other electrophysiological techniques (Kang and Hilgemann, 2004; Kang et al.,
2003). Moreover, Ca2þ electrodes are economical, easy to prepare, and they can
3. Calcium Selective Mini- and Microelectrodes 71
easily be used for measurements of free [Ca2þ] in experimental solutions or
biological fluids. Indeed, for those studying Ca2þ-dependent processes, there is
no practical or economical reason why Ca2þ electrodes should not be used as
routinely as pH electrodes.
III. Methods
A. Preparation of Minielectrodes
Ca2þ-selective minielectrodes can be prepared by dipping polyethylene (PE)
tubes (typically �5 cm) in a membrane solution (see composition below). We
have tried other types of tubing but polyvinyl chloride (PVC) tubing appears to
absorb ETH 129 from the Ca2þ-selective membrane, resulting in a faster loss of
sensitivity as compared to the PE tubing. On the other hand, materials such as
Teflon tubing absorb little ETH 129 but the PVC membrane does not adhere well
to the tubing. As a result, the electrodes are more easily damaged, although they
may have a longer lifetime if handled with care.
The dimensions of the tubing vary from a diameter less than 1 to �3 mm.
Electrodes prepared with the membrane solution described below result in Ca2þ-selective membranes that are a few hundred micrometers thick. With diameters
larger than 5 mm, the Ca2þ-selective membrane bursts more easily during
handling, but this problem may be overcome by inserting a ceramic plug into the
tubing, before dipping it in the membrane solution as described by Orchard et al.
(1991). For general purposes we have used inner electrode diameters of 1.67 mm
(PE 240, Clay Adams).
After dipping the PE tubing in the membrane solution, the Ca2þ-selectivemembrane is allowed to dry overnight. Then the electrode is filled with an appro-
priate filling solution, which should correspond to experimental conditions (see
below). After filling the electrode, it is allowed to equilibrate for at least 3 days in a
glass vial containing the filling solution (but see below).
1. Preparation and Use of the Ca2þ-Selective Ligand
The Ca2þ-selective membrane can be prepared as described by Schefer et al.
(1986) (all from Fluka/Sigma-Aldrich; Table I).
ETH 129 is dissolved inN-phenyl-octyl-ether (NPOE) under vigorous stirring in
a small glass vial (Solution 1). At the same time, PVC is dissolved in the THF;
when completely dissolved, potassium TCPB is added (Solution 2). When the
components of Solutions 1 and 2 are completely dissolved, the two solutions are
mixed, and the membrane solution is ready to use, or can be stored in a glass vial,
closed with a Teflon screwcap and protected from light. If THF evaporates from
the membrane solution during storage, a small amount of THF can be added to
achieve the desired viscosity of the membrane solution.
Table IPreparation of the PVC-based Ca2þ-selective ligandminielectrodes
Component* Amount
Solution 1
ETH 129 25 mg
N-phenyl-octyl-ether (NPOE) 451.5 ml
Solution 2
Polyvinyl chloride (PVC) 250 mg
Kþ-tetrakis chlorophenyl borate (TCPB) 12.9 mg
Tetrahydrofuran (THF) �5 ml
*Components can be obtained from Sigma-Aldrich (St. Louis, MO)
72 L. Hove-Madsen et al.
The filling solution used for the minielectrode depends on the experimental
solutions. Generally, the ionic composition should mimic the environment in
which measurements of Ca2þ are planned, and the [Ca2þ] of the filling solution
should also be in the range of the measured values. With measurements of low
[Ca2þ], Ca2þ in the filling solution can be buVered with EGTA to the desired free
[Ca2þ]. However, we have not obtained good results with filling solutions with a
pCa higher than 7.5. We typically use a Ca2þ-EGTA buVer with 1 mM free Ca2þ as
a filling solution.
2. Electrode Characteristics
The resistance of the minielectrodes is 1–2 MO, which normally makes it possi-
ble to use a standard pH/ion meter to monitor the electrode potential. We have
used either a commercial pH/ion meter (Orion pH/ion analyzer) or preamplifier
(A311J, Analog Devices). When using commercial pH/ion meters, it is normally
necessary to make an adapter cable that connects the minielectrode to the meter
input. We have used a chart recorder (Soltec) or an electronic data acquisition
device (Linseis) for continuous monitoring of the electrode potential.
The lifetime of the minielectrode depends on the [Ca2þ] the electrode is used to
measure, and on the composition of the experimental solutions. For measurements
in solutions without protein or interfering ions, the detection limit for the electro-
des increases slowly with time and the electrodes will have a detection limit in the
subnanomolar range and a Nernstian response down to 10 nM for at least 1 month
after preparation. Figure 2 shows the change in the response of a Ca2þ electrode
with time.
Two days after filling, the electrode response was still ‘‘super-Nernstian’’ at low
[Ca2þ] (i.e., >29 mV per 10-fold change in [Ca2þ]), but normal within 7 days after
filling.Notice thatwedid still obtainaNernstian responsedowntopCa8 for2months
after filling the electrode. The response time at low [Ca2þ], however, got slower with
Pot
entia
l (m
V)
2 Months
2 Days
50
25
0
−25
−50
−75
−100
pCa
8 7 6 5 4
Fig. 2 Electrode potential of a Ca2þ-selective minielectrode 2, 7 and 60 days after filling of the
minielectrode.Measurements were performed in a KCl buVer as in Fig. 1. Notice the ‘‘super-Nernstian’’
response of electrodes 2 days after filling (circles). Seven days after the filling (squares), electrode
response was linear down to a free [Ca2þ] of less than 10 nM. The slope of the regression line was
�28.4 mV/pCa. Two months after filling (diamonds), the electrode response was linear to pCa 7.5, but
response time was slowed at high pCa.
3. Calcium Selective Mini- and Microelectrodes 73
time and measurements below 30 nM are only practical with fairly fresh electrodes.
We generally fill electrodes once a week to obtain the best results. However, if the
Ca2þ electrodes are used to measure micromolar or higher [Ca2þ] in protein-free
solutions, the same electrode can be used for longer periods (up to several months).
In the presence of cellular proteins, a small oVset in the electrode response is seen
at the first exposure to protein. Then, no further alteration of the electrode
response occurs, but the response time of the Ca2þ electrode is increased after
exposure to protein, and measurements of free [Ca2þ] below 10 nM are more
diYcult in the presence of protein concentrations higher than 10 mg/ml.
The response time of the Ca2þ electrodes can be of critical importance in some
applications. Figure 3 compares the response times of a Ca2þ electrode and indo-1
fluorescence to a decrease in the [Ca2þ] in a suspension of permeabilized myocytes
(3 mg/ml) where cellular Ca2þ uptake processes have been blocked with thapsigar-
gin and ruthenium red.
We examined the response to a decrease in [Ca2þ], as this may be a more
stringent test than an increase in [Ca2þ]. Notice that when 2 mM EGTA was
added to lower the free [Ca2þ], the electrode response was 94% complete in 1 s
while the indo-1 signal was 97% complete in 1 s. A slow final phase, lasting several
seconds, is apparent in the electrode signal only (see amplified inset).
In Fig. 3 the response time was examined under experimental conditions where
spatial inhomogeneities in the myocyte suspension are minimized by buVeringCa2þ with indo-1 and oxalate. However, under some experimental conditions, an
50Indo
Elec
ElecEGTA
40
30
20
Free
[Ca]
(mM
)
Free
[Ca]
(mM
)
10
−10 10 15−5 0
Time (s)
5
1.5
1
0.5
0
0 10Time (s)
0
Fig. 3 Comparison of response time of a Ca2þ-selective minielectrode and indo-1. Free [Ca2þ] wasmeasured in a suspension of permeabilized rabbit ventricular myocytes. Ca2þ uptake into sarcoplasmic
reticulum and mitochondria was inhibited with thapsigargin and ruthenium red, respectively. The initial
free [Ca2þ] was 32 mMwhich is near saturation for indo-1, resulting in a very noisy trace. That is because
a small change in fluorescence ratio corresponds to a large in [Ca2þ] at this level (see Fig. 1). At the
arrow, 2 mMEGTAwas added to the cell suspension to lower free [Ca2þ]. Both the electrode and indo-1signal were more than 90% complete in 1 s. The inset shows the response of indo-1 and Ca2þ electrode at
low [Ca2þ]. Notice that the indo-1 signal was 100% complete in 2 s (and actually undershoot slightly)
while the final completion of the electrode response was slower.
74 L. Hove-Madsen et al.
apparently slower electrode response may result from inhomogeneities rather than
a slower electrode response per se. This is illustrated in Fig. 4, where free [Ca2þ]was monitored with Ca2þ electrode and indo-1 simultaneously in a myocyte
suspension in the absence and presence of oxalate.
In Fig. 4A, Ca2þ addition to the cells causes a rapid increase in [Ca2þ], which is
subsequently sequestered by the sarcoplasmic reticulum (SR). In the absence of
10 mM oxalate (Fig. 4A), the electrode response appears to be slower than the
corresponding indo-1 signal. However, when oxalate is subsequently added to the
cell suspension (Fig. 4B), the measured change in free [Ca2þ] after a Ca2þ addition
is similar for indo-1 and the Ca2þ electrode. It should be noted that oxalate not
only buVers the free [Ca2þ], but also increases the Ca2þ uptake rate in the SR, and
thereby the removal of Ca2þ from the cell suspension. Thus, despite inducing a
faster rate of change in free [Ca2þ], oxalate eliminates the diVerence between Ca2þ
electrode and indo-1 signal by eliminating spatial inhomogeneities in free [Ca2þ] inthe myocyte suspension. Indeed, indo-1 is expected to be less sensitive to spatial
inhomogeneities as it diVuses into the permeabilized cells and binds to cellular
proteins (Hove-Madsen and Bers, 1992, 1993b). In contrast, the Ca2þ electrode
can only measure the Ca2þ outside the permeabilized cells, and inhomogeneities
during uptake or release of Ca2þ from the cells are therefore likely to occur,
resulting in erroneous measurements with the Ca2þ electrode.
2.5
0
2
1.5
1
0.5
0
15
10
5
0Fr
ee [C
a] (mM
)
Free
[Ca]
(mM
)
250 500
Time (s) Time (s)
750
Ca electrode
Ca electrodeIndo-1 Indo-1
1000 0 50 100 150
Fig. 4 EVect of inhomogeneities in [Ca2þ] on a Ca2þ-selective minielectrode and indo-1 response in
permeabilized rabbit ventricular myocytes. Panel A shows simultaneous measurements of Ca2þ uptake
in digitonin-permeabilized myocytes with both a Ca2þ-selective minielectrode and indo-1. Ca2þ uptake
in mitochondria was inhibited with ruthenium red. Ca2þ was added at time¼0 and was largely
accumulated by the SR. Under these conditions, the response of the Ca2þ electrode was slower than
indo-1. Panel B shows Ca2þ uptake by precipitating intra-SR Ca2þ and thereby preventing buildup of a
[Ca2þ] gradient. Notice that Ca2þ uptake is much faster in the presence of oxalate with no apparent
diVerence between electrode and indo-1 response (from Hove-Madsen and Bers (1993a) with
permission).
3. Calcium Selective Mini- and Microelectrodes 75
3. Storage of Minielectrodes
After PE tubes have been dipped in an ETH 129 membrane solution and allowed
to dry overnight, the dry electrodes can be stored in a closed glass vial for long
periods. We have filled minielectrodes that had been stored for 3 years and the
electrodes made with PE tubing still had a resistance of 1–2 MO with a linear
response down to less than 10 nM Ca2þ after filling. Electrodes made with PVC
tubing had higher resistance (�50 MO) but were also functional, although slower
and less sensitive. Storage of the electrodes in plastic vials results in ‘‘Ca2þ-selectiveplastic containers,’’ as the ETH 129 slowly diVuses into the container. Once the
Ca2þ electrodes are filled with the filling solution, however, the response time
increases and the electrodes gradually lose sensitivity.
B. Application of Minielectrodes
Minielectrodes can be used for a number of purposes. The most straightforward
application is the preparation of solutions where Ca2þ is buVered with chelators
such as EGTA, EDTA, or BAPTA as described by Bers (1982). We have developed
a spreadsheet that allows calculation of the actual pCa of these solutions, based on
the Nernstian response of the minielectrodes (see Chapter 1). Furthermore, the
spreadsheet allows determinations of the Kd and the purity of the Ca2þ chelator
used to prepare the solution. Thus we have used the minielectrodes to determine
76 L. Hove-Madsen et al.
the Kd for EGTA, BAPTA, and oxalate in buVer solutions (Bers, 1982; Harrison
and Bers, 1987, 1989; Hove-Madsen and Bers, 1993a). More comprehensive pro-
grams to calculate the amount of Ca2þ and Ca2þ buVer needed to prepare solu-
tions have been developed (e.g., MaxChelator by C. Patton, Marine Biology
Institute, Monterey, CA; http://maxchelator.stanford.edu/downloads.htm; cf.
Chapter 1).
We have also used the minielectrodes to characterize the binding of Ca2þ to
indo-1 in vitro and in cell suspensions, in order to calibrate the indo-1 and furaptra
signals when used in cell suspensions (Hove-Madsen and Bers, 1992; Hove-
Madsen et al., 1998; Shannon and Bers, 1997). In agreement with previous studies
of fura-2 (Konishi et al., 1988) we found that indo-1 binds extensively to cellular
proteins and causes a � fourfold increase in the Kd for Ca2þ-indo-1 in permeabi-
lized myocytes.
The minielectrodes can also be used to titrate the passive Ca2þ binding sites in
permeabilized myocytes where the cellular Ca2þ uptake and release process are
inhibited. Using the same titration method, we have measured total Ca2þ uptake in
the SR in permeabilized myocytes, by inhibiting Ca2þ uptake in the mitochondria
and release of Ca2þ through the SR Ca2þ release channels (Hove-Madsen and
Bers, 1993a).
Finally, we have used the minielectrodes together with indo-1 for online mea-
surements of the Ca2þ uptake rate in the SR in permeabilized ventricular myocytes
(Hove-Madsen and Bers, 1993b) and to examine the eVects of phospholamban
phosphorylation and temperature on the uptake rate (Hove-Madsen et al., 1998;
Mattiazzi et al., 1994), and we have determined the inhibition of Ca2þ uptake by
thapsigargin in order to measure the number of SR Ca2þ pump sites (Hove-
Madsen and Bers, 1993b). In experiments measuring Ca2þ uptake rates caution
should, however, be taken when using minielectrode because of the above men-
tioned possibilities of inhomogeneities in Ca2þ in cell suspensions and slowing of
the electrode response.
C. Preparation of Ca2þ-Selective MEs
1. Glass Tubing Preparation
We have used nonfilamented capillaries (150 mm outer diameter, 15 cm long,
from Clark Electromedical Instruments, UK or World Precision Instruments,
USA). The glass is cut in the middle with a diamond pen or glass scorer (with
care to avoid deposition of dirt). Both ends of the capillaries (now 7.5 cm long) are
lightly fire-polished; a whole batch can be prepared and kept in a small glass
beaker, preferentially in a dust-proof container. Cleaning of the micropipettes
prior to pulling has been a matter of debate, but as 99.8% of the glass after pulling
is newly exposed (Deyhimi and Coles, 1982), we do not find such procedures to be
necessary.
3. Calcium Selective Mini- and Microelectrodes 77
2. Microelectrode Pulling and Silanization
MEs can be prepared ‘‘on-demand’’ and in ‘‘batch’’ methods. The first method
consists in preparing (i.e., pulling and silanizing) MEs on the experimental day,
which has the obvious advantage of having ‘‘freshly’’ prepared electrodes and of
being able to manufacture as many as desired a day. Protocols have been described
(reviewed in Ammann, 1986). The batch method that we use consists of preparing
a batch of MEs that can be kept for several days in a dry, dust-proof container.
MEs are pulled on a programmable horizontal stage puller (Model P80 PC or
comparable, Sutter Instruments, USA). By trial-and-error, and according to the type
of biological preparations studied, a satisfactory shape ofMEcanbe found.However,
several aspects ofMEpulling have to be taken into account when designing its shape.
As ion-selective MEs have an intrinsically high resistance (i.e., >50–100 GO), thesignal-to-noise ratio has to be minimized. This can be achieved in two ways: the first,
and most obvious, is to increase the tip diameter, within certain limits, which are
dictatedby the sizeof the cell type. Inour case,we impale cardiac cells inwholemuscle.
Cell length is typically between 50 and 150 mm and tip diameters less than 0.5 mm are
needed. However, there is a trade-oV between tip diameter and detection limit of the
electrode (see below). That is, increasing tip diameter improves electrode response (in
terms of detection limit and speed of response), but is more likely to damage the cell
during impalement. Another simple way to decrease the resistance is to decrease the
length of the ME shank. This further helps to reduce capacitative artifacts, that are
encountered when the level of physiological solution fluctuates in the experimental
bath (Vaughan-Jones and Kaila, 1986), and also helps electrolyte filling (see below).
In contracting muscular preparations, the shank should also possess some flexibility
to avoid dislodging of the ME during a contraction. Again, by trial-and-error, an
adequate shape that fulfills all these requirements can be found. Their shape was
designed to reduce their resistance by making them steeply tapered (having a shank
length of approximately 150 mm and diameter of 20 mm at 10 mm above the tip).
Under lightmicroscopical observation, the tip diameterwas estimated tobe�0.5 mm.
MEs are dehydrated, tip up in an aluminium block, at 200 �C for 12 h. Our
experience, and of others (Vaughan-Jones and Wu, 1990), has been that a better
silanization and longer lifetime of the MEs are achieved this way.
The silanization protocol consists of spritzing 300 ml of N,N-dimethyltrimethyl-
silylamine (Fluka) onto the aluminum block and rapidly placing a glass lid on top
of the dish. Care must be taken not to inhale vapor from the silane vial or during its
introduction in the dish. In our hands, once opened, the silane vial can be kept for
at least 2 months without losing its properties. Silanization procedure lasts for
90 min. The lid is then removed and the MEs are baked for another 60 min, which
drives oV the excess silane vapor. The aluminum block and the MEs are then
placed into an air-tight plastic container, also containing desiccant. We advise not
to keep the MEs more than a week, because repetitive openings of the container
and insuYcient seal quality will progressively make the electrodes lose their
hydrophobicity.
78 L. Hove-Madsen et al.
3. Electrolyte Filling of the ME
The filling or conducting electrolyte is introduced in the ME from the back
(back-filling), but because of the hydrophobicity of the glass wall (due to the
silanization), the tip does not get filled immediately. The strategy to fill the ME
completely depends on the absence or presence of the ligand at its very tip.
In the case where the ME is first filled with the electrolyte, the back of the ME is
connected, via a flexible tubing, to a 50-ml syringe, whose plunger has been
previously pulled to 5 ml. Positive pressure is then applied and, as assessed by
microscopic observation, the electrolyte creeps along the ME wall and fills the tip.
The biggest air bubbles are then removed by gently flicking the ME, held tip down.
Although we have initially applied this procedure to filamented MEs, a similar
success rate (more than 90%) has been obtained with nonfilamented MEs
(surprising, because silanization is expected to limit aqueous filling ease).
Whatever the method used, if the electrolyte column was interrupted by air
bubbles then, gentle heating of the tip of theME, under microscopic control, with a
tungsten–platinum wire, according to the device described by Thomas (1982), can
remedy this problem. Once filled with electrolyte to the tip, the hydrophobic ligand
can be introduced (see below).
In the case where the electrolyte is added after the Ca2þ ligand, additional
problems exist. In fact, caution has to be taken to avoid the presence of air at the
electrolyte–ligand interface. If a traditional whisker is used, great care has to be
taken not to accidentally disrupt the column of ligand, which can lead to mixing of
oil and water, making unstable ion-selectiveMEs. If a heating filament is used, care
has to be taken not to heat the ligand because it is likely that the local high
temperature may damage the ligand properties.
We have used a filling solution that has an ionic composition mimicking the
intracellular medium (in mM): Naþ: 10; Kþ: 140; HEPES: 10; EGTA: 1; pH 7.1
(at 30 �C) and pCa 7. This solution is in fact identical to the calibrating solution
having the same pCa (see below and also Orchard et al. (1991)) for additional
comments).
Our experience has been that it is best to minimize the time between electrolyte
and ligand filling. We prefer to fill the ME with the ligand as soon as the ion-
selective ME is filled with the electrolyte (although we managed to draw the ligand
in the tip 2–3 h after electrolyte filling). If MEs are left overnight with the filling
solution, it can be very diYcult to draw ligand into the tip; this observation could
be explained by a progressive glass hydration, causing it to lose its capability to
retain the ligand.
4. Preparation and Use of the Ca2þ-Selective Ligand
For repetitive and long-lasting Ca2þ measurements with Ca2þ-selective MEs
dissolving the Ca2þ ligand in a ‘‘cocktail’’ containing PVC is useful (Tsien and
Rink, 1981). Although the cocktail available from Fluka (Cocktail II containing
Table IIPreparation of the PVC-based Ca2þ-selective ligand MEs
Componenta Amount
Solution 1
ETH 129 (or ETH 1001) 27.5 mg
NPOE 500 mlNaþ-tetraphenyl borate 5.53 mg
Solution 2
Solution 1 200 mlPVC 36 mg
THF 400 ml
aComponents can be obtained from Sigma-Aldrich (St. Louis,
MO) and see Table I for abbreviations.
3. Calcium Selective Mini- and Microelectrodes 79
94% (w/w) NPOE, 5% (w/w) ETH 129, and 1% (w/w) sodium TPB) is satisfactory
to start with (Ammann et al., 1987), small volumes sometimes provided (0.1 ml) do
not facilitate handling. We prefer to make up our own cocktail, in a larger NPOE
volume, with the same proportions. The composition, for 500 ml of NPOE is given
in Table II.
Because of the small volumes and required stirring, it is preferable to work with
flat-bottomed, small volume glass vials, and miniature stirring bars. ETH 129 and
TPB are dissolved with vigorous stirring in NPOE, in a 2-ml glass vial (Solution 1).
Solution 1 can be kept at room temperature for several months, with a Teflon
screw cap, protected from light. When the final cocktail (Solution 2) is prepared,
PVC is dissolved in THF with stirring. 0.2 ml of Solution 1 is then added, stirred,
and finally sonicated. THF is allowed to partially evaporate to approximately half
of the initial volume and the final cocktail is finally poured in a 0.5-ml conical vial
(Clark Electromedical Instruments).
Before dipping the ME tip, THF is allowed to evaporate until the mixture has
the consistency of a thick syrup. Experience helps to determine the adequate
consistency of the ligand. In fact, if not enough THF has evaporated, the ETH
129 sensor is too diluted and the evaporation in theMEmay cause retraction of the
gel, yielding poor responses. On the other hand, in some instances, we have
managed to fill the electrodes even if the ligand appeared to be solidified (as a
rule, the thickest mixture which will fill the tip is best).
Because of the small diameter of the tip and the viscosity of the ligand, negative
pressure is required at the back of the electrode. This is achieved by a >10-ml
syringe connected, via a 3-way stopcock, to a flexible Teflon tubing connected to
the back of the electrode with soft tubing. Vacuum is then applied by pulling the
plunger out and by blocking it with a rod/block or collar placed along the plunger
(care should be taken to regularly check the vacuum of the system). Observation of
the ME and measurements of the ligand column height are performed under
80 L. Hove-Madsen et al.
microscopic control, with a microforge (e.g., as described by Thomas (1982). In
brief, the microscope body is laid on its back so that the stage (removed) would be
vertical and the eyepieces are oriented upwards. A long-working distance objective
(40�) is used and the ME and the ligand vial are held independently by two
micromanipulators.
Depending on glass, shape, and tip diameter 10–30 min of negative pressure is
typically required to fill the tip. Vacuum is slowly released before lifting the
electrode from the ligand. A column height of less than 300 mm is preferable,
because it decreases the electrode sensitivity to changes in temperature and level
of the bath in the experimental chamber (Vaughan-Jones and Kaila, 1986). Our
experience is that, depending on the tip diameter, column heights between 50 and
250 mm yielded acceptable electrode responses.
Once the electrode is filled with both the ligand and the electrolyte, we prefer not
to let the ME equilibrate in Tyrode or high pCa solutions, because this favors the
deposition of dirt on the tip of the ME and might contribute to clogging. Rather,
we place them, tip up, in a drilled plastic plate, protected by an upside down glass
beaker. Before calibration, the column height is rechecked because THF in the
column continues to evaporate, leading to shrinkage of the PVC gel.
As THF evaporates, the stock solution becomes even thicker. Periodically
enough THF must be added to decrease the viscosity of the mixture. This process
is hastened by mixing it with a glass rod and then on a Vortex mixer (maximal
setting). Sonication can also be used, but does not give better results.
5. Double-Barreled Ca2þ-Selective MEs
For Ca2þ-selective MEs measurements, one must measure both the potential of
the Ca2þ electrode and the local voltage (typically with a KCl-filled ME; see
below). Both electrodes can be built into a single double-barreled electrode,
where one barrel is the Ca2þ electrode and the other is the voltage electrode. We
have not had much luck using these for intracellular recording, but they can be
extremely useful for measurement of local extracellular or interstitial [Ca2þ] inmulticellular preparations (Bers, 1983, 1985, 1987; Bers and MacLeod, 1986;
Shattock and Bers, 1989).
Double-barreled electrodes can be pulled from 2 to 2.5 mm diameter theta-style
tubing (R and D Optical Systems, MD) on a Brown–Flaming P-77 micropipette
puller (Sutter Instruments, CA, USA). For extracellular recording, the tips of the
electrodes are carefully broken under microscopic observation to have 4–12 mmoverall tip diameters. Two methods of silanization of the Ca2þ barrel are practical.
(1) Distilled water is injected into the reference barrel. A hypodermic needle
containing tri-n-butylchlorosilane is introduced into the Ca2þ barrel, �1 ml ofsilane is displaced into the shank of each electrode, and electrodes are placed
in a 200 �C oven, tips up, for 5 min and then cooled. (2) a stream of silanizing N,
N-dimethyltrimethylsilylamine (TMSDMA vapor is passed through the Ca2þ
barrel (with or without warming), while a stream of nitrogen gas is passed through
3. Calcium Selective Mini- and Microelectrodes 81
the reference barrel (under pressure) to prevent silanization of the reference barrel
(which would result in both barrels being Ca2þ-sensitive).The larger tips of these electrodes make the filling easy. Both barrels can be
easily backfilled. The silanized barrel is backfilled with a reference solution con-
taining 10 mMCaCl2 and 100 mMKCl and the nonsilanized barrel with a solution
containing 140 mM NaCl. A column of the neutral Ca2þ ion-exchange cocktail
ETH 1001 or 129 (Fluka Chemical, Ronkonkoma, NY) 50–250 mm long is easily
drawn into the silanized barrel. Ca2þ electrodes with these tip diameters exhibit
Nernstian behavior over the range 10 mM–10 mM Ca2þ (Bers and Ellis, 1982).
The resistance of these MEs was typically 1–5 GO for the Ca2þ-sensitive barrel and1–4 MO for the reference barrel.
The impedance of the two barrels is very diVerent, but their fast response allowsrelatively rapid interstitial [Ca2þ] monitoring. To match signal response kinetics to
voltage steps a variable-passive R-C filter can be added to the reference barrel
signal after the signal has come from operational and oVset amplifiers. This filter is
adjusted while a square voltage pulse is fed into the bath until the best matching
with the Ca2þ barrel response is obtained (Bers, 1983). These Ca2þ electrodes
typically exhibit Nernstian behavior at least over the range 10 mM–10 mM Ca2þ.This is satisfactory for typical extracellular [Ca2þ] measurements and the double-
barreled electrodes are easier to calibrate and use than intracellular impalements
(described in more detail below).
6. Calibrating Bath and Solution Perfusion
It is preferable to calibrate ion-selective MEs in the experimental chamber in
which measurements are made or to have the calibrating bath as close as possible in
design and proximity. Our calibration chamber is a ‘‘flow-through’’ type (volume:
0.1 ml), immediately adjacent to the experimental chamber. Note that it is conve-
nient that the experimental chamber is viewed from the front, and not from above.
7. Calibration Procedure
The bath electrode is either an Ag wire (chlorided by dipping it in bleach for
15–20 min) or an agar bridge. Ideally, a conventional electrode (3 M KCl filled)
should also be immersed in the bath, and the diVerential voltage (ion-selective ME
minus conventional) should be read. We use a commercial amplifier (FD-223 from
WPI) or a home built amplifier using varactor bridge preamplifiers (AD311J,
Analog Devices) as described by Thomas (1982).
We have adopted the following method to quickly select suitable MEs. The ME
is mounted in its holder and advanced into the calibrating bath, allowing the trace
to stabilize. If the device used to measure the signal has a resistance measurement
feature, it is worth measuring this parameter. In fact, our experience has been that
for the sharp Ca2þ MEs having resistances ranging between 100 and 250 GO were
suitable for our experiments, in terms of linearity and detection limit of the
82 L. Hove-Madsen et al.
calibration curve. Within this range, the higher the resistance, the lower the
detection limit for a given batch of MEs. At resistances higher than 300 GO, thedetection limit decreased sharply, probably because of the small tip diameter
(Ammann, 1986). Finally, although low resistance MEs tended to also have low
detection limits, they were not suitable for our experiments, because of the large tip
diameter that could seriously damage the cell membrane during impalement. By
contrast, we have found the height of the ligand column not to be a valuable
predictor of ME performance (although this height was always kept 50–250 mm).
In our hands, an ME can be used a few minutes after ligand filling.
After equilibration in the control physiological solution (in our case, an HEPES-
based Tyrode, containing 2 mM Ca2þ), ion-selective ME potential is adjusted to
0 mV. Our 2 mM Ca2þ Tyrode gives a voltage reading corresponding to an
intracellular calibrating solution of pCa 2.6. Then, flow is switched to a solution
of high pCa (between 7.5 and 9). At 30 �C (our experimental temperature), the
theoretical slope of the relationship between voltage and pCa is �30 mV/pCa, so
that, between pCa 2.6 and pCa 8, the theoretical voltage should be �162 mV.
However, as the electrode detection limit decreases at high pCa, a practical
compromise is often necessary for acceptability (e.g., readings more negative
than �150 mV). A Ca2þ-selective ME meeting this criteria may then be calibrated
over a wider range of [Ca2þ]. After the calibration is completed, the ME is moved
into the experimental chamber and equilibrated until stable. Conventional,
3 M KCl-filled MEs are pulled from the same glass and with the same character-
istics as the Ca2þ electrodes, but are not silanized.
D. Application of Ca2þ-Selective MEs
1. Sharp Ca2þMEs for [Ca2þ]i Measurement
We and others have had some, but limited success in making reliable [Ca2þ]imeasurements in cardiac myocytes with these electrodes (Bers and Ellis, 1982;
Marban et al., 1980), and a few other groups have had some luck with other cell
types such as skeletal muscle fibers (Allen et al., 1992; Blatter and Blinks, 1991;
Lopez et al., 2000), Aplysia neurones (Gorman et al., 1984), and photoreceptors
(Levy and Fein, 1985) using an earlier developed Ca2þ ionophore (ETH 1001;
Sigma-Aldrich 21192) instead of ETH 129 in the cocktail. ETH 1001-based MEs
generally do not make electrodes that have quite as low a detection limit as ETH
129 (Schefer et al., 1986), but for some reason ETH1001 seems to be of greater
practical utility for intracellular Ca2þ MEs. We have done some preliminary
cardiac muscle experiments that are consistent with this notion (resting
pCa¼6.34�0.15; mean�SD; n¼10 determinations). However, with the excellent
fluorescent Ca2þ indicators available now that are easy to use, one would need a
compelling reason to tackle this challenging electrophysiological approach for
[Ca2þ]i and the references above should then help.
Control
200 ms 100 ms
Rabbit ventricle Rat ventricle
[Ca]
o mM
Control
Control550
Control
+Citrate
+Citrate
+Citrate
+Citrate
500480
5202.5 m
N/m
m2
5 mN
/mm
2
500
490
A B
Fig. 5 Measurements of [Ca2þ]o with double-barreled Ca2þ-selective MEs during single steady state
contractions in (A) rabbit and (B) rat ventricular muscle (0.5 Hz, 30 �C). The [Ca2þ]o and tension are
shown in the absence and presence of 10 mM citrate (which limits [Ca2þ]o depletion by buVering
[Ca2þ]o. Bath [Ca2þ]o¼0.5 mM (dotted line). Data was from Shattock & Bers, (1989), as presented in
Bers (2001) (with permission).
3. Calcium Selective Mini- and Microelectrodes 83
2. Measuring Extracellular [Ca2þ] with Double-Barreled MEs
Double-barreled Ca2þ MEs can record rapid changes in extracellular [Ca2þ]([Ca2þ]o) between cells in multicellular preparations such as isolated cardiac tra-
beculae (Bers, 1983, 1985, 1987; Bers and MacLeod, 1986; Shattock and Bers,
1989). Figure 5A shows that one can detect small [Ca2þ]o depletions during
individual steady state rabbit cardiac action potentials and contractions. More-
over, when [Ca2þ]o is buVered by the low aYnity fast buVer citrate these depletionscan be suppressed. Note that these [Ca2þ]o depletions reflect net cellular Ca2þ
influx (in excess of eZux) early in the contraction and net Ca2þ eZux later in the
contraction, such that [Ca2þ]o returns to the bath level. In cardiac myocytes the
depletion is driven mainly by Ca2þ influx via Ca2þ channel current and to some
extent by Naþ/Ca2þ exchange (which can mediate Ca2þ influx at positive Em when
[Ca2þ]i is low). As [Ca2þ]i rises in the cell during the heartbeat because of Ca2þ
entry and SR Ca2þ release, it causes enhanced Ca2þ eZux (mainly via Naþ/Ca2þ
exchange in cardiac myocytes), and this allows [Ca2þ]o to recover. Note that action
potential repolarization greatly enhances the driving force for Ca2þ eZux via Naþ/Ca2þ exchange, further enhancing the recovery of [Ca2þ]o to the bath level.
In rat ventricular muscle the [Ca2þ]o signals are remarkably diVerent (Fig. 5B).In the rat there is only a very brief phase of [Ca2þ]o depletion (for �20 ms), which
gives way to a large rise in [Ca2þ]o during the contraction. At first this result
seemed surprising in light of the rabbit results in Fig. 5A. However, when we
consider the diVerences in action potential shape and that [Naþ]i is higher in rat
ventricular myocytes (Shattock and Bers, 1989), the explanation became clear. The
rat (and mouse) ventricle exhibit very short action potential duration compared to
30
60
−30
0
30
−90
−60
0
30
0
−60
−30
A
Em
or
EN
a/C
a (m
V)
Na/
Ca
exch
ange
driv
ing
forc
e(E
Na/
Ca−
Em
) m
V
84 L. Hove-Madsen et al.
rabbit (or human ventricle), and this drives rapid Ca2þ extrusion via Naþ/Ca2þ
exchange at a time when [Ca2þ]i is very high (Fig. 6B). In the rabbit ventricle, the
longer action potential plateau keeps Naþ/Ca2þ exchange in check, delaying
extrusion until a later time where [Ca2þ]i is lower. Another implication of
Fig. 5B is that there is net Ca2þ eZux during the contraction in rat (vs. net influx
in rabbit). This means that there must be net Ca2þ influx between contractions in
rat ventricle, and the [Ca2þ]o trace in Fig. 5B is actually going below the bath by the
end of the trace to restore the steady state balance before the next beat (�1.5 s
later). Note that during a steady state heartbeat, total Ca2þ influx must equal total
Ca2þ eZux (i.e., there is no net gain or loss of Ca2þ at the steady state).
Extracellular Ca2þ-MEs are also useful for assessing nonsteady state Ca2þ fluxes
on a longer time scale (Bers and MacLeod, 1986; MacLeod and Bers, 1987).
Figure 7A shows that when 0.5 Hz stimulation is stopped there is a very slow
small rise in [Ca2þ]o over many seconds (net Ca2þ eZux), and upon resumption of
stimulation (now at 1 Hz) that there is a net [Ca2þ]o depletion which develops over
Rabbit ventricle Rat ventricleB
Ca efflux
250 500
Ca influx
Time (ms)
0 250 500
Time (ms)
aNai= 7.2 mM aNai= 7.2 mM
ENa/CaENa/Ca
Em
Em
Fig. 6 Changes in the reversal potential of the Naþ/Ca2þ exchange (ENa/Ca) during the action
potential (Em) and Ca2þ transient in rabbit and rat ventricle. Changes in electrochemical driving force
for Naþ/Ca2þ exchange (ENa/Ca�Em) are shown in the bottom panels, assuming a 3:1 stoichiometry of
Naþ/Ca2þ exchanger and aNai are measured Naþ activity values (Shattock & Bers, 1989). Ca2þ
transients driving the contraction are assumed to be the same for both species (resting [Ca2þ]i¼150 nM,
peak [Ca2þ]i¼1 mM, 40 ms after the AP initiation). Note that Ca2þ eZux is low during rest in
rabbit myocytes because of the low [Ca2þ]i (despite a significant driving force). Based on data in
Shattock & Bers, (1989), as modified in Bers (2001), with permission.
Tension200mN
A Control
B Ryanodine
0.2 mM[Ca]o
[Ca]o1 mV
[Ca]o1 mV
Tension80mN
0.2 mM [Ca]o
30 s
Fig. 7 Measurements of [Ca2þ]o using double-barreled extracellular Ca2þ MEs in rabbit ventricular
muscle. Local [Ca2þ]o and tension are shown starting during steady state stimulation at 0.5 Hz with a
couple of pauses (0.5 and 1 min) and a period of 1 Hz stimulation. Spikes on [Ca2þ]o trace are stimulus
artifacts, and bath [Ca2þ]o was 0.2 mM. The same protocol was used before (A) and after equilibration
with 1 mM ryanodine (B; from Bers & MacLeod, (1986), with permission).
3. Calcium Selective Mini- and Microelectrodes 85
several beats. What we showed with other experiments was that this depletion
reflects the filling of SR Ca2þ stores (i.e., net transfer of Ca2þ from the extracellular
space to the SR). Moreover, the rise in [Ca2þ]o during rest reflects the gradual loss
of SR Ca2þ that depends on diastolic leak of Ca2þ from the SR and net Ca2þ
extrusion by Na/Ca exchange (thermodynamically favored in resting rabbit ven-
tricular muscle; Fig. 6A). This also allowed new insight at the time as to exactly
how ryanodine works in cardiac myocytes. Figure 7B shows that loss of cellular
Ca2þ during rest is much faster after ryanodine exposure, but that during 1 Hz
pacing the cell (and SR) can take up Ca2þ. This showed that ryanodine makes the
SR leaky, but not so much as to abolish SR Ca2þ uptake (i.e., Ca2þ could still be
driven into the SR). Once the repeated high [Ca2þ]i signals stop driving SR Ca2þ
uptake, the Ca2þwithin the SR is lost very quickly. Thus dynamics of cellular Ca2þ
flux balance can be readily assessed by double-barreled Ca2þ-MEs, both with
relatively high time resolution during steady state conditions and for longer
changes that occur in nonsteady state conditions. This makes them a nice comple-
ment to fluorescent indicators and voltage clamp studies.
3. Troubleshooting Ca2þ-Selective MEs
The ME cannot be filled with the ligand:
1. The ligand may be too thick because of the THF evaporation. Redilute PVC
by adding small amounts of THF and stirring the mixture to homogeneity.
86 L. Hove-Madsen et al.
2. The tip diameter is too small. This may entail preparing MEs with diVerentshapes and testing them (ligand filling and calibration) on the same day. Relying on
ME resistance may give misleading results because it is also aVected by the
geometry of the shank.
3. ME silanization may be insuYcient, because of insuYcient time of silaniza-
tion (or insuYcient beaker seal during exposure to silane vapors), old silane, glass
rehydration (storage problem), and insuYcient dehydration.
The ME gives bad calibration curves (subNernstian slopes, low detection limit).
1. Make sure that the calibrating solutions are adequate. Check them with
commercial macroelectrodes or the above described ETH 129-based minielec-
trodes, as described in previously published methods (e.g., Bers, 1982).
2. The ligand may be too old. This is a common occurrence. Ligand lifetime is
sensitive to exposure to light and air.
3. The ETH 129 is too diluted. This can occur by not letting THF evaporate
enough before ligand filling. In this case, it is better to let all the THF evaporate
and then, to re-add small amounts (few tens of microliter) of THF and stir the
mixture until a syrup-like solution is obtained. It can also happen if after adding
THF to a gelled cocktail, the cocktail is not mixed to homogeneity (by vortexing or
sonication).
IV. Discussion
We here describe the design and preparation of Ca2þ-selective mini- and micro-
electrodes, based on the ligand ETH 129 in a PVC matrix. Both mini- and micro-
electrodes have excellent responses during in vitro calibration with a response time
and detection limit superior to that of most commercially available minielectrodes.
These electrodes can have multiple applications. We do note, however, that for
measuring [Ca2þ]i, the ligand ETH 1001 may be preferable.
Other Ca2þ-selective ionophores with a higher selectivity for Ca2þ against other
ions, such as K23E1 (Suzuki et al., 1995), have been developed after ETH 129 but
their detection limit for calcium is considerably inferior to that of ETH 129-based
electrodes (Suzuki et al., 1995). Thus, K23E1 may be useful for clinical analysis of
calcium in plasma or other biological fluids, but ETH 129-based electrodes remain
superior for calcium measurements in the submicromolar concentration range.
ETH 129-based minielectrodes are economical, easy to prepare, and have suc-
cessfully been used for purposes where the response time of the electrode is appro-
priate. This includes preparation of calibration solutions, determination of the Kd
for EGTA, BAPTA, and oxalate in buVer solutions (Bers, 1982; Harrison and Bers,
1987; Harrison and Bers, 1989; Hove-Madsen and Bers, 1993a), and calibration of
indo-1 and furaptra signals cell suspensions (Hove-Madsen and Bers, 1992; Hove-
Madsen et al., 1998; Shannon and Bers, 1997). The minielectrodes have also been
3. Calcium Selective Mini- and Microelectrodes 87
used to titrate the passive Ca2þ binding sites in permeabilized myocytes where the
cellular Ca2þ uptake and release process are inhibited. The same titration method
has also been used to measure total Ca2þ uptake in the SR of permeabilized
cardiomyocytes (Hove-Madsen and Bers, 1993a).
Ca2þ minielectrodes have also been used together with indo-1 for online mea-
surements of the Ca2þ uptake rate in the SR in permeabilized ventricular myocytes
(Hove-Madsen and Bers, 1993b), to examine the eVects of phospholamban phos-
phorylation and temperature (Hove-Madsen et al., 1998; Mattiazzi et al., 1994)
and to measure the number of SR Ca2þ pump sites by titration with the selective
pump inhibitor thapsigargin (Hove-Madsen and Bers, 1993b; Hove-Madsen et al.,
1998). In these applications, the electrode response time can become a limiting
factor and it is important to use a fresh Ca2þ electrode for each experiment, and to
minimize inhomogeneities in cell suspensions that the calcium electrodes cannot
detect.
Although the use of Ca2þ selective electrodes for measurements of dynamic
changes in some biological systems is limited by their response time (particularly
at submicromolar concentration), the potential benefits of combining Ca2þ selec-
tive electrodes with other experimental techniques are underexplored. Indeed,
calcium selective electrodes have successfully been used to monitor [Ca2þ] insidepatch pipettes (Kang and Hilgemann, 2004; Kang et al., 2003). New Ca2þ sensors
based on coating of microcantilevers with ion-selective self-assembled monolayers
have also been developed (Ji and Thundat, 2002) and may be useful in the mapping
of Ca2þ channels and transporters on the cell surface. Indeed, measurement of the
change in extracellular ion concentration with ion-selective MEs has also been
shown to provide a noninvasive means for functional mapping of the location and
density of potassium channels (Korchev et al., 2000; Messerli et al., 2007) and for
the quantification of transmembrane Ca2þ flux (Bers, 1983, 1985, 1987; Bers and
MacLeod, 1986; Shattock and Bers, 1989; Smith et al., 1999).
Thus, in spite of the overwhelming predominance of fluorescent Ca2þ indicators,
Ca2þ-selective electrodes and biosensors still remain a valuable supplement to
many imaging and electrophysiological techniques in molecular and cellular
physiology.
Acknowledgments
This work was supported by a grant from the National Institute of Health (HL30077) to DMB and a
grant from the Spanish Ministry of Science and Technology (SAF2007-60174) to LHM.
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CHAPTER 4
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
Construction, Theory, and PracticalConsiderations for using Self-referencingof Ca2þ-Selective Microelectrodes forMonitoring Extracellular Ca2þ Gradients
Mark A. Messerli and Peter J. S. SmithBioCurrents Research CenterCellular Dynamics ProgramMarine Biological LaboratoryWoods Hole, Massachusetts, USA
A
OGY,All rig
bstract
VOL. 99 0091hts reserved. 91 DOI: 10.1016/S0091
-679X-679X
I. In
troduction II. C aSM ConstructionA.
Micropipette Fabrication B. Silanization C. Microelectrode ConstructionIII. P
roperties of CaSMs A. Response to Ion Activity B. Selectivity C. Spatial Resolution D. Response TimeIV. S
elf-referencing of CaSMs A. DiVerential Concentration Measurement B. DiVerential Concentration Determination C. Calculation of Flux D. Correction for Ca2þ BuVering E. Measurement of Voltage Gradients F. Positional ArtifactsV. C
a2þ Flux Measurements R eferences/10 $35.00(10)99004-3
92 Mark A. Messerli and Peter J. S. Smith
Abstract
Ca2þ signaling in the extra- and intracellular domains is linked to Ca2þ
transport across the plasma membrane. Noninvasive monitoring of these result-
ing extracellular Ca2þ gradients with self-referencing of Ca2þ-selective micro-
electrodes is used for studying Ca2þ signaling across Kingdoms. The quantitated
Ca2þ flux enables comparison with changes to intracellular [Ca2þ] measured with
other methods and determination of Ca2þ transport stoichiometry. Here, we
review the construction of Ca2þ-selective microelectrodes, their physical charac-
teristics, and their use in self-referencing mode to calculate Ca2þ flux. We also
discuss potential complications when using them to measure Ca2þ gradients near
the boundary layers of single cells and tissues.
I. Introduction
Regulation of resting [Ca2þ]i and the control of spatial and temporal dynam-
ics during Ca2þ signaling require coordinated transport between membrane-
separated compartments, giving rise to Ca2þ fluxes across organelles and the
plasma membrane. Movement of Ca2þ across the plasma membrane via trans-
porters, exchangers, or channels gives rise to minute gradients of [Ca2þ] in the
extracellular boundary layer that reflect changes in [Ca2þ]i. The near real-time
extraction of these gradients requires a detection method that is not disturbing
to the local chemical environment, functions over a wide dynamic range, and
possesses high sensitivity, selectivity, and spatial resolution. For these reasons
extracellular Ca2þ gradients have been monitored with self-referencing of Ca2þ-selective microelectrodes (CaSMs), enabling noninvasive characterization of
Ca2þ transport and signaling events. Unlike most fluorescent or luminescent
indicators, CaSMs were originally developed for measuring both intracellular
and extracellular [Ca2þ] (listed in Lanter et al., 1982). Measurement of minute
Ca2þ gradients on the outside of cells was limited by electrical drift in the
system. For this reason, a modulation technique was introduced (Kuhtreiber
and JaVe, 1990) that enabled reduction of drift and provided a simple means
for calculating Ca2þ flux. The method was later coined ‘‘self-referencing’’ and
has been extended to other ion-selective microelectrodes and amperometric
microelectrodes enabling characterization of fluxes of many diVerent analytes
(Messerli et al., 2006; Smith et al., 2007). Measurement of Ca2þ fluxes with self-
referencing has enabled direct comparison of Ca2þ fluxes measured with other
techniques including radioactive tracers, fluorescent and luminescent ion indi-
cators, and voltage clamp. We will first discuss the construction and general
properties of CaSMs before discussing their use with the self-referencing
approach.
4. Electrochemical Measurement of Ca2+ Flux 93
II. CaSM Construction
A. Micropipette Fabrication
Ion-selective microelectrodes are based on an ion-selective solvent or liquid
membrane, immobilized in the tip of a glass micropipette with a backfilling
electrolyte. The glass micropipette housings are pulled from 1.5 mm outer diameter
borosilicate (TW150-4 World Precisions Instruments, Sarasota, FL), aluminosili-
cate (A150-100-10 Sutter Instruments, Novato, CA), or quartz glass (Q150-110-10,
Sutter Instruments). Inner filaments, commonly used to load electrolyte solutions
to the tips of micropipettes, are avoided. Although the glass body is fragile, it
provides distinct advantages over other materials including low cost, excellent
resistive properties necessary for use with the high-resistance liquid membranes,
and easy fabrication of small tips. Micropipettes are pulled, silanized, and stored in
bulk, �50 per wire rack. The glass is pulled down to a final edge slope of 0.15–0.17
and an inner tip diameter of 2–3 mm. Borosilicate and aluminosilicate micropip-
ettes are pulled on a horizontal heated filament puller (P-97, Sutter Instruments)
while quartz pipettes are pulled on a horizontal laser puller (P-2000, Sutter Instru-
ments). Latex gloves are worn during handling of the glass before silanization.
B. Silanization
The hydrophilic glass surfaces are coated with a hydrophobic silane to enable
adhesion and high electrical resistance between the glass and the hydrophobic
liquid membrane. While many forms of silanization exist, we prefer vapor deposi-
tion of N,N-dimethyltrimethylsilylamine (cat# 41716 Sigma-Aldrich, St. Louis,
MO) as it enables rapid and uniform coating of numerous micropipettes, simulta-
neously. A wire rack of micropipettes is placed in a small solid wall metal box
(8 cm�8 cm�10 cm) with a swinging door within the oven so that the silane vapor
can be trapped in a small region around the pipettes. Prior to coating, the glass is
dried for 20 min at 240 �C under vacuum (28 in Hg). This shortens drying time and
decreases loss of hydroxyl groups (Deyhimi and Coles, 1982; Munoz et al., 1983).
Higher temperatures may dry glass more quickly as well; however, this silane has
ignited two out of four times at �250 �C. After drying, atmospheric pressure is
recovered by purging the oven with Argon. A small volume of silane (20 mL) isdropped into a tiny glass beaker in the metal enclosure and the door to the
enclosure is closed before the oven door is closed. The glass is exposed to the silane
vapor for 20 min before removing and placing the micropipettes in a sealed bell jar
with desiccant in the bottom. Functional CaSMs have been produced from micro-
pipettes that have been stored in this manner for up to a month. This method has
reduced variation in the quality of silanization.
94 Mark A. Messerli and Peter J. S. Smith
C. Microelectrode Construction
Standard, electrolyte-based CaSMs consist of a short column of liquid membrane
(�30 mm) with a longer column of Ca2þ containing electrolyte (5 mm) used to make
electrical contact with the voltage recording headstage via a Ag/AgCl wire. Commer-
cially available vented pipette holders (WPI, Sarasota, FL, Warner Instruments,
Hamden, CT) are used to immobilize the CaSMs while loading and recording from
the high input impedance electrometers �1015 Ω (BioCurrents Research Center,
Woods Hole, MA; Molecular Devices, Sunnyvale, CA; Warner Instruments, Ham-
den,CT). CaSMs are constructed by first backfilling a fewmillimeters of the electrolyte
into a silanized micropipette with a long blunt needle and syringe, before tip loading
the liquid membrane. The backfilling electrolyte has varied from 100 nM Ca2þ buV-ered with 5 mM EGTA, 10 mMHEPES with 90 mMKCl (Tsien and Rink, 1981) to
simply 100 mM CaCl2 (Kuhtreiber and JaVe, 1990). However, based on further
discussion below it will be shown that the backfilling solution should be based on the
bath [Ca2þ] with additional electrolyte, 100 mMKCl, to make electrical contact with
the Ag/AgCl wire. Ca2þ-selective liquid membranes can be mixed in the lab or
purchased premixed (cat# 21048 (ETH1001), cat# 21196 (ETH129); Sigma-Aldrich,
St. Louis, MO).
Tip loading of the liquid membrane is performed under microscopic control,
displayed in Fig. 1A. The electrolyte filled micropipette on the right is positioned
near a loading pipette on the left, a tip brokenmicropipette that has been dip-loaded
with liquidmembrane. Both the loading pipette and the CaSM are connected to air-
filled syringes with plastic tubing so that pressure can be applied. The threaded
plunger (TP) syringe in Fig. 1A allows a small controlled pressure to create a small
bulge of liquid membrane away from the loading pipette which aids loading of the
CaSM.A plastic syringe (PS) with a three-way valve for the CaSM enables applying
and venting pressure before loading and before removing the CaSM from the
electrode holder. After positioning both the loading pipette and the CaSM within
the field of view under the microscope objective, Fig. 1B, pressure is applied to the
back of the CaSM to push the electrolyte to the tip. Pressure is vented and the tip of
the CaSM is immediately positioned within the liquid membrane bulge held in the
loading pipette. Surface tension will immediately draw the liquid membrane into
the silanizedmicropipette. A combination of pressure and suction is used to achieve
a liquid membrane column of the desired length (�30 mm). After the desired length
is achieved, the CaSM tip is removed from the liquid membrane, the back of the
CaSM is vented to atmospheric pressure, and the CaSM is removed from its holder.
III. Properties of CaSMs
A. Response to Ion Activity
The potential across the Ca2þ-selective liquid membrane in the tip of the CaSM
is comprised of two phase boundary potentials, between the interfaces of the liquid
membrane with (1) the backfilling solution and (2) the extracellular medium, and
Loading pipette
A B
Loading pipetteCaSM CaSM
TPPS
Fig. 1 Ca2þ-selective microelectrode tip filling station. (A) Micropositioners on each side of an
upright microscope are used to position the tips of a loading pipette and a CaSM in the field of view.
The stage has been removed. The threaded plunger (TP) syringe and the plastic syringe (PS) are
connected to the loading pipette and CaSM via plastic tubing enabling application of pressure and
suction to control the length of liquid membrane loaded into the CaSM from the loading pipette.
(B) Higher magnification of A showing the close positioning of the glass loading pipette and glass
CaSM. The system is mounted on a large metal plate to reduce vibration during loading.
4. Electrochemical Measurement of Ca2+ Flux 95
also the diVusion potential between the two ends of the column of liquid mem-
brane (Bakker et al., 1997). The diVusion potential is considered negligible as bulk
movement of Ca2þ across the liquid membrane does not occur during common
usage with high-impedance electronics and no current flow. The inner phase
boundary potential is considered constant due to the rigorous clamping of Ca2þ
with buVers or with high concentrations of Ca2þ in the backfilling electrolyte. The
external phase boundary potential, for an ideal ion-selective microelectrode, is
related to the extracellular ion activity by the Nernst equation,
E ¼ EO þ S logai ð1Þwhere ‘‘Eo’’ is the sumof constant potential contributions, ‘S’is theNernstian slope¼(2.3RT) / (ziF) (R, T, and F hold their standardized meanings) and ‘‘ai’’ is the activity
of the primary ion. Constant potential contributions are comprised of the boundary
potentials and liquid junction potentials that exist across the circuit comprising the
reference and measuring electrodes. The valence ‘‘zi’’ of Ca2þ produces a slope only
half as steep (�29 mV/order magnitude change in Ca2þ) compared to monovalent
ions. The high selectivity of the twoCa2þ liquidmembranes discussed here alongwith
the generally standard physiological media that are used enables us to perform flux
96 Mark A. Messerli and Peter J. S. Smith
calculations according to the Nernst equation listed above. However, in complex
media with significantly interfering ions, the slope of response can be reduced. The
decrease in slope can be predicted using the Nicolsky–Eisenman equation for ions of
similar valence and the extended Nicolsky–Eisenman equations for ions of diVerentvalence (Bakker et al., 1994). In some cases it may be more practical to perform an
empirical determination of the slope of the line describing the relationship between
measured voltage and the change in ionic activity. This determination is performed
by making up the working medium with slightly higher and lower concentrations of
Ca2þ and determining the slope of response. A sub-Nernstian response may reflect
the presence of an interfering ion or of a substance that is fouling the microelectrode.
According to the Nernst equation, the voltage output is dependent on ionic
activity. However, as ion activity is directly proportional to ion concentration, via
the activity coeYcient, and the changes that occur to the activity coeYcient due to
changes in ionic strength are negligible during self-referencing in physiological
saline, we will use concentration in place of activity for further discussion.
B. Selectivity
A primary motivation for early development of CaSMs was to monitor intracel-
lular [Ca2þ] (Lanter et al., 1982; Tsien and Rink, 1981). This required a liquid
membrane with high selectivity for monitoring the low resting [Ca2þ]i (�100 nM)
in the presence of higher concentrations of potentially interfering ions including
Kþ (�120 mM), Naþ (�10 mM) and Mg2þ (�1 mM). Accordingly, two diVerentCa2þ ionophores with very high selectivity were reported (Ammann et al., 1975,
1987; Lanter et al., 1982). Some of their selectivity coeYcients for Ca2þ over other
common cations are listed in Table I. Selectivity for Ca2þ over these cations is
relatively good compared to liquid membranes for other ions. However, not all
inorganic or organic ions have been tested and may therefore act as interferents.
Not only do interfering ions reduce the electrode’s voltage response to the primary
ion but they also slow the response time of the electrode (Bakker et al., 1997). This
point is particularly important when using the electrodes in self-referencing mode
where a temporal component is part of the modulation approach. In biological
applications, it is critical for the investigator to empirically test the voltage
response of a CaSM in the medium in which the experiments are to be performed.
Simple solutions of the primary ion are not suYcient. Additionally, the CaSM
should be tested for interference or fouling due to the addition of transport
blockers or cellular poisons.
C. Spatial Resolution
Small, micron-sized sensors give rise to high spatial resolution. The spatial
resolution is defined first by the external surface area of the Ca2þ-selective liquid
membrane, but also by the sampling time and the distance between the source of
the Ca2þ transport and the CaSM. This holds true for the high-impedance
Table ISelectivity coeYcients of two diVerent Ca2þ-selective liquidmembranes
Interfering ion (M)
Selectivity coeYcients (log CaMPot)
ETH1001 ETH129a
Kþ �5.4b �7.2
Naþ �5.5b �5.8
Mg2þ �4.9b �6.7
NH4þ �5.0c �3.6
Hþ �4.4c �2.5
aAmmann et al. (1987).bLanter et al. (1982).cAmmann et al. (1975).
4. Electrochemical Measurement of Ca2+ Flux 97
headstages �1015 Ω that are typically used, which help to decrease the bulk
movement of Ca2þ between the medium and liquid membrane. Spatial resolution
is decreased due to diVusion of Ca2þ in the bulk medium from nearby transport
events. DiVusion of Ca2þ from 10 to 20 mm away will reach the CaSM in only �20
and �80 ms, indicating that the sampled volume is much larger than the immedi-
ate dimensions of the CaSM tip. As these events are diVusing from regions further
away, the local concentration change that they produce near the tip of the CaSM
will be much smaller (proportional to 1/r2) than the signals from events immedi-
ately in front of the CaSM. The decay in signal with distance is evident from
measurements of extracellular Kþ gradients due to eZux through single Kþ
channels (Messerli et al., 2009). The sampling domain of the CaSM is therefore
slightly larger than the surface area of the liquid membrane and decays rapidly
with increasing distance from the surface.
D. Response Time
Self-referencing of CaSMs requires the use of CaSMs with relatively short
response times so that the CaSM can reach equilibrium in a short period of time
at its new position. The response time of CaSMs is governed by the ability to
provide charge to the sensing node. In an ideal measuring system, diVusionthrough the unstirred layer at the surface of the electrode defines the response
time of the sensors when the liquid membrane is equilibrated with the salt of an ion
to which the electrode responds (Bakker et al., 1997). For ion-selective microelec-
trodes, this process may occur so quickly that the electronics of the system slow the
measured response (Ammann, 1986). Low input impedance of the amplifier and
parasitic capacitances in the circuit will draw more charge than an ideal system
therefore slowing the response time of the system. Amplifier input impedances of
Table IICaSMs based on ionophore ETH1001 possess short response times inphysiological saline over a range of [Ca2þ]
Response times (t95%ms)
0.1–1 mM 1–10 mM 10–1 mM 1–0.1 mM
48�7 53�10 58�9 81�10
Physiological saline consists of (in micromolar) 120 NaCl, 5 KCl, 2 MgCl2,
10 HEPES with the CaCl2 concentration listed above. CaSMs remained stationary
during the experiment, while three adjacent streams of media (1 mL/min) were rapidly
positioned (<8 ms) in front of the measuring electrode. These measurements describe
the response time of the entire measuring system for 4 CaSMs.
98 Mark A. Messerli and Peter J. S. Smith
�106 GO are typically used to accommodate ion-selective microelectrodes that
have high resistances, 1–20 GO (Ammann, 1986). Even with the best electronics,
the time constant (RC, resistance times capacitance) of the CaSM itself imposes a
low pass filter. Resistance is primarily dependent on the tip diameter and length of
the column of liquid membrane and capacitance is primarily dependent on the
thickness and dielectric constant of the wall of the glass micropipette. To reduce
the resistance of the CaSMs, they are fabricated with relatively large tips of 2–3 mminner diameter and with short columns of liquid membrane �30 mm. The short
columns are achieved by tip loading the liquid membrane as discussed above.
Capacitance can be lowered by using thicker walled borosilicate glass (1.5 mm O.
D. 0.84 mm I.D. cat# 1B150-6, WPI Sarasota, FL). The construction design listed
above has produced CaSMs with response times shorter than 100 ms, Table II.
In practice, a slight deviation from the expected length does not change the
response time very much, at least when considering the use of these electrodes in
the self-referencing application.
IV. Self-referencing of CaSMs
A. DiVerential Concentration Measurement
Self-referencing of CaSMs was developed to measure extracellular Ca2þ gradi-
ents/currents that may have existed near previously characterized extracellular
voltage gradients (Kuhtreiber and JaVe, 1990). For example, relatively steady
eZux of Ca2þ across the plasma membrane gives rise to a gradient of Ca2þ with
a higher concentration near the cell. Self-referencing of CaSMs is implemented by
measuring the [Ca2þ] at two points in that Ca2þ gradient. The electrical variation
of a single CaSM due to thermal noise, �100–200 mV, of the high-impedance
sensors and chemical drift is too large to enable measurement of such small
diVerences in extracellular Ca2þ. As a result, a frequency sensitive method of
4. Electrochemical Measurement of Ca2+ Flux 99
detection was explored based on response times of about 1 s reported for CaSMs
commonly used at that time (Ammann, 1986). The general measuring protocol
includes intermittent collection of ion concentration by a single CaSM at a position
near the biological preparation and then at a position some distance away orthog-
onal to the source. A CaSM is shown in Fig. 2A and B at the two positions, next to
a mouse pancreatic islet. The excursion distance in this case is 20 mm but can vary
between 5 and 50 mm depending on the size of the cell or cellular preparation.
At each pole of excursion, the CaSM is allowed to reach equilibrium (�0.25 s)
before recording the average local ion concentration for about 1 s. Considering
that the CaSMs possess response times of less than 0.1 s, 0.25 s is plenty of time to
reach equilibrium. The CaSM is then immediately positioned to the opposite pole,
and allowed to reach equilibrium before recording the local ion concentration. The
movement of the CaSM between the two positions is controlled by stepper motors
set to move the sensor at a rate of 40 mm/s such that it takes 0.25 s to reach its new
Near pole (E1)
A
Near pole
Far poleE2 E1
ΔE = E1− E2
ΔE
E2 E1d d d d
C
B
Far pole (E2)
50mm
Fig. 2 Ca2þ flux measurements performed with self-referencing of a Ca2þ-selective microelectrode
near a mouse pancreatic islet. (A) In the near pole the CaSM collects the average [Ca2þ]-dependentpotential for 1 s, E1. (B) After movement to the far pole and equilibration, the average [Ca2þ]-dependentpotential is collected again for 1 s, E2. (C) Data collection scheme portraying Ca2þ eZux. The auto-
mated determination of the diVerential [Ca2þ]-dependent potential, DE, is used to determine Ca2þ flux.
This measuring scheme continues, as defined by the user. The [Ca2þ]-dependent potential is discarded,(d), during periods of movement (�0.25 s) and during equilibration in the new position (�0.25 s).
100 Mark A. Messerli and Peter J. S. Smith
position. A diVerential concentration recording between the two poles of excursion
is collected, about every 3 s. The measurement scheme continues until stopped by
the user. The diVerential recording possesses peak to peak noise of �10 mV while
longer periods of signal collection and averaging can enable extraction of concen-
tration diVerences that give rise to 1 mV diVerences or a 0.008% diVerence from the
background [Ca2þ]. Figure 2C illustrates this collection scheme during Ca2þ eZux
where E1 and E2 are the recorded ion concentration–dependent potentials at the
two poles. The recording collected during movement and equilibration at the new
pole is discarded, labeled ‘‘d’’ in Fig. 2C. A diVerential Ca2þ measurement is
collected over a period of about 3 s, which is faster than the low frequency drift,
thus reducing its influence on the measurement. Signal averaging at each pole over
a period of 1 s reduces the influence of the high frequency noise. Measurement of
the diVerential ion concentration–dependent voltage, DE, between the two posi-
tions over time enables further enhancement of the signal-to-noise ratio.
This modulation approach was termed self-referencing, referring to the fact that
the measurements, collected by a single CaSM, are compared to each other in
order to determine the concentration diVerence between the two points. The signal
collected by a single CaSM at one point in space and time is referenced to the
signal collected by that same CaSM at a diVerent point in space and time in order
to reduce electrical drift of the measuring system. The CaSM has its own
bath reference electrode. While this diVerential measurement could be achieved
with two similar, CaSMs, positioned at known distances from the source, the
sensitivity would suVer from the signal drift and noise inherent to two separate
measuring systems.
Measured diVerential voltages of �10s mV are extracted from relatively large
oVset potentials�100s mV by using a combination of amplification methods. Low
gain must be used with the large oVset potentials in order to keep the signal within
the dynamic range of the amplifier. As a result, low resolution digital systems will
not be able to register small changes in the diVerential voltage. A 12-bit system
with a dynamic range of �10 V provides only 4.9 mV/bit resolution while a 16-bit
system provides only 0.3 mV/bit. Additional amplification prior to digitization is
necessary to resolve signals at or below 1 mV. Two separate methods for amplifica-
tion are used; (1) a nearly equal and opposite electrical oVset is supplied before
amplification (sample hold mode) and (2) a running average of the low gain
measurement is subtracted from the real-time input before amplification (RC
subtract mode). Sample hold mode applies a known voltage that is selected either
manually or automatically from the signal after a set duration of time to null the
oVset potential before applying 103 times gain. The primary disadvantage for this
mode is that drift can take the system back out of the dynamic range of the
amplifier so that a new potential must be applied regularly. The advantage is
that it does not need an additional correction factor to compensate for the signal
lost due to the filtering that occurs in RC subtract mode. In RC subtract mode, a
high-pass filter is used to collect a running average potential that is subtracted
from the potentials collected in the near and far pole. The signals are then amplified
4. Electrochemical Measurement of Ca2+ Flux 101
103 times before digitizing. Typically, this mode employs a high-pass filter with a
time constant of 10 s. RC subtract allows amplification for systems with large drift
but involves a correction factor to oVset the high-pass filter. The correction factor
will be dependent on the time constant of the high-pass filter and the period of data
acquisition. For standard conditions, a period of 3.3 s (0.3 Hz translation frequen-
cy), 40 mm/s translation speed, 10 s time constant of the high-pass filter along with
data selection of the last 70% of the half cycle, we calculate that the signal is 7%
smaller than a square wave with similar rise time.
Automated, repetitive positioning of the CaSMs is controlled by three stepper
motors arranged in an X, Y, Z configuration with the Z plane parallel to the plane
of the stage of the microscope. Smooth linear motion is obtained by coupling each
of the stepper motors to a lead screw controlling the position of three small
translational plates connected together to form a three-dimensional positioner
(BioCurrents Research Center, Woods Hole, MA). Low voltage control of the
stepper motors prevents electrical feedback to the high-impedance headstage of the
CaSM. Positioning can be achieved over a working distance of 3–4 cm with
submicron resolution and repeatability (Danuser, 1999). A computer interface
enables repetitive motion and positional control with the Faraday cage closed.
B. DiVerential Concentration Determination
The relationship between the measured diVerential voltage and the diVerentialion concentration between the two poles of excursion for an ideal CaSM can be
determined using the Nernst equation.
E1 � E2 ¼ EO þ S logCið Þ1 � EO þ S logCið Þ2DE ¼ S logCið Þ1 � S logCið Þ2DE ¼ logCS1 � logCS2
i 1ð Þ i 2ð ÞCS1
i 1ð Þ !
ð2Þ
DE ¼ logCS2
i 2ð Þ
‘‘E1’’, ‘‘Ci(1)’’, and ‘‘S1’’ are the measured voltage, [Ca2þ] and slope of the voltage–
log(Ci) graph for the near pole of excursion. The subscript 2 labels the same
parameters for the far pole of excursion. The slow changing constant potential
contributions ‘‘Eo’’ are reduced if not eliminated by calculating the diVerencebetween potentials over short periods of time.
Equation (2) enables a clear picture of the relationship of the sensitivity of
detection to the background ion concentration during measurements. For a
given [Ca2þ] change due to cellular flux, the concentration in the position next to
the cell, Ci(1), is the sum of the background ion concentration and the concentra-
tion change generated by the source while Ci(2), in most cases, is close to the
background ion concentration. It is easier to generate a larger DE when the
background concentration of the measured ion is lower as the ratio of Ci(1)/Ci(2)
102 Mark A. Messerli and Peter J. S. Smith
will be much larger/smaller for the same Ca2þ eZux/influx on lowered background
[Ca2þ]. This has led to the lowering of the background [Ca2þ] in order to generate
DE with a greater signal-to-noise ratio, see Table IV. Care must be taken to ensure
that changing the background concentration does not interfere with normal cellu-
lar activity.
Rearrangement of Eq. (2) relates the [Ca2þ] in the near pole of excursion to the
[Ca2þ] at the far pole of excursion.
Cið1Þ ¼ CS2S1
ið2Þ � 10DES1 ð3Þ
For an ideal electrode, the voltage output is close to the Nernstian slope over a
wide range of [Ca2þ] so S1 ¼ S2 ¼ S ¼ 2:3RTziF
. Therefore, Eq. (3) simplifies to
Ci 1ð Þ ¼ Ci 2ð Þ � 10DES ð4Þ
For minute fluxes that are typically measured with self-referencing, the average
concentration of Ca2þ at the far pole, position 2, is not too diVerent from the
average concentration of Ca2þ in the bulk solution. Therefore the diVerence in
[Ca2þ] between the two points of excursion can be described as follows:
DC ¼ Ci 1ð Þ � Ci 2ð Þ ¼ Cbath � 10DES � Cbath ð5Þ
A primary assumption here is that the excursion distance is small compared to the
extent that the gradient extends out into the bulk solution so that the concentration
diVerence between the two excursion points is linear. For minute gradients
measured from small cells (�10 mm diameter), an excursion of 10 mm will most
likely sample over a distance in which the concentration diVerence is not linear andtherefore will lead the investigator to underestimate the true flux. Incorrect estima-
tion of the true flux could also occur during a two-point measurement in a more
intense, extended gradient, where the concentration of the ion in the far pole is
substantially diVerent from the background concentration of the ion. In both of
these cases, a three-point measurement should be performed in order to (1) more
carefully map the concentration gradient with a third point to ensure a linear
relationship or determine a more accurate nonlinear relationship and (2) to deter-
mine the concentrations in the gradient relative to the background concentration
of the ion in the bath.
The selectivity of Ca2þ liquid membranes is relatively good compared to liquid
membranes for other ions. Therefore, measurement of Ca2þ gradients in the
presence of a constant concentration of an interfering ion or in the presence of
a gradient of an interfering ion is not a major concern. However, specific
circumstances may require the use of higher concentration of an interfering
ion and the two cases need to be addressed. Details necessary to account for
these situations have been addressed previously (Messerli et al., 2006; Smith
et al., 2007).
4. Electrochemical Measurement of Ca2+ Flux 103
C. Calculation of Flux
The diVerential concentration measurement is converted to flux to provide a
direct representation of the number of ions passing through a unit area per unit
time. Calculation of flux enables comparison of Ca2þ transport between diVerentsystems as it takes into account the diVusion coeYcient of Ca2þ, the distance overwhich the diVerential concentration measurement was acquired, the surface geom-
etry of the source, and the distance of measurement from the source. It also
provides a value for comparison of Ca2þ flux measured with self-referencing of
CaSMs to other methods for monitoring Ca2þ including intracellular fluorescent
and luminescent ion indicators and radioactive tracer flux studies. For planar
sources where the measuring electrode is relatively close to a large source, such
as a tissue, sheet of cells or large diameter cell, and the diVerential concentrationis measured over a small distance ‘‘Dx’’ within the gradient next to the source,
flux (J) is
J ¼ �DDCDx
ð6Þ
where ’’D’’ is the diVusion coeYcient of Ca2þ. By this model, at equilibrium the
flux measured at some distance from the source is the same as the flux at the surface
of the source. According to this equation, eZux, a higher concentration of Ca2þ
near the source, is identified by a negative flux.
In order to determine flux at the cell surface for known surface geometries, it is
useful to calculate analyte flow, that is, the quantity of substance (Q) moving per
unit time (Henriksen et al., 1992). Flow is the same for all concentric surfaces
surrounding the source surface. Flux at the source surface is the flow divided by the
surface area of the source. Therefore, radially emanating flow from a cylindrical
surface is
Flow ¼ Q
t¼ � 2pD
ln b=að Þ DCð Þ ð7Þ
where ‘‘D’’ is the diVusion coeYcient of the analyte and ‘‘a’’ and ‘‘b’’ are the radial
distances between the center of the cylinder and the electrode tip at the near and far
poles, respectively. These equations have been adapted from Crank (1967). Ana-
lyte flux at the surface of the cylinder is then determined by dividing by its surface
area 2prl. A caveat of this approach is the assumption that the flow is equal at all
points around the cylinder and along the shaft of the cylinder. An alternative is to
calculate flux per unit length by dividing by 2pr (Henriksen et al., 1992).
The flow from a spherical source is
Flow ¼ Q
t¼ �4pD
ab
b� aDCð Þ ð8Þ
Flux at the cell surface can then be determined by dividing by the spherical surface
area 4pr2.
104 Mark A. Messerli and Peter J. S. Smith
D. Correction for Ca2þ BuVering
The presence of Ca2þ buVers or binding agents with the appropriate aYnity can
lead to collapse of Ca2þ gradients by shuttle buVering (Speksnijder et al., 1989).
Ca2þ can diVuse from the surface of the cell in either its free state or bound to the
buVer. CaSMs only measure the free concentration of Ca2þ. The actual Ca2þ
flux at a source is the sum of the measured free Ca2þ flux and the unmeasured
Ca2þ flux diVusing as Ca2þ bound to buVer.
JCa total ¼ JCa measured þ JCa Buffer ð9ÞKnowing the conditions under which the Ca2þ flux was measured including the
[Ca2þ] of medium, dissociation constant, Kd, of the buVers and concentration of
the buVers present, a simple relationship can be derived to determine the ratio of
Ca2þ diVusing bound to buVer compared to the freely diVusing Ca2þ. ShuttlebuVering of Hþ is a bigger concern than for Ca2þ due to the larger number of
Hþ buVers that are used in physiological media. The equations that exist to correct
for shuttle buVering of Hþ (Messerli et al., 2006; Smith et al., 2007) can be adapted
for use with Ca2þ flux correction and may be necessary under specific
circumstances.
xi ¼ DB
DCa2þB½ � Kd
Kd þ Ca2þ� �� �2 ð10Þ
The correction factor, ‘‘xi’’, is the ratio of the Ca2þ bound buVer flux to the free
Ca2þ flux. Therefore
JCa total ¼ JCa measured 1þ xi þ þ xnð Þ ð11Þwhere a number of diVerent Ca2þ buVers (xi þ þ xn) may be collapsing the Ca2þ
gradient. The correction factor is based on three criteria, the ratio of the diVusioncoeYcients of the Ca2þ-buVer complex to free Ca2þ, the Ca2þ buVer concentration,and the dissociation constant, Kd, of the Ca
2þ buVer compared to the [Ca2þ] of themedium. The Kd is the inverse of the more commonly used ‘‘stability constant’’ also
known as the association constant. Only Ca2þ buVers/binding agents that have Kd
values near the range of the extracellular [Ca2þ] will act as eVective shuttle buVersduring self-referencing of CaSMs. Table III lists a few of these compounds. Note
that two of the compounds, ADA and Bicine, are commonly used as Hþ buVers.Generally, the Ca2þ Kd values of other Good buVers are not in the range of normal
extracellular [Ca2þ] or are very poor Ca2þ chelators (Dawson et al., 1986).
E. Measurement of Voltage Gradients
The use of CaSMs with self-referencing is subject to a similar problem that
occurs with the use of intracellular CaSMs; specifically they detect not only
changes in ion concentration but also voltage. Extracellular voltage gradients
Table IIIList of Ca2þ binding compounds that can act as shuttle buVers for extra-cellular Ca2þ in the 0.1–1.0 mM range
Ligand log(Kd)
Pyrophosphate �5.0
N-(2-acetamido)iminodiacetic acid (ADA) �4.01
ATP �3.8
Citric acid �3.5
Oxalic acid �3.0
Polyphosphate �3.0
N,N-bis(hydroxyethyl)-glycine (Bicine) �2.8
Values were obtained fromDawson et al. (1986) except as noted 1(Lance et al., 1983).
4. Electrochemical Measurement of Ca2+ Flux 105
have been mapped near many diVerent systems (Borgens et al., 1989; Nuccitelli,
1986) Extracellular electric fields generated by cells are generally very small espe-
cially in high conductivity media such as animal saline. However, in lower conduc-
tivity saline ion transport can give rise to relatively large electric fields >1 mV/10 mm which can be detected with self-referencing microelectrodes. Plants for
example, drive transcellular currents through them, as a result of ion transport
across single cells or tissues. In low conductivity medium, these currents generate
substantial voltage gradients next to the cells, coexisting with the concentration
gradients of the transported ions. The diVerential voltage measured by the CaSM
will be the sum of the voltage diVerences due to the [Ca2þ] diVerence and the
voltage diVerence. For example, a peak voltage diVerence during oscillating cur-
rent influx of about 9 mV would occur over a 10-mm distance immediately in front
of a lily pollen tube. Peak current density around 0.4 mA/cm2 was measured at a
distance of about 20 mm from the cell surface with a medium resistivity of about
5000 O cm (Messerli and Robinson, 1998). This voltage diVerence is just above thebackground noise of the system used at that time,�5 mV for Ca2þ, (Messerli et al.,
1999). The voltage signals detected by the self-referencing CaSM peaked about six
times larger than the diVerential voltage due to current flux indicating that the
extracellular electric field could have contributed to the calculated Ca2þ flux by up
to 15%.
F. Positional Artifacts
Self-referencing of CaSMs near solid objects can generate position dependent
artifacts. Movement of Ca2þ across the external interface between the liquid
membrane and bathing medium may occur through current driven and zero net
current mechanisms (Bakker and MeyerhoV, 2000). Release of Ca2þ by the CaSM
restricts its sensitivity in bulk medium by leading to a modification of the local ion
106 Mark A. Messerli and Peter J. S. Smith
concentration at the tip of the microelectrode. During self-referencing, when the
CaSM is positioned near a solid object, released Ca2þ can accumulate between the
CaSM and the object in a short period of time leading to an artificially higher
concentration of Ca2þ in the constrained space. Likewise, uptake of Ca2þ by the
CaSM can lead to a depletion of Ca2þ in the constrained space. These artifacts are
most apparent in solutions of low background [Ca2þ]. Figure 3 shows examples of
both extremes where CaSMs are self-referenced near a 100-mm diameter glass
bead. The CaSM is moving in a path so that the plane of its tip is always parallel
to the near surface of the bead to enable the ISM to get closer to the surface. The
ion trapping eVect is reduced when the path of excursion orients the plane of the tip
of the CaSM perpendicular to the near surface of the solid object, as shown in
Fig. 2A and B, because the liquid membrane surface cannot get as close to the solid
object. EZux of Ca2þ from the microelectrode tip occurs when constructed with
100 mM CaCl2 backfilling solution, originally performed by Kuhtreiber and JaVe(1990). Accumulation of the released Ca2þ in less than 1 s can be detected when the
bath [Ca2þ] is 50 mM but not when it is 2 mM, giving rise to an artificial eZux of
Ca2þ from the solid glass bead. Reducing the concentration of the primary ion in
the backfilling solution is one method of reducing the ion leak (Bakker and
MeyerhoV, 2000). However, when used with self-referencing this can lead to an
artifact of the opposite polarity shown in Fig. 3. The CaSM constructed with
−40
−30
−20
−10
0
10
20
30
40
50
0 5 10 15 20 25 30
Distance from bead (mm)
Diff
eren
tial v
olta
ge (mV
)
100 mM Ca2+ backfill, 50mM Ca2+ bath
100 mM Ca2+ backfill, 2 mM Ca2+ bath
100 nM Ca2+ backfill, 50mM Ca2+ bath
100 nM Ca2+ backfill, 2 mM Ca2+ bath
50mM Ca2+ backfill, 50mM Ca2+ bath
Fig. 3 Ca2þ movement across the tip of a CaSM can be detected in low background [Ca2þ] near asolid object. Electroneutral exchange of Ca2þ out of the tip of a CaSM (filled box) or into the tip of the
CaSM (filled circle) can give rise to accumulation or depletion of the local [Ca2þ] between a solid object
and the tip of the CaSM. In higher bath [Ca2þ] (empty box, empty circle) the accumulation or depletion
is insignificant compared to the background [Ca2þ] and is therefore not detected. In lowered bath [Ca2þ]careful balancing of the backfilling [Ca2þ] with the bath [Ca2þ] can reduce (filled triangle) if not
eliminate net movement of Ca2þ across the liquid membrane.
4. Electrochemical Measurement of Ca2+ Flux 107
100 nM Ca2þ in the backfilling solution generates an influx of Ca2þ into the CaSM
tip, depleting the local [Ca2þ] in the bath and giving rise to an artificial Ca2þ influx
into the glass bead. Again this can be detected in the 50 mM Ca2þ solution but not
the 2 mM Ca2þ containing bath solution. The artifact can be reduced by matching
the backfilling [Ca2þ] with the [Ca2þ] in the bath. Other methods for eliminating
Ca2þ flux across the tip of the CaSM include current clamping (Lindner et al.,
1999; Pergel et al., 2001), or using the solid contact ion-selective electrode design
(Lindner and Gyurcsanyi, 2009).
V. Ca2þ Flux Measurements
Extracellular Ca2þ flux measurements have been performed on a number of
diVerent systems some of which are listed in Table IV, ranging from animal
neurons and muscle to tip growing root hairs, pollen tubes, and fungi. Measured
Ca2þ fluxes are relatively small ranging between 0.1 and 10 pmol cm�2 s�1 encour-
aging measurements from cells in reduced background [Ca2þ] 0.1 mM. The limit
of flux sensitivity for a typical self-referencing CaSM with �10 mV near real-time
variation performed in 1 mM bath [Ca2þ] is about �6.3 pmol cm�2 s�1, an order
of magnitude higher than in 0.1 mM bath [Ca2þ]. Considering the large trans-
plasma membrane electrochemical driving force on Ca2þ, reduction of extra-
cellular [Ca2þ] by an order of magnitude did not cause noticeable problems
for the diVerent preparations, at least over the few hour period during
which measurements were acquired as noted by multiple authors listed in
Table IV.
While eZux of Ca2þ in cells at rest is expected to be relatively small, the
measured influx of Ca2þ, presumably through channels, is also relatively small.
Active single 0.5 pS Ca2þ channels at a density of 1 mm�2 should give rise to a Ca2þ
influx of about 47 pmol cm�2 s�1. Although as noted by Hille (2001) voltage-gated
Ca2þ channels exist at low density and low open probabilities (<0.1) even with
strong depolarizing potentials indicating that low channel density and activity is
suYcient to account for measured changes in [Ca2þ]i. The channel density and
activity used above may be overestimates of actual Ca2þ channel density. Also,
weak influx may also be a result of the Ca2þ amplification cascades that exist to
release Ca2þ from intracellular stores after influx through the plasma membrane.
Additional directions for the use of self-referencing with CaSMs include the
study of electroneutral Ca2þ transporters/exchangers and extracellular Ca2þ sig-
naling (Breitwieser, 2008). Ca2þ selective microelectrodes have been instrumental
in providing the sensitivity for defining the complex transport of the Naþ/Ca2þ
exchanger (Kang and Hilgemann, 2004) and the P-type plasma membrane Ca2þ
pump (PMCA) in neurons (Thomas, 2009). With self-referencing of ion-selective
microelectrodes, transport stoichiometries could be determined noninvasively
from the outside of intact cells.
Table IVCalcium flux measurements acquired from preparations representing multiple Kingdoms
Preparation Ca2þ flux (pmol cm�2 s�1) Conditions
Bath [Ca2þ](mM) Reference
Aplysia californica bag cell �1 to �5 Rest, H2O2 0.1 Duthie et al. (1994)
�1 to �5 Rest, thapsigargin None added
(0.5 mM
EGTA)
Knox et al. (1996)
Rana catesbeiana hair cell �0.5 Rest 0.05 Yamoah et al. (1998)
þ5.0 Stimulated
Callinectes sapidus olfaction �2.5 15% ASWa 0.1 Gleeson
et al. (2000b)
�4.0 AFWb 0.1 Gleeson
et al. (2000a)
Sclerodactyla
briareus smooth muscle
�1.0 to �4.0 Rest, Ach.c 0.1 Devlin and Smith
(1996)
�1 to �7.5 Muscarinic
agonists
0.1 Devlin et al. (2003)
Busycon canaliculatum
cardiac muscle
�1 to �4 Rest, FMRFamide 0.1 Devlin (1996)
Mouse ova �0.02 Rest 0.05 Pepperell
et al. (1999)þ0.6 Bepridil additiond
�0.2 Replenished Naþ
þ0.08 to þ0.35 Addition of EGFe None added Hill et al. (1999)
Lilium longiflorum
pollen tubes
þ2 to þ20 Germination 0.1 Pierson et al. (1994)
þ5 to þ38f Oscillating
tip growth
0.13 Messerli et al. (1999)
Root hairs þ4.3 to þ7.2 (alfalfa) Tip growth,
nod factor
Not listed Herrmann and Felle
(1995)
þ2.5 (S. alba) Tip growth 0.1
þ0.07 to þ1.2 (A. thaliana) Osmotic regulation 0.1 Lew (1998)
Neurospora crassa hyphae �0.1 Voltage
dependence
0.05 Lew (2007)
þ0.1 to þ1.5 Osmotic regulation 0.05 Lew and Levina
(2007)
Ceratopteris richardii spores �3.5 top Gravity sensing 0.02–0.05 Chatterjee
et al. (2000)þ0.5 bottom
Physcomitrella
patens filaments
þ1–3 Gravity sensing 0.1 Allen et al. (2003)
aArtificial seawater.bArtificial freshwater.cAcetylcholine.dBepridil was added to block the plasma membrane Naþ/Ca2þ exchanger.eEpidermal growth factor.fCalculated flux at cell surface.
108 Mark A. Messerli and Peter J. S. Smith
Acknowledgments
The BioCurrents Research Center is funded by NIH:NCRR grant P41 RR001395.
4. Electrochemical Measurement of Ca2+ Flux 109
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CHAPTER 5
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
Practical Aspects of Measuring IntracellularCalcium Signals with Fluorescent Indicators
Joseph P. Y. Kao, Gong Li, and Darryl A. AustonCenter for Biomedical Engineering and Technology, andDepartment of PhysiologyUniversity of Maryland School of MedicineBaltimore, Maryland, USA
OGY,All rig
A
Vhts
bstract
OL. 99 0091reserved. 113 DOI: 10.1016/S0091
-679X-679X
I.
I ntroduction II. F luorescent Ca2þ Indicators III. L oading Indicators into CellsA.
Limited Aqueous Solubility of AM Esters B. Dye Compartmentalization: Loading of Indicator into SubcellularCompartments Other than the Cytosol
(
C.
Dye Leakage or Extrusion from Cells D. Procedure for LoadingIV.
M anipulation of [Ca2þ] A. Using EGTA and BAPTA as Extracellular Ca2þ BuVers B. Lowering Extracellular [Ca2þ] C. Divalent Cation Ionophores D. BuVering Changes in Intracellular [Ca2þ]V.
C onversion of Indicator Fluorescence Signal into Values of [Ca2þ] A. Calibrating a Nonratiometric Fluorescent Indicator B. Calibrating a Ratiometric Fluorescent IndicatorVI.
R eporting Indicator Fluorescence Intensity Changes without Calibration A. Reporting Relative Changes in Fluorescence: F/F0 and DF/F0 B. Caveat in Interpreting Relative Fluorescence Changes: IndicatorFluorescence is Not a Linear Function of [Ca2þ]
V II. M easuring [Ca2þ] in MitochondriaA.
Estimating the Fraction of Intracellular Rhod-2 Indicator that Resides inMitochondriaB.
Minimizing Rhod-2 Loading in the Cytosol C. Monitoring Cytosolic and Mitochondrial [Ca2þ] Simultaneously/10 $35.0010)99005-5
114 Joseph P. Y. Kao et al.
V
III.1
conc2
while
emis
excit
over
C
Sy
en
A
e
sio
at
a
oncluding Remarks
mbols used: Ca2þ, free calcium ion; [Ca2þ], concentration of free calcium ions; [Ca
tration of free calcium ions.
n excitation spectrum is taken by monitoring fluorescence emission intensity at a fixe
xcitation light is scanned through a wavelength range over which the sample can abso
n intensity is plotted as a function of the excitation wavelength. To collect an emissi
ion light at a fixed wavelength is delivered to the sample while the emission intensity
wavelength range. Here, the emission intensity is plotted as a function of emission w
2þ]i,
d wa
rb l
on s
is m
avel
R
eferencesAbstract
The use of fluorescent indicators for monitoring calcium (Ca2þ) signals and for
measuring Ca2þ concentration ([Ca2þ]) in living cells is described. The following
topics are covered in detail: (1) ratiometric and nonratiometric fluorescent indicators
and the principles underlying their use, (2) techniques for loadingCa2þ indicators and
Ca2þ buffers into living cells, (3) calibration of indicator fluorescence intensity
measurements to yield values of intracellular [Ca2þ], (4) analysis of nonratiometric
fluorescence intensity data and caveats relating to their interpretation, (5) techniques
for manipulating intracellular and extracellular [Ca2þ], and (6) the use of fluorescent
indicators tomonitorCa2þ signals inmitochondria. The chapter aims topresent these
fundamental topics in a manner that is practically useful and intuitively accessible.
The origins of key mathematical equations used in the article are outlined in two
appendices.
I. Introduction
In the application of anymeasurement technique, a body of practical knowledge is
shared by experienced practitioners. Although important for making successful
measurements, such lore, which sometimes seems arcane, often is not described
explicitly or explained in journal publications. In this respect, measuring [Ca2þ]1
with fluorescent indicators is no exception. The purpose of this chapter is to gather in
one place some of the most common and useful practical information relevant to the
use of fluorescent Ca2þ indicators. Such a collection of information is hoped to
alleviate the frustration of those who are novices at using fluorescent indicators.
II. Fluorescent Ca2þ Indicators
The commonly available fluorescent indicators for Ca2þ fall into two operation-
al classes: dual-wavelength ratiometric dyes and single-wavelength nonratiometric
dyes (Table I). Chemical structures of some of the indicators listed in Table I are
shown in Fig. 1. For nonratiometric indicators, a change in [Ca2þ] brings about acorresponding change in the intensity of the indicator’s fluorescence excitation and
emission spectra,2 whereas the wavelengths of the excitation and emission spectral
cytosolic
velength
ight. The
pectrum,
onitored
ength.
Table IProperties of common fluorescent Ca2þ indicatorsa
Indicator type Kd (nM)
Absorption maxima
(nm) Emission maxima (nm)
Ca2þ-free Ca2þ-bound Ca2þ-free Ca2þ-bound
Nonratiometric
Monomeric
Quin2 115b 352 332 492 498
Fluo-2/Fluo-8c 380 – 492 – 514
Fluo-3 400 503 506 526 526
Fluo-4 345 491 494 �516 516
Calcium Green-1TM 190 506 506 531 531
Calcium Green-2TM 550 506 503 536 536
Calcium Green-5NTM 14d 506 506 532 532
Oregon Green 488 BAPTA-1TM 170 494 494 523 523
Oregon Green 488 BAPTA-2TM 580 494 494 523 523
Rhod-2 1.0d 556 553 576 576
Calcium OrangeTM 328 549 549 575 576
Calcium CrimsonTM 185 589 589 615 615
Dextran-conjugatede
Fluo-4 dextran (MW 10,000) �600 �494 �494 �518 �518
Calcium Green-1 dextran (MW 3000–70,000)f �240–540f �509 �509 �534 �534
Oregon Green 488 BAPTA-1 dextran (MW 10,000) �265 �496 �496 �524 �524
Ratiometric
Monomeric
Fura-2 224b 362 335 512 505
Fura RedTM 140 473 436 670 655
Indo-1 250b 349 331 485 410
Dextran-conjugatede
Fura(-2) dextran (MW 10,000) �240 364 338 501 494
aData from Tsien (1980), Grynkiewicz et al. (1985), Minta et al. (1989), Haugland (1992), and Molecular Probes, The
Handbook (web publication, Invitrogen Corporation).bEVective Kd in the presence of 1 mMMg2þ. (Generally, competition byMg2þ slightly reduces the aYnity of any indicator
for Ca2þ.)cDiVerent names for the same molecule.dmM.eThe Kd and absorption and emission maxima of dextran-conjugated indicators can vary from lot-to-lot and is dependent
on the molecular weight of the dextran used as well.fKd is reported to be diVerent between low- and high-MW versions: �540 nM for MW 3000, �240–250 nM for MW
10,000 and 70,000.
5. Measuring [Ca2þ] with Fluorescent Indicators 115
peaks remain essentially unchanged. Excitation spectra of Fluo-3 (Minta et al.,
1989), a nonratiometric indicator, at saturating and ‘‘zero’’ [Ca2þ] are shown in
Fig. 2. It can be seen that peak excitation occurs at �505 nm irrespective of
whether the indicator is Ca2þ-free or Ca2þ-bound—the defining characteristic of
a nonratiometric indicator. In contrast, ratiometric indicators exhibit not only
intensity changes with changing [Ca2þ] but the Ca2þ-free and Ca2þ-bound forms
CO2−
CO2− CO2
−CO2
−
CO2−
CO2−
−O2C
−O2C
CO2−
CO2− −O2C
−O2C
CO2−
CO2− −O2C
−O2C
−O2C −O2C−O2C
CO2−
CO2−
−O2C−O2C
CO2−
CO2−
−O2C−O2C
−O2C
−O2C−O2C
N N N N
N
N
O O O O O
CO2−
CO2−
CO2−
CO2−
N
N
N
OOO
O
O
N
N N
OO
N N
OO
−O2C
N N
OO
O
O
CO2−
N
NH
N
N
N+
H
N
O
O Rhod-2
Indo-1Fura-2
BAPTAEGTA Quin2
NH
O
O
O
Calcium green-2
Oregon green 488 BAPTA-2ClF
O
O
O
O
X
XFluo-
R
X
2 or −8*
3
4
H H or CH3
CH3
CH3
Cl
F
R
X
X X
X
X
O−
O−
−O
Fig. 1 Structures of selected fluorescent Ca2þ indicators and the Ca2þ chelators, EGTA, and BAPTA.
All molecules are represented in their polycarboxylate, Ca2þ-sensitive forms. The following conventions
have been used in these structural drawings: (1) Implicit carbon: Every unlabeled vertex, whether
internal or terminal, represents a carbon atom. (2) Implicit hydrogen: Every carbon has a suYcient
number of (undrawn) hydrogens to make the total number of bonds to that carbon equal to 4. (3)
Explicit heteroatoms: non-carbon, non-hydrogen atoms (e.g., O, N) are labeled explicitly; hydrogens
attached to the heteroatom are also explicitly drawn. For example, OH is equivalent to
CH3–CH¼CH–CH2–OH. *DiVerent names for the same molecule.
116 Joseph P. Y. Kao et al.
350
Ca2+-bound
Ca2+-free
400
Wavelength (nm)
Flu
ores
cenc
e in
tens
ity
450300
Fig. 3 Excitation spectra of Ca2þ-bound and Ca2þ-free forms of Fura-2 (lemission¼505 nm).
440420 460
Wavelength (nm)
Ca2+-free
Ca2+-bound
Flu
ores
cenc
e in
tens
ity
480 500 520
Fig. 2 Excitation spectra for Fluo-3 (lemission¼525 nm). The Ca2þ-free form of Fluo-3 is �100 times
less bright than the Ca2þ-bound form.
5. Measuring [Ca2þ] with Fluorescent Indicators 117
of the indicator actually have distinct spectra, the maxima in which occur at
diVerent wavelengths (the spectra show wavelength shifts). The two ratiometric
indicators most commonly used are Fura-2 and Indo-1 (Grynkiewicz et al., 1985).
For Fura-2, significant shifts are observed in the excitation spectra (Fig. 3) but not
in the emission spectra. Indo-1 shows a significant shift primarily in its emission
spectra. For nonratiometric indicators, because intensity monitored at a single-
118 Joseph P. Y. Kao et al.
wavelength is the only experimental measurement that is related to [Ca2þ], intensitychanges arising from factors unrelated to changes in [Ca2þ] (e.g., changes in cell
thickness, leakage of indicator from the cell) can confound interpretation of the
intensity data. In contrast, because the Ca2þ-free and Ca2þ-bound forms of ratio-
metric indicators are characterized by spectral peaks at diVerent wavelengths,
intensity measurements can be made at two diVerent wavelengths, and the ratio
between these intensities is quantitatively related to [Ca2þ] (Grynkiewicz et al.,
1985). Obtaining a ratio minimizes the eVect of many artifacts that are unrelated
to changes in [Ca2þ]—for example, a change in cell thickness or indicator loss from
the cell would aVect intensities at the two wavelengths equally, so the eVect wouldcancel when the two intensities are ratioed. The two commonly used ratiometric
indicators, Fura-2 and Indo-1, require excitation in the ultraviolet (UV) range,
whereas most of the common nonratiometric dyes use visible excitation light.
Although the ratiometric dyes can be calibrated more reliably (Section V), some-
times avoiding using UV light for excitation may be necessary (e.g., UV can excite
significant autofluorescence in some biological preparations and can photolyze
photosensitive ‘‘caged’’ compounds). Clearly, in practice, instrumentation for
using ratiometric indicators is more complex than that for nonratiometric
indicators.
Quin2 (Tsien, 1980; Tsien et al., 1982) is the archetypal tetracarboxylate indicator
listed in Table I (structure in Fig. 1). Its properties and applications as a nonratio-
metric indicator have been reviewed in detail (Tsien and Pozzan, 1989). However,
Quin2 has been superseded by new generations of nonratiometric and ratiometric
indicators. Of the nonratiometric indicators listed in Table I, the Fluo and Calcium
Green series as well as Oregon Green 488 BAPTA-2 incorporate fluorescein chro-
mophores and are, therefore, excited at wavelengths typical of fluoresceins. The
Fluo dyes, Calcium Green-2 and Oregon Green 488 BAPTA-2 exhibit the largest
intensity changes in their transition from Ca2þ-free to Ca2þ-bound forms (�100-
fold; Haugland, 1992;Minta et al., 1989). This change can be an advantage because,
for a given rise in [Ca2þ], these indicators give a larger increase in brightness
compared to other nonratiometric indicators. Because fluorescence quantum
eYciency3 can range only from 0 to 1, the large intensity diVerence between Ca2þ-bound and Ca2þ-free forms implies that the Ca2þ-free forms of the two indicators
must be only weakly fluorescent. Some researchers find this fact annoying because
cells with relatively low resting [Ca2þ]i (cytosolic free Ca2þ concentration) would
have most of the indicator in the Ca2þ-free form and therefore would be quite dim.
Rhod-2, Calcium Orange, and Calcium Crimson are indicators that incorporate
rhodamine-type chromophores and therefore are excited at much longer wave-
lengths than are the Fluo and Calcium Green dyes. When the acetoxymethyl (AM)
3 Fluorescence quantum eYciency, symbolized as FF or QF, is the fraction of total light absorbed
that is emitted as fluorescence. Fluorescence quantum eYciency may also be thought of as the proba-
bility that a molecule will emit fluorescence after absorbing a photon. Being a probability, the quantum
eYciency can have a value between 0 and 1.
5. Measuring [Ca2þ] with Fluorescent Indicators 119
ester is used to load cells, Rhod-2 loads well into mitochondria; up to 80% of the
intracellular dye is located in these organelles. The use of Rhod-2 to monitor Ca2þ
signals in mitochondria is outlined in Section VII.
A choice of ratiometric indicator can be made on practical grounds. Typically,
Fura-2 is excited alternately at two diVerent wavelengths, whereas the emission is
collected at a single fixed wavelength. Therefore, the pair of intensity measure-
ments, whether in imaging or in single-cell microfluorometry, must be collected
sequentially. Indo-1, on the other hand, usually is excited at a fixed wavelength
whereas emission is monitored simultaneously at two diVerent wavelengths, that is,emission from the Ca2þ-bound and Ca2þ-free forms of the indicator can be
collected simultaneously. Therefore, Indo-1 potentially can give better temporal
resolution. However, in conventional imaging, Indo-1 can be more diYcult to use
because the two emission images, usually collected through slightly diVerentoptical paths, can be diYcult to keep in spatial registration. Fura-2 has been the
most widely used ratiometric Ca2þ indicator, both in conventional imaging and in
single-cell measurements. Indo-1 has, however, been used successfully in UV laser-
scanning confocal imaging applications (Motoyama et al., 1999; Niggli et al., 1994;
Sako et al., 1997). Fura RedTM is touted as a ratiometric indicator whose excita-
tion and emission wavelengths are both in the visible range. This indicator suVersfrom having very low fluorescence quantum eYciency (�0.013 in the Ca2þ-freeform; J.P.Y. Kao, unpublished results4). Fura Red diVers from the other ratio-
metric indicators because its fluorescence intensity decreases upon binding Ca2þ.The relatively low quantum eYciency implies that higher indicator concentrations
and/or higher excitation light intensities are required.
The dextran-conjugated dyes are biopolymers with pendant indicator molecules.
The dextran-conjugated indicators listed in Table I are available with dextran
molecular weights of 3000, 10,000, or 70,000 (Invitrogen Corporation, Molecular
Probes Brand). Being membrane-impermeant, dextran conjugates must be loaded
into cells by an invasive technique such as microinjection. Whereas the monomeric
indicators can leak out of cells at a steady rate (Section III.C), dextran-conjugated
indicators tend to have long residence times in cells. Therefore, dextran-conjugated
dyes can be useful in applications in which long-term monitoring of [Ca2þ]i isrequired. Instances also occur in which cells rapidly transport monomeric dyes
into internal organelles (Hepler andCallaham, 1987) but do not do so when dextran
conjugates are used. Because the conjugates are made by covalent attachment of
monomeric indicators to dextran polymers, individual indicator monomers can
reside in slightly diVerent local microenvironments on the polymer. Therefore, the
conjugates, rather than having a unique Kd and identical spectral properties, are
characterized by a range of microscopic Kds and a distribution of spectral proper-
ties. These characteristics provide a likely explanation for lot-to-lot variations inKd
and spectral characteristics.
4 The quantum eYciency of the Ca2þ-free form of Fura Red was determined relative to carboxy-
SNARF-1.
Esterases
IntracellularExtracellular
Indicator
Indicator
Membrane-permeant
Ca2+
AMO2C
AMO2CAMO2C
AMO2CCO2AM
CO2AMCO2AM
CO2AM
Trapped inside cell
Indicator
Ca2+-sensitive-insensitive
−O2C−O2C
CO2−
CO2−
Fig. 4 Schematic representation of how incubation with the acetoxymethyl (AM) ester results in
intracellular accumulation of a polycarboxylate indicator. The hydrophobic (lipophilic) AM ester
readily diVuses into the cell through the cell membrane. Abundant cellular esterases cleave the AM
ester groups to generate the Ca2þ-sensitive form of the indicator which, being a polyanion, cannot
escape through the cell membrane and is, therefore, trapped inside the cell.
120 Joseph P. Y. Kao et al.
III. Loading Indicators into Cells
The common fluorescent indicators for Ca2þ are polycarboxylate anions that
cannot cross lipid bilayer membranes and therefore are not cell-permeant. In the
negatively charged form, the indicators can be introduced into cells only by
microinjection or through transient cell permeabilization, procedures that require
some special equipment and skill.5 By far the most convenient way of loading an
indicator into cells is incubating the cells in a dilute solution or dispersion of the
AM ester of the indicator. This process is represented schematically in Fig. 4. The
AM group is used to mask the negative charges on the carboxyl groups present in
the indicator molecule. The AM ester form of the indicator is uncharged and
hydrophobic. Consequently, it can pass through the cell membrane and enter the
cell interior. The carboxyl groups in the indicator, however, are essential to the
ability of the indicator molecule to sense Ca2þ; therefore, the AM groups must be
removed once the AM ester has entered the cell. Because the AM group is labile to
enzymatic hydrolysis by esterases present in the cell, the AM esters are processed
intracellularly to liberate the Ca2þ-sensitive polycarboxylate form which, being
multiply charged, becomes trapped inside the cell. Trapping of the polyanionic
form of the indicator allows cells to accumulate up to hundreds of micromolar of
5 A variety of techniques for loading membrane-impermeant species into cells is discussed byMcNeil
(1989, 2001).
5. Measuring [Ca2þ] with Fluorescent Indicators 121
the Ca2þ-sensitive form of the indicator when incubated with micromolar concen-
trations of the AM ester in the extracellular medium. Several factors influence the
eYciency and quality of indicator loading via the AM ester, and will be discussed
subsequently.
A. Limited Aqueous Solubility of AM Esters
AM esters of the common Ca2þ indicators have molecular weights in excess of
1000. Being large uncharged organic molecules, these esters have very low solubili-
ty in aqueous media. For example, at 25 �C, the solubility of the AM ester of Fura-
2 in pure water is only 0.11 mM (Kao et al., 1990). In biological media, in which the
ionic strength is typically �0.15 M, the solubility of Fura-2 AM would be even
lower. Addition of AM ester in excess of the solubility limit simply would result in
precipitation of solid AM ester, which is eVectively unavailable for loading cells. Inaddition, fine particles of solid AM ester often adhere well to the outer surfaces of
cells or to the extracellular matrix and can contribute large Ca2þ-insensitivefluorescence signals to the measurement.6 A convenient solution to the solubility
problem is the use of Pluronic F-127, a mild nonionic surfactant,7 as a dispersing
agent for AM esters. Typically, aliquots of Pluronic and AM ester stock solutions
in dimethylsulfoxide (DMSO) are mixed intimately before dispersal into an aque-
ous medium.8 The Pluronic is presumed to sequester the AM ester in micellar form,
thus preventing precipitation, and the micelles serve as a steady source to replenish
AM esters taken up by cells. The net result is significantly improved loading of
indicators into cells. Details of the loading procedure are described in Section III.D
below.
B. Dye Compartmentalization: Loading of Indicator into Subcellular CompartmentsOther than the Cytosol
1. Minimizing Compartmentalization
In typical experiments, one usually wishes to monitor changes in the concen-
tration of Ca2þ in the cytosol; therefore, ideally, one would like the indicator to
be loaded exclusively into the cytosol. This ideal situation is almost never
realized for two reasons. First, because the AM ester form of the indicator is
membrane-permeant, it can enter not only the cytosol but all subcellular
6 This is a problemwith AM ester that are fluorescent (e.g., Fura-2 AM and Indo-1 AM) but not with
nonfluorescent AM esters (e.g., AM ester of the Fluo series).7 Pluronic F-127 is manufactured by BASF Wyandotte Corporation, Wyandotte, Wisconsin. Being
a surfactant used on the industrial scale, it is inexpensive.8 With slight warming, Pluronic F-127 can be dissolved in DMSO at almost 25% (w/v). Such a highly
concentrated stock solution is inconvenient to use, however: as the solution absorbs moisture from the
air, the Pluronic will precipitate, and the resulting particulate suspension is diYcult to pipette. A 15–20%
solution in DMSO is a convenient formulation.
122 Joseph P. Y. Kao et al.
membrane-enclosed compartments as well. Although this process occurs to a
large extent in the cytosol, enzymatic hydrolysis of AM esters also can take
place within subcellular organelles. Therefore, some fraction of the indicator
molecules tend to be trapped in noncytosolic compartments. Second, some cell
types actively endocytose material from the incubation medium (Malgaroli
et al., 1987), including dispersed AM esters, which are then hydrolyzed to
release fluorescent indicator molecules within organelles of the endocytotic
pathway. Presumably, indicator molecules liberated in this way can end up in
a variety of organelles that are connected to the endocytotic pathway by
vesicular traYc. Because most subcellular organelles [e.g., endoplasmic reticu-
lum (ER), lysosomes] tend to have high intraorganellar [Ca2þ] (>mM), indica-
tors confined to these organelles would be saturated with Ca2þ and would
contribute a high-[Ca2þ] fluorescence signal that would not vary with changes
in cytosolic [Ca2þ]. Therefore, the net eVect of compartmentalized dye is
biasing measured cytosolic free Ca2þ concentration toward higher values.
The first cause of dye compartmentalization just stated is a reflection of an
inherent imperfection of the AM ester loading technique and cannot be remedied
easily. The second cause is a cell biological process and can be attenuated.
Because endocytosis is a temperature-dependent process, cells loaded at lower
temperatures with AM ester tend to show less compartmentalization of indicator
(Malgaroli et al., 1987). The following results illustrate this point. In REF52
fibroblast cells, roughly 30% of total intracellular Fura-2 was in noncytosolic
compartments when loading via the AM ester was carried out at 37 �C, whereasonly 10% was compartmentalized when loading was performed at 23 �C. Although
endocytosis is known to be blocked at 10 �C in mammalian cells and at 4 �C in
amphibian cells, loading at the lowest biologically permissible temperature does
not necessarily yield the best results, because processing of AM esters by esterases
in the cytosol is also temperature-dependent. At very low temperatures, the con-
centration of indicator accumulated in the cytosol can be quite low. Optimal
loading temperature is determined empirically to be the temperature at which
dye compartmentalization is minimized while good cytosolic loading is main-
tained. In practice, convenience often dictates loading at room temperature as a
reasonable compromise.9
2. Assessing Extent of Compartmentalization
The extent of indicator compartmentalization can be estimated through a simple
semiquantitative procedure that is based on the observation that micromolar
concentrations of digitonin primarily permeabilizes the plasma membrane
9 When loading is done in air, the incubation medium should be bicarbonate-free (i.e., some other
buVer such as HEPES should be used to maintain the pH of the medium). Otherwise, steady loss of CO2
will shift the CO2–HCO3�–CO3
2� equilibrium and rapidly alkalinize the medium.
0
Flu
ores
cenc
e in
tens
ity (
a.u.
)
100 200
Digitonin
Triton X-100
Fi
Fd
Fb
Time (s)
300 400
Fig. 5 Procedure for assessing dye compartmentalization. REF52 fibroblast incubated with 1 mMFura-2 AM dispersed with Pluronic F-127 in Minimal Essential Medium (MEM) for 60 min in an air
incubator at 37 �C.Measurementwas done inHanks’ Balanced Salt Solution (HBSS) containing 2.6 mM
EGTA (suYcient to reduce extracellular [Ca2þ] to <1 mM), and buVered at pH 7.4 with HEPES.
The concentration of digitonin was 20 mM; that of Triton X-100, 1%, w/v. (a.u. = arbitrary units).
5. Measuring [Ca2þ] with Fluorescent Indicators 123
and release cytosolic dye, whereas 1% Triton X-100 can permeabilize and release
dye from subcellular organelles (e.g., see Kao et al., 1989). The procedure consists
of monitoring total fluorescence from a cell or a field of cells, preferably bathed
in low-Ca2þ medium,10 and treating the cells first with digitonin and then
with Triton X-100. Figure 5 shows the procedure being applied to a cell loaded
with Fura-2 AM. The fluorescence measured before treatment (Fi) represents
contributions from cytosolic and compartmentalized dye plus background.
After digitonin release of cytosolic dye, the fluorescence measured (Fd) represents
compartmentalized dye plus background. The fluorescence level attained after
Triton X-100 treatment is considered the background (Fb). The fraction of total
intracellular dye that is compartmentalized in organelles is simply (Fd�Fb)/
(Fi�Fb).
10 Nominally Ca2þ-free medium (i.e., medium from which Ca2þ salts have been omitted) or, better
still, medium containing suYcient EGTA to make [Ca2þ]< 1 mM (see Section IV.B.1). It is important to
keep [Ca2þ] low in this experiment because when [Ca2þ] is elevated, permeabilized cells lose their
integrity rapidly, and dye molecules may leak from organelles even before Triton is applied.
B
Frac
tion
rem
aini
ng
0 5 10 15
Time (min)
20
Proben.
Sulfin.
Noinhibitor
35�C
25
1.0
0.9
0.8
0.7
0.6
1.0A
0.9
0.8
Frac
tion
rem
aini
ng
0.7
0.6
0.50 10 20
35 �C
30 �C
Time (min)
30
Fig. 6 (A) Time course of indicator loss at 30 and 35 �C from REF52 cells loaded with Fura-2. (B)
EVect of probenecid (1 mM) and sulfinpyrazone (75 mM) on indicator extrusion from REF52 cells at
35 �C; arrowmarks the time of drug application. For all measurements, the cells were loaded with Fura-
2 by incubation with AM ester/Pluronic dispersion in HEPES-buVered HBSS at 25 �C. Experiments
were done in HBSS at the specified temperatures. Total Fura-2 fluorescence (FT¼F340þF380) is
monitored over time. Each trace was normalized by dividing by the value of FT at time 0.
124 Joseph P. Y. Kao et al.
C. Dye Leakage or Extrusion from Cells
Once an indicator is loaded into cells, it leaks out at a rate that is strongly tempera-
ture-dependent. For mammalian cells, the loss rate is maximal at 37 �C and drops oVsharply as temperature is lowered,11 as shown in Fig. 6A. Loss of indicator occurs by
an extrusion mechanism for organic anions (Di Virgilio et al., 1988) and can be
blocked eVectively by inhibitors of uric acid transport such as probenecid and
sulfinpyrazone (DiVirgilio et al., 1988, 1990), as illustrated inFig. 6B. Sulfinpyrazone
at tens to hundreds ofmicromolar has been found to be eVective, whereas probenecidworks at millimolar dosage. High concentrations of inhibitor virtually can stop
indicator extrusion but also can induce signs of cellular stress such as blebbing.
Therefore, using the minimum concentration of inhibitor necessary to reduce the
rate of indicator loss to a tolerable level without attempting to block the process
altogether is advisable. In general, slowing indicator loss by lowering the temperature
at which experiments are conducted is preferable to using transport inhibitors.
11 The sharp increase in rate of indicator leakage with rising temperature is another reason why
compartmentalization is a more severe problem when cells are incubated with AM esters at higher
temperatures. Higher temperature accelerates loss of dye from the cytosol but not from organelles, so
compartmentalized dye becomes a larger proportion of the total intracellular dye content.
5. Measuring [Ca2þ] with Fluorescent Indicators 125
Probenecid and sulfinpyrazone are both hydrophobic organic acids and, as such, are
practically insoluble in water. These molecules must be neutralized with a stoichio-
metric amount of base (e.g., NaOH) before an aqueous stock solution can be
prepared.
D. Procedure for Loading
Stock solutions of the AM ester of the indicator can be made with dry DMSO as
solvent. Typical concentrations can be in the range of 0.1–10 mM. Such DMSO
solutions may be stored safely in screw-capped polypropylene microcentrifuge
tubes in the freezer for many months without apparent degradation. Dry DMSO
is used for making a solution of Pluronic F-127 at a concentration of 15–20% (w/w
or w/v is not crucial). This stock solution can be stored at room temperature. When
exposed to air, this concentrated stock solution slowly absorbs moisture until
Pluronic F-127 begins to precipitate. Because a heterogeneous Pluronic stock is
diYcult to transfer, preparing a fresh stock is preferable at this stage. Usually one
can load cells by incubation in medium containing (nominally) 0.1 to a few tens of
micromolar AM ester in a Pluronic dispersion. Typically 0.5–1 mL of Pluronic
stock is suYcient for dispersing 1–10 nmol AM ester. Thus, 1 mL of 20% Pluronic
stock is adequate for dispersing 1 mL of 10 mM AM ester stock into 1 mL of
medium to yield a 10 mM AM ester solution (1000-fold dilution). Using a dilution
of �1:1000 minimizes the DMSO concentration in the final loading medium (to a
few parts per thousand). Premixing the requisite volumes of AM ester stock and
Pluronic stock is advisable since this minimizes the chances of AM ester precipita-
tion during aqueous dispersal. The Pluronic-AM mixture is then dispersed into
aqueous loading medium.
Serum proteins [e.g., bovine serum albumin (BSA)] often have a salutary eVecton cells and also can improve loading eYciency, possibly by acting as hydrophobic
carriers for the AM ester and preventing its precipitation. Thus, including BSA
(0.5–1%) or serum (a small percentage) in the incubation medium may be
advantageous.
Enzymatic processing of an AM species to yield the Ca2þ-sensitive indicator
typically consists of sequential hydrolysis of four to eight AM ester groups,
depending on the particular indicator chosen. Only when all the AM groups on a
molecule are hydrolyzed does the molecule become properly Ca2þ-sensitive. Forthis reason, after removal of AM-containing loading medium, allowing the cells
some extra time (e.g., 20 min at room temperature) to complete intracellular
processing of the most recently trapped, partially hydrolyzed AM species can be
useful in some cases.
It is reassuring to convince oneself that suYcient AM ester is present in the
loading medium, so the amount of indicator taken up by cells is not limited by the
amount available. For a typical example, assume that 1 million cells are loaded in
2 mL medium containing 2 mMAM ester. The total amount of AM ester available
126 Joseph P. Y. Kao et al.
is 4 nmol. If the cells are 20 mm in diameter, then the 1 million cells have a total
intracellular volume of 4.2 mL. Further, if cells are loaded to a final intracellular
indicator concentration of 150 mM (a generous estimate), then the total amount of
AM uptake by cells is 0.63 nmol, which is still much less than the 4 nmol available
in the loading medium.
IV. Manipulation of [Ca2þ]
In studying Ca2þ-dependent cellular processes, raising or lowering intracellular
or extracellular [Ca2þ] is frequently desirable. Conventional techniques for achiev-ing these ends require the use of Ca2þ buVers or ionophores and will be discussed in
this section.
A. Using EGTA and BAPTA as Extracellular Ca2þ BuVers
Because it is highly selective for binding Ca2þ over Mg2þ,12 EGTA is the most
commonly used Ca2þ buVer. However, because two of the ligand atoms in EGTA
are tertiary alkylamino nitrogens, the two highest pKas of EGTA are 8.90 and
9.52,13 implying that at physiological pH EGTA will exist primarily as protonated
species—a fact that is illustrated more quantitatively in Fig. 7. For example, Fig. 7
shows that, at pH 7.2, �98% of EGTA in solution exists as H2EGTA2�, �2% as
HEGTA3�, and only a negligible fraction is in the EGTA4� form. Therefore, the
Ca2þ-binding reaction near physiological pH is fairly represented as
H2EGTA2� þ Ca2þ > CaEGTA2� þ 2Hþ
That two Hþ ions are liberated in the binding reaction means that the binding of
Ca2þ by EGTA should have very steep pH dependence, as a plot of pK0d(Ca)
14
versus pH indeed shows (Fig. 8). For a concrete example, a drop in pH from 7.2 to
7.1 changes the K0d(Ca) of EGTA by a factor of �1.6, that is, small errors in pH
can lead to significant uncertainties in the dissociation constant. In contrast,
12 ForEGTA,DpKd¼ pKd(Ca2þ)� pKd(Mg2þ)¼ 5.58; therefore, EGTAbindsCa2þmore tightly than
Mg2þ by a factor of 380,000 (i.e., 105.58). For comparison, in the case of EDTA, DpKd ¼ 1.78, which
represents only a 60-fold diVerence in EDTA’s aYnity for Ca2þ and Mg2þ. BAPTA [1,2-bis(o-aminophe-
noxy)ethane-N,N,N0,N0-tetraacetic acid] has a selectivity similar to that of EGTA: DpKd ¼ 5.20.13 At 25�C and 0.10M ionic strength. Data pertaining to EGTA that are used in this section are from
Martell and Smith (1974).14 In the metal chelator literature, Kd is used for the ‘‘absolute’’ (or intrinsic) dissociation constant
and represents the dissociation constant characterizing the fully deprotonated form of the chelator. K0d
represents Kd that has been corrected for the weakening eVect of acidic pH (thus K0d is the working
dissociation constant at a specific pH). This convention (Kd vs. K0d) is not followed consistently in the
applications literature. Details of how pH correction is applied to convert Kd into K0d are described in
Appendix 1.
100
H2EGTA2−
Free
EG
TA e
xist
ing
asH
2EG
TA2−
, HE
GTA
3−, o
r E
GTA
4− (%
)
EGTA4−
HEGTA3−
50
60
7 8 9pH
10 11 12
Fig. 7 Percentage of free EGTA existing as H2EGTA2�, HEGTA3�, and EGTA4� as a function of
solution pH. Calculations performed with data for EGTA (at 0.1 M ionic strength, 25 �C) tabulated by
Martell and Smith (1974); see Appendix 1 for algebraic details.
5. Measuring [Ca2þ] with Fluorescent Indicators 127
knowing that the two highest pKas of BAPTA are 5.47 and 6.36 (Tsien, 1980), one
infers that the ability of BAPTA to bind Ca2þ should be only very weakly
dependent on pH, as shown in Fig. 8. Comparison of the two traces in Fig. 8
shows that BAPTA has the advantage of being only weakly pH dependent in the
physiological pH range. The fact that the two traces cross between pH 7.2 and 7.3
implies that EGTA has the potential advantage of being a progressively stronger
binder of Ca2þ above the crossover point (e.g., about two- and ninefold stronger
than BAPTA at pH 7.5 and 7.8, respectively). The pH insensitivity of BAPTA
makes it a less troublesome Ca2þ buVer to use, although it is more costly than
EGTA.
B. Lowering Extracellular [Ca2þ]
In an experiment, lowering extracellular [Ca2þ] is often desirable. Depending on
how low one wishes to clamp the extracellular [Ca2þ], one of the approaches
described in the following sections may be adopted. The procedures require either
a stock solution of 1 M Na2H2EGTA at a pH near neutral or a stock solution of
1 M Na4BAPTA.
12
12
11
11
10
10
9
9
pH
BAPTA
EGTA
8pK� d
8
7
7
6
6
5
4
Fig. 8 Plot of pK0d(Ca) versus pH for EGTA and BAPTA. Calculations performed with data for
EGTA from Martell and Smith (1974) and for BAPTA from Tsien (1980).
128 Joseph P. Y. Kao et al.
1. Lowering [Ca2þ] to <1 mM but not Approaching 0
a. Using EGTANa2H2EGTA can be added directly to calcium-containing medium at a concen-
tration that is 3–4 times15 the concentration of Ca2þ in the medium. Because binding
of Ca2þ to H2EGTA2� releases protons and acidifies the medium, simultaneously
adding TRIS base [tris(hydroxylmethyl)aminomethane] at a concentration equal to
�2.2 times the Ca2þ concentration in the medium is also necessary. This amount of
TRIS scavenges protons released from H2EGTA2� and maintains the pH of the
medium at nearly the level before EGTA addition. This procedure can be applied
confidently down to pH �6.8. At pH levels that are much lower, the strong pH
sensitivity weakens EGTA and makes it inconvenient to use as a Ca2þ scavenger.
b. Using BAPTAAdd Na4BAPTA at a concentration equal to 3 times the Ca2þ concentration in
the medium. Essentially no pH adjustments are necessary. Because of its relative
insensitivity to decreasing pH (Fig. 8), BAPTA can be used more conveniently for
scavenging Ca2þ at lower pH values than EGTA.
15 3 times if pH � 7.0, 4 times if pH < 7.0.
5. Measuring [Ca2þ] with Fluorescent Indicators 129
2. Lowering [Ca2þ] to a Level Approaching 0
If one desires a medium in which the free Ca2þ concentration approaches
‘‘zero,’’ one can prepare nominally calcium-free medium16 that also contains
EGTA or BAPTA at millimolar concentrations. Again, to avoid problems of
rapid weakening of the Ca2þ aYnity of EGTA with decreasing pH, using
BAPTA at pH<7 is more convenient.
3. Setting Extracellular [Ca2þ] to a Precisely Known Value
When the extracellular [Ca2þ] must be known precisely, medium containing a
well-defined Ca2þ buVer system at a fixed pH must be made. The preparation of
such solutions is detailed in Chapter 9.
C. Divalent Cation Ionophores
1. Properties of Br-A23187 and Ionomycin
Br-A2318717 and ionomycin are ionophores that form complexes with divalent
metal cations; the complexes are lipid-soluble and thus can cross cellular mem-
branes. These two ionophores are commonly used to increase the permeability of
biological membranes to Ca2þ. Understanding the diVerences between the two
makes it possible to make a judicious choice during an experiment.
Ionomycin can lose two acidic hydrogens and, as a dianion, can form an
uncharged 1:1 complex with a divalent metal ion such as Ca2þ or Mg2þ. Br-A23187 can lose a single acidic hydrogen and form an uncharged 2:1 complex
with a divalent metal ion. This diVerence makes ionomycin potentially more
eVective in binding and transporting divalent cations (e.g., two molecules of Br-
A23187 are needed to bind and carry a single Ca2þ whereas only one molecule of
ionomycin is suYcient).
Compared with Br-A23187, ionomycin has somewhat better selectivity for Ca2þ
over Mg2þ; ionomycin prefers Ca2þ by a factor of �2, whereas Br-A23187 shows
essentially no preference for one cation over the other (Liu and Hermann, 1978). In
addition, these ionophores actually do not bind Ca2þ very tightly [e.g., Kd(Ca2þ)�
100 mM for ionomycin; J.P.Y. Kao, unpublished results.18] These factors suggest
that the two ionophores would be ineYcient in mediating Ca2þ transport when
relatively low Ca2þ concentrations are involved (i.e., at [Ca2þ]�Kd, e.g., <1 mM),
because at such low Ca2þ concentrations, only a minute fraction of total
16 Because calcium is a ubiquitous ‘‘contaminant’’ in the environment, nominally calcium-free
solutions still can contain micromolar levels of Ca2þ.17 The parent compound, A23187, is fluorescent and should be avoided in fluorescence work. The
presence of the bromine atom in 4-bromo-A23187 (Br-A23187) eVectively quenches the intrinsic
fluorescence of the ionophore and makes the molecule useful in fluorescence microscopy.18 Determined by absorption spectroscopy at pH 11, at which all ionomycin in solution would be
present as the fully deprotonated dianion.
130 Joseph P. Y. Kao et al.
ionophore is actually engaged in Ca2þ binding and transport. This problem
becomes evident when either ionophore is used in calibrating Ca2þ indicators in
cells (see Section V.B).
The most significant diVerence between the two ionophores lies in the pH
dependence of their ability to transport Ca2þ (Liu and Hermann, 1978). Transport
of Ca2þ by Br-A23187 approaches a maximum at pH 7.5, whereas Ca2þ transport
by ionomycin does not reach a maximum until pH 9.5. The pH at which half-
maximal transport is achieved is �6.4 for Br-A23187 and �8.2 for ionomycin.
Therefore, if one desires to increase transport of extracellular Ca2þ into cells in
acidic media (pH<7.0), Br-A23187 is a much better choice than ionomycin.
2. Using Br-A23187 and Ionomycin
Ionomycin can be obtained as either the free acid or the Ca2þ salt. Br-A23187 is
available as the free acid. All forms are soluble in dry DMSO, which can be used to
prepare stock solutions. Because these ionophores are very hydrophobic, they are
bound avidly by serum proteins. Serum proteins such as BSA, when present in the
medium, greatly reduce the eVectiveness of ionophores and, if possible, should be
left out of the experimental medium when ionophores are to be used. Otherwise,
much higher concentrations of ionophore must be used. Br-A23187 and ionomycin
have been used at concentrations ranging from 10�7 to 10�5 M. In addition to
increasing Ca2þ flux across the plasma membrane, Br-A23187 and ionomycin also
transport Ca2þ out of intracellular calcium stores into the cytosol. Therefore, in
the presence of these ionophores, intracellular calcium stores are rapidly depleted
(Kao et al., 1990).
D. BuVering Changes in Intracellular [Ca2þ]
1. Increasing Intracellular Ca2þ BuVering Capacity by BAPTA Loading
When a change in [Ca2þ]i (a Ca2þ signal) is correlated with a biological process,
one can ascertain whether the Ca2þ signal is essential in the process by blocking the
change in [Ca2þ]i with a calcium chelator. By far the easiest way to introduce extra
Ca2þ buVering capacity into cells is by incubation with BAPTA AM in Pluronic
dispersion. Compared with the AM esters of common Ca2þ indicators, BAPTA
AM has much higher aqueous solubility—15 mM at 25 �C (Kao et al., 1990).
Therefore, BAPTA can be loaded eYciently into cells via the AM ester. BAPTA
AM is loaded into cells in precisely the same way that AM esters of indicators are
loaded. Cells can be loaded with AM esters of BAPTA and an indicator simulta-
neously. Figure 9 illustrates the eVect of intracellular BAPTA loading on normal
changes in [Ca2þ]i. Figure 9A shows the changes in [Ca2þ]i in a REF52 cell loaded
with Fura-2 in response to sequential application of 1 mM bradykinin and 1 mMBr-A23187. Figure 9B shows the responses in a similar cell loaded with Fura-2 and
BAPTA. These results clearly demonstrate that the presence of suYcient BAPTA
practically eliminates the rapid and transient rises in [Ca2þ]i elicited by an agonist.
1.2
1.0
0.8
0.6
[Ca2+
] i (m
M)
0.4
0.2
0.00 10
A B
Bra
dyki
nin
Bra
dyki
nin
Br–
A23
187
Br–
A23
187
20 30
−BAPTA +BAPTA
0 10 20
Time (min)Time (min)
30
Fig. 9 BuVering action of intracellular BAPTA on changes in [Ca2þ]i. (A) Changes in [Ca2þ]i of aREF52 cell treated with 1 mMbradykinin and then 1 mMBr-A23187. (B) Changes in [Ca2þ]i of a REF52
cell, preloaded with BAPTA, in response to the same treatments as in (A). Cells were loaded with 1 mMFura-2 AM in Pluronic dispersion for 85 min at 25 �C. For (B), 20 mMBAPTA AMwas also present in
the incubation medium. Experiments were done in HBSS.
5. Measuring [Ca2þ] with Fluorescent Indicators 131
Indeed, even the massive rise resulting from a combination of Ca2þ influx and
discharging of intracellular calcium stores mediated by Br-A23187 is suppressed
substantially by the buVering action of BAPTA.
2. Possible Controls for the Use of BAPTA
Similar to EGTA, BAPTA is a chelator not only for Ca2þ but also for other
multivalent metal cations. Thus, one may wish to ensure that any inhibitory eVectobserved when using BAPTA is caused strictly by the ability of BAPTA to buVerCa2þ, and not because it is scavenging other biochemically important metal ions
such as Zn2þ. The reagent used to control for heavy metal scavenging by BAPTA is
TPEN (N,N,N0,N0-tetrakis(2-pyridylmethyl)ethylenediamine) (Fig. 10), a mem-
brane-permeant metal ion chelator that shows a marked preference for binding
heavy metal cations over Ca2þ (Anderegg et al., 1977). Whereas the Kd(Ca2þ) of
TPEN is 40 mM (Arslan et al., 1985), Kd(Zn2þ) is 2.6�10�16 M (Anderegg and
Wenk, 1967). This enormous selectivity for binding heavy metal ions over Ca2þ
enables TPEN to scavenge heavy metal ions very eVectively, even in the presence of
N N
NN
N
N N
O O
BAPTATPEN Half-BAPTA
OH3C
CO2−
CO2− CO2
−−O2C−O2C −O2C
N N
Fig. 10 Structures of TPEN, BAPTA, and half-BAPTA.
132 Joseph P. Y. Kao et al.
millimolar levels of Ca2þ. That TPEN is membrane-permeant means it can be
applied without using any special procedures. Dry DMSO can be used to prepare
stock solutions of TPEN. Typically, TPEN is used in aqueous medium at a
concentration of 10�6–10�5 M.
BAPTA loading through the AM ester is very eYcient; high concentrations of
BAPTA may be accumulated intracellularly. Thus, ascertaining that observed
inhibitory eVects are not the result of cytotoxicity arising from the presence of
high concentrations of a foreign organic anion may be important. In this case, the
control reagent is N-(o-methoxyphenyl)iminodiacetic acid,19 sometimes referred
to as ‘‘half-BAPTA.’’ As can be seen from Fig. 10, half-BAPTA is essentially
chemically identical to BAPTA except that the molecule is only half of BAPTA.
Because the full tetracarboxylate structure of BAPTA is crucial for Ca2þ binding,
half-BAPTA, lacking such a structure, shows only very weak aYnity for Ca2þ
(Kd�3 mM; J.P.Y. Kao, unpublished results). Half-BAPTA is thus expected to
mimic BAPTA in all chemical respects except for the ability to buVer Ca2þ at
physiological concentrations. The AM ester of half-BAPTA is sporadically
available from commercial vendors. Cell loading via the AM ester can be done
as described previously for other AM esters.20
V. Conversion of Indicator Fluorescence Signal intoValues of [Ca2þ]
Although raw fluorescence signals from intracellularly trapped Ca2þ indicators
can be informative in a qualitative way, one still must perform some calibration
before even semiquantitative estimates of [Ca2þ]i can be made. Basic principles of,
19 A trivial name is anisidine-N,N-diacetic acid.20 Because it is processed by esterases to generate only the Ca2þ-insensitive half-BAPTA and yet it is
processed intracellular in the same way that all AM esters are, half-BAPTA AM can also be used as a
control for possible artifacts from AM ester hydrolysis.
5. Measuring [Ca2þ] with Fluorescent Indicators 133
as well as experimental procedures for, calibration for ratiometric and nonratio-
metric indicators are discussed in this section.
A. Calibrating a Nonratiometric Fluorescent Indicator
For a nonratiometric indicator that increases fluorescence emission on binding
Ca2þ, the free Ca2þ concentration is given by
Ca2þ� � ¼ Kd
F � Fmin
Fmax � F
� �ð1Þ
where Fmin is the indicator fluorescence intensity at zero [Ca2þ] (when all
indicator molecules in the sample are Ca2þ-free), Fmax is the indicator fluores-
cence at saturatingly high [Ca2þ] (when all indicator molecules are present as
the Ca2þ-bound form), and F is the measured fluorescence intensity for which
we wish to find a corresponding value of [Ca2þ]. To arrive at a correspondence
between measured F and [Ca2þ]i, Kd, Fmin, and Fmax all must be known.
Whereas Kd usually is predetermined in vitro, Fmin and Fmax must be obtained
in situ. The most straightforward approach would be to try to equilibrate the
indicator-loaded cell with solutions that contain ‘‘zero’’ [Ca2þ] and then high
[Ca2þ]. In practice, however, deficiencies inherent in a nonratiometric indicator
make this approach unattractive. Interpretation of intensity changes is con-
founded by dye leakage, which causes the total indicator fluorescence from the
cell (and therefore Fmin and Fmax) to decrease with time. This basic flaw of
nonratiometric indicators means that obtaining good quantitative estimates of
[Ca2þ]i in cells in which dye leakage or extrusion occurs at significant rates
would be diYcult. In such cases, a laborious calibration would not be justified.
An alternative semiquantitative calibration procedure developed for Fluo-3 is
discussed next.
The calibration procedure for Fluo-3 depends on the fact that, in vitro,
FMn¼0.2Fmax, where FMn is the fluorescence intensity when Fluo-3 is saturated
completely with Mn2þ (Kao et al., 1989; Minta et al., 1989). That Fmax¼100Fmin is
also known from in vitro measurements. Because both Fmax and Fmin can be
expressed in terms of FMn, the only parameter that must be determined experimen-
tally is FMn. In situ calibration then consists of:
1. applying micromolar levels of ionomycin or Br-A23187 to increase perme-
ability of the cell to divalent metal ions;
2. adding suYcient MnCl2 (typically twice the concentration of Ca2þ in solu-
tion)21 to ensure saturation of intracellular Fluo-3; and
21 In Step 2, it is best if the medium contains no carbonate, bicarbonate, or phosphates, which can
form insoluble precipitates with Mn2þ and thus reduce the concentration of free Mn2þ. Moreover, the
light scattering by the particulate precipitates can add considerable noise to the fluorescence signal.
134 Joseph P. Y. Kao et al.
3. permeabilizing the cell with digitonin to release Fluo-3 to permit estimation
of fluorescence background.
Because the fluorescence intensity measured after Step 3 is just the background
signal (including cellular autofluorescence), whereas the intensity after Step 2 is
(FMnþbackground), one can obtain FMn by subtraction. The calibration proce-
dure described here is based on the following assumptions: (1) indicator fluores-
cence intensity is not diminishing rapidly as a result of leakage; (2) the fluorescence
properties (Fmin, Fmax, and FMn) of the indicator are known from in vitromeasure-
ments and are the same in cells as in vitro; and (3) the Kd of the indicator is also the
same in cells as in vitro.
Quin2 is an example of an indicator the fluorescence of which is quenched
completely by heavy metal ions. For calibration of such an indicator, see the
review on Quin2 by Tsien and Pozzan (1989).
B. Calibrating a Ratiometric Fluorescent Indicator
A dual-wavelength ratiometric indicator allows excitation spectral intensity or
emission spectral intensity of the indicator to be monitored at two diVerentwavelengths. If F1 is the fluorescence intensity at wavelength l1, F2 is the fluores-
cence intensity at wavelength l2, and R¼F1/F2, then the free Ca2þ concentration
can be shown to be (Grynkiewicz et al., 1985)
½Ca2þi ¼ Kd
R� Rmin
Rmax � R
� �sf ;2sb;2
� �ð2Þ
where Rmin is the limiting value of the ratio R when all the indicator is in the Ca2þ-free form and Rmax is the limiting value of R when the indicator is saturated with
Ca2þ.22 Experimentally, the factor sf,2/sb,2 is simply the ratio of the measured
fluorescence intensity when all the indicator is Ca2þ-free to the intensity measured
when all the indicator is Ca2þ-bound, with both intensity measurements taken at
l2. On the right side of Eq. (2), with the exception of Kd, which is an intrinsic
property of the indicator, all other terms are ratios of intensities; in forming these
ratios, problems associated with cell shape and dye concentration changes cancel.
Using Eq. (2) to calculate [Ca2þ] requires that Kd be known and that Rmin, Rmax,
and sf,2/sb,2 be determined experimentally. A typical calibration entails
1. increasing Ca2þ permeability of the cell with ionomycin or Br-A23187 in the
presence of ‘‘zero’’ extracellular Ca2þ (EGTA or BAPTA in nominally Ca2þ-free medium; Section IV.B.2), so all intracellular Ca2þ could be depleted;
22 Usually, l1 and l2 are chosen so that intensity measured at l1 consists mostly of fluorescence
emitted by the Ca2þ-bound form of the indicator, whereas intensity at l2 consists mostly of fluorescence
from the Ca2þ-free form. Choosing the wavelength pair in this way increases the diVerence betweenRmin
and Rmax, making it possible to map [Ca2þ] onto a wider range of R values.
5. Measuring [Ca2þ] with Fluorescent Indicators 135
2. increasing extracellular [Ca2þ] in the presence of Ca2þ ionophore so that
Ca2þ could enter the cell to saturate intracellular indicator;
3. permeabilizing the cell with digitonin (at concentrations prescribed in
Section III.B.2) to release cytosolic dye so the background signal may be
measured.23
Although the procedure seems straightforward, a few empirical findings are
helpful in performing a successful calibration:
1. Many cell types do not tolerate severe calcium deprivation well. During Step
1, these cells often become fragile or leaky or, in the case of adherent cells, detach
from the substrate. In many cases, however, one can compensate for the total
absence of Ca2þ by supplementation with elevated concentrations of Mg2þ. Thus,raising the extracellular [Mg2þ] to 5–20 mM can help maintain cell integrity during
a long calibration. Although Mg2þ should bind Fura-2 to a limited extent and
slightly alter its fluorescence spectrum (Grynkiewicz et al., 1985), in practiceRmin is
not aVected significantly by Mg2þ supplementation.
2. Because ionomycin and Br-A23187 become very ineYcient at Ca2þ transport
when intra- and extracellular free Ca2þ concentrations are below micromolar
levels, depleting the cell of Ca2þ entirely is quite diYcult. Therefore, one often
must wait a long time for trueRmin to be reached in Step 1. In the example shown in
Fig. 11, the interval between ionomycin addition and attainment of Rmin was in
excess of 90 min.
3. Step 2 could be performed in two ways. One could add, in combination with
fresh ionophore if desired, suYcient Ca2þ to bind to all the EGTA or BAPTA that
was introduced in Step 1 and still have a large excess of free extracellular Ca2þ, aswell as suYcient TRIS base to counteract any acidification arising from the Ca2þ-EGTA binding reaction. Alternatively, one could replace the medium from Step 1
with nominally Ca2þ-free medium and then add a large excess of Ca2þ in combi-
nation with a fresh dose of ionophore. The aim is to initiate massive Ca2þ influx
into the cell at a rate that overcomes any Ca2þ extrusion mechanism that the cell
may mobilize. In practice, concentrations of ionophore ranging from 10�6 to
10�5 M and external [Ca2þ] in the range of one to several tens of millimolar can
be used.
23 At controlled concentrations, digitonin releases primarily cytosolic dye whereas compartmenta-
lized dye remains with the permeabilized cell and would be subtracted out as background. The
assumption is that intraorganellar [Ca2þ] is significantly higher (> mM) than cytosolic [Ca2þ], socompartmentalized indicator would be essentially completely Ca2þ-bound and thus contribute a con-
stant background to the measured 340- and 380-nm fluorescence signals from the cell. This assumption
would fail if significant amounts of dye are compartmentalized into organelles that do not have high
luminal [Ca2þ]. An alternative approach to obtaining a background reading is to add MnCl2 (at a
concentration equal to or greater than the Ca2þ concentration in the medium) at the same time as the
digitonin so that compartmentalized dye can also be quenched as ionophores transport Mn2þ into the
organelles. Using such an approach assumes that cellular autofluorescence is the true background and
ignores the contribution of compartmentalized dye to the background.
136 Joseph P. Y. Kao et al.
4. A stable baseline is obtained quickly only if the dye released from cells by
digitonin permeabilization is swept rapidly away from the region directly above the
microscope objective. Otherwise, fluorescence from the released dye still will be
captured by the objective and, thus, contribute to the measured background. Once
swept away and diluted into the bulk medium, the released dye contributes
negligibly to the background.
0
A45 6
F�340
F�380
F�f,380
BG340
BG380
20
Flu
ores
cenc
e in
tens
ity (
a.u.
)
40 60
Time (min)
80 100 120
B
1 3
R=
F34
0/F
380
20
10
0
20 40 60 80 100
6
Rmax
Rmin
0 20 40
0.8
60
Time (min)
80 100 120
0.7
0.6
0.5
1 2 3
5
2
F�b,380
Fig.11 (Continued)
2.0C
1 2 3
1.0
0.00 10 20
Time (min)
30 40
[Ca2+
] i (m
M)
Fig. 11 Procedure for in situ calibration of intracellular Fura-2. (A) Fluorescence intensity traces
acquired at 340- and 380-nm excitation. Time marker arrow correspond to (1) addition of 50 nM
vasopressin; (2) exchange into Ca2þ-free phosphate-buVered saline (PBS) containing 10 mM MgCl2,
2 mM EGTA, pH 7.4; (3) addition of 10 mM ionomycin; (4) exchange into nominally Ca2þ-free saline,pH 7.4; (5) addition of 10 mM ionomycinþ20 mM CaCl2; and (6) 20 mM digitonin. Dotted lines mark
fluorescence levels corresponding to various parameters discussed in Section V.B. (a.u. = arbitrary
units). (B) F340/F380 ratio trace derived from the data in (A). Dotted lines markRmin andRmax (0.566 and
16.6, respectively, in this experiment). The parameter sf,2/sb,2 is 10.7. Inset. The portion of the trace from
20 to 118 min at higher resolution on the vertical scale to reveal the gradualness with which Rmin is
approached. (C) [Ca2þ]i trace derived from the ratio trace by using Eq. (2) in Section V.B. Only the first
40 min of the experiment are shown. This REF52 cell was incubated with 1 mMFura-2 AM in Pluronic
dispersion in HBSS for 90 min at 25 �C before being transferred to fresh HBSS for measurement.
5. Measuring [Ca2þ] with Fluorescent Indicators 137
Elevation of [Ca2þ]i by ionophore can lead to rapid cell lysis and loss of indica-
tor, sometimes before Rmax can be determined confidently. Almost paradoxically,
raising the extracellular [Ca2þ] to 10–30 mM (rather than just a few mM) in this
procedure appears, in some cases, to have a protective eVect on cell structure so
lysis is deferred and Rmax can be reached. If high extracellular [Ca2þ] is used, themedium should be free of phosphate salts, bicarbonate/carbonate, and even sul-
fate, since these ions can form precipitates with Ca2þ.Typical data from an experiment performed on a REF52 cell loaded with Fura-
2 are shown in Fig. 11. Shown in Fig. 11A are the two raw data traces, F 0340 and
F 0380, collected when the cell is excited alternately with 340-nm and 380-nm light.
The fluorescence signals measured after digitonin permeabilization are the back-
ground intensities, BG340 and BG380, that must be subtracted from the respective
traces to yield the true F340 and F380 (i.e., F340¼F 0340�BG340 and
F380¼F 0380�BG380). The ratio trace is simply a point-by-point division,
138 Joseph P. Y. Kao et al.
R¼F340/F380 and is shown in Fig. 11B. Rmin is the limiting value of R that is
reached during Ca2þ deprivation, whereas Rmax is the limiting value of R reached
after treatment with ionophore at high [Ca2þ].24 The factor sf,2/sb,2 is essentially
(F 0f,380�BG380)/(F 0
b,380�BG380). Using these experimentally derived parameters
and a predetermined Kd (224 nM; Grynkiewicz et al., 1985) in Eq. (2), one can
convert the F340/F380 ratio trace into a plot of [Ca2þ]i as a function of time
(Fig. 11C).
This procedure has the advantage that all spectroscopically derived parameters,
namely Rmin, Rmax, and sf,2/sb,2, that are especially sensitive to environmental
changes are determined in situ with the indicator residing in the intracellular
environment. Only the equilibrium dissociation constant is determined in vitro.
Rmin determined by Ca2þ deprivation is assumed to be the true value. In view of the
ineVectiveness of currently available ionophores at low [Ca2þ], one would be
justified in concluding that true Rmin would be diYcult to reach25 and that Rmin
is easy to overestimate. An overestimate of Rmin results in underestimation of
[Ca2þ].Finally, it is worthwhile to examine the eVects of errors in Rmin, Rmax, and
sf,2/sb,2 on the derived value of [Ca2þ]. For simplicity, one assumes that errors in
the three parameters are independent. Because sf,2/sb,2 is related linearly to [Ca2þ](see Eq. (2)), a percentage error in sf,2/sb,2 translates into the same percentage error
in [Ca2þ]. Inspection of Eq. (2) reveals that errors in Rmin should aVect primarily
low values of [Ca2þ] (corresponding to R values near Rmin). Error in Rmax, on the
other hand, aVects the way in which all the R values are scaled and, therefore,
should influence all derived values of [Ca2þ]. These expectations are borne out bycalculation.26
24 From Fig. 11B, the ratio values near Rmax are seen to oscillate significantly because, at saturating
[Ca2þ], the fluorescence of the indicator excited at 380 nm (Fb,380 ¼F 0b,380 � BG380) is very weak and
cannot be determined with high precision. In forming the ratio, because Fb,380 is a small number and
occurs in the denominator, noise fluctuations in Fb,380 become magnified into large-amplitude fluctua-
tions in Rmax. Therefore, one must average a large number of points to obtain a reliable estimate of
Rmax. Alternatively, the fluorescence intensity data (both F340 and F380) can be smoothed first before a
ratio is formed.25 Rather than estimating Rmin directly from the lowest values attained in the ratio trace, curve-
fitting the portion of the ratio trace that represents the slow descent towards Rmin is also a reasonable
approach. As expected, Rmin obtained by exponential curve-fitting is somewhat lower than that
estimated directly from the ratio trace.26 When one uses parameters similar to those for Fura-2 inREF52 cells as determined onour instrument
(Rmin¼ 0.5,Rmax¼ 15, and sf,2/sb,2¼ 12), a 10% overestimation ofRmin leads to�19% underestimation of
[Ca2þ] at 50 nM, �10% at 100 nM, and �2% at 500 nM. A 10% overestimation of Rmax leads to
underestimation of [Ca2þ] by �9.5% at 50 nM, �10.9% at 500 nM, and �12.5% at 1 mM. A 10% under-
estimation ofRmax results in overestimation of [Ca2þ] by�11.8% at 50 nM,�14% at 500 nM, and�16.5%
at 1 mM.
2.5
2.0
1.5
1.0
0.5
0 10
Time (s)
ΔF
F
F/F
0 o
r Δ
F/F
0
F0
F0
20 30
0.0
Fig. 12 Two conventions, F/F0 and DF/F0, for reporting fluorescence changes relative to baseline
fluorescence intensity. Note: DF/F0¼F/F0�1.
5. Measuring [Ca2þ] with Fluorescent Indicators 139
VI. Reporting Indicator Fluorescence Intensity Changeswithout Calibration
A. Reporting Relative Changes in Fluorescence: F/F0 and DF/F0
With the widespread use of nonratiometric indicators, which are diYcult to
calibrate, it has become common to report not [Ca2þ], but rather indicator fluores-cence changes. The convention is to report either the fluorescence intensity relative
to baseline intensity (F/F0), or the change in fluorescence intensity relative to
baseline intensity (DF/F0¼ (F�F0)/F0). Figure 12 illustrates these two conventions.
From the above definitions and from the graphs in Fig. 12, it is apparent that the
two reporting conventions are simply related: DF/F0¼F/F0�1. It is important to
stress that in order for these relative measurements to be meaningful, F and F0
should be intensities that have been background-subtracted.
B. Caveat in Interpreting Relative Fluorescence Changes: Indicator Fluorescence is Not aLinear Function of [Ca2þ]
Because a nonratiometric indicator becomes brighter when it binds Ca2þ, anincrease in indicator fluorescence implies an increase in [Ca2þ]. Once fluorescence
intensity data have been converted into relative changes, however, there is perhaps
140 Joseph P. Y. Kao et al.
a natural tendency to regard the relative change in intensity as reflecting an
equivalent relative change in [Ca2þ]. For example, a doubling of intensity relative
to baseline (F/F0¼2 or DF/F0¼1) is often used to infer a doubling of [Ca2þ]. Suchan inference should never be made because it is always incorrect. A quantitative
analysis is presented below.
As shown in Appendix 2, the total fluorescence, FT, emitted by a solution of
Ca2þ indicator is governed by the expression
FT / QCaIneCaInfCaIn þQIneIn 1� fCaInð Þ ð3Þwhere QCaIn and QIn are the fluorescence quantum eYciencies of the Ca2þ-boundand Ca2þ-free forms of the indicator, respectively, eCaIn and eIn are the extinctioncoeYcients of the two forms of the indicator at the excitation wavelength, and fCaInis the fraction of the indicator that is in the Ca2þ-bound form. Knowing that
fCaIn¼ [Ca2þ]/([Ca2þ]þKd), we can rewrite the expression to show its dependence
on [Ca2þ] more explicitly:
FT / QIneIn þ QCaIneCaIn �QIneInð Þ Ca2þ� �
Ca2þ� �þKd
ð4Þ
The only variable in the expression is [Ca2þ]; all other parameters, being intrinsic
characteristics of a particular indicator, are constants. The above expression shows
that whereas [Ca2þ] can range from 0 to any arbitrary positive value, the total
fluorescence, FT is bounded. This behavior is shown in Fig. 13.When [Ca2þ]¼0, all
of the indicator is Ca2þ-free, and the fluorescence has a minimum value that
depends on the intrinsic brightness (QIneIn) of the Ca2þ-free form of the indicator.
At saturating [Ca2þ] ([Ca2þ]Kd), all of the indicator is Ca2þ-bound, and the
fluorescence has a maximum value that depends on the intrinsic brightness
(QCaIneCaIn) of the Ca2þ-bound form of the indicator. Once the indicator molecules
are saturated, further increasing [Ca2þ] brings no increase in fluorescence. There-
fore, as can be seen from Eq. (4) and Fig. 13, fluorescence intensity is a nonlinear
function of [Ca2þ]. This nonlinearity is the reason that a relative change in indica-
tor fluorescence does not imply an equal relative change in [Ca2þ]. Figure 13 showsthat the discrepancy depends on the extent to which the indicator is already bound
to Ca2þ: Starting from a relatively low [Ca2þ], increasing [Ca2þ] by an increment,
DCa1, results in a fluorescence increase, DF1. From the now-higher [Ca2þ], a
further identical increment of DCa2 (¼DCa1) brings a much smaller fluorescence
increase, DF2.
The error in using relative fluorescence changes to infer relative [Ca2þ] changescan be analyzed quantitatively for a specific example. Fluo-4 is a nonratiometric
indicator that is commonly used with 488-nm excitation. The extinction coeYcient
of Fluo-4 changes only by a few percent upon binding Ca2þ
(eIn� eCaIn¼77,000 M�1 cm�1 at 488 nm); the Ca2þ-bound form is at least 100
times more fluorescent than the Ca2þ-free form (QCaIn¼0.14, QIn�0.0014); and
Kd¼345 nM. The quantitative relationship between [Ca2þ] and fluorescence can
Upper bound ∝ QCaIn εCaIn
ΔF2
ΔCa 1
ΔCa 2
ΔF1
[Ca2+]
00
FT
Lower bound ∝ QIn εIn
Fig. 13 Indicator fluorescence intensity is a nonlinear function of [Ca2þ]. At [Ca2þ]¼0, all indica-
tor molecules in solution are in the Ca2þ-free form, and indicator fluorescence is at the lower
bound. Whereas [Ca2þ] can range from 0 to any arbitrarily large value, indicator fluorescence
cannot exceed an upper bound, which is reached when all indicator molecules in solution are in
the Ca2þ-bound form. The relationship between fluorescence intensity and [Ca2þ] is hyperbolic.
The consequence is that successive equal increments in [Ca2þ] do not result in equal increments of
fluorescence intensity (compare the fluorescence increments DF1 and DF2 resulting from two equal
increments in [Ca2þ]).
5. Measuring [Ca2þ] with Fluorescent Indicators 141
be calculated by using these parameters in Eq. (4). Figure 14 shows the relative
change in Fluo-4 fluorescence for diVerent increments in [Ca2þ], up to a 10-fold
change ([Ca2þ]/[Ca2þ]0¼10). Because resting [Ca2þ]i is typically in the range 50–
100 nM, the starting [Ca2þ] was assumed to be [Ca2þ]0¼75 nM for the calculation.
Figure 14 shows clearly that the relative change in fluorescence is never a good
measure of the true relative change in [Ca2þ]. F/F0 significantly underestimates
[Ca2þ]/[Ca2þ]0, and the error increases severely as the change in [Ca2þ] becomes
larger.
VII. Measuring [Ca2þ] in Mitochondria
As mentioned in Section II, when cells are incubated with the AM ester of
Rhod-2, the indicator preferentially loads into mitochondria. The structures of
two fluorescent dyes, TMRM and TMRE, which also accumulate preferentially
into mitochondria, and Rhod-2 AM are shown in Fig. 15A. The positively charged
structure in these molecules that enables preferential loading into mitochondria
is highlighted with thick lines in Fig. 15A. Figure 15B shows that, rather than
A
B
Fig. 15 (A) Structures of Rhod-2 AM (bromide salt), as well as TMRM and TMRE (percholorate
salts), two dyes that accumulate preferentially into mitochondria. Note that in each case, the
dye molecule bears a permanent positive charge. (B) A series of related resonance structures
showing that the positive charge can be located on diVerent atoms in the molecule; that is, the charge
is delocalized.
2
2
1
1
03
3
4
4
5
5
6
6
7
7
8
8
9
9
10
10
2
1
0
3
4
5
6
7
8
9
10
[Ca2+]/[Ca2+]0
FF0
[Ca2+]0= 75 nM
[Ca2+]
[Ca2+]0
Fig. 14 Specific example illustrating that a relative change in fluorescence (F/F0) of the indicator,
Fluo-4, does not accurately reflect the true relative change in [Ca2þ] ([Ca2þ]/[Ca2þ]0).
5. Measuring [Ca2þ] with Fluorescent Indicators 143
being isolated on a single atom, the positive charge can reside on many
diVerent atoms in the structure—that is, the positive charge is delocalized over
the entire highlighted structure. A hydrophobic organic ion whose charge
is delocalized can pass through lipid membranes. Mitochondria maintain a re-
markably negative membrane potential—the mitochondrial lumen is typically at
�150 to �200 mV relative to the cytosol. Therefore, Rhod-2 AM, which is a
hydrophobic organic cation whose positive charge is delocalized, can permeate
through the plasma membrane and the mitochondrial membranes and preferen-
tially partition into the negative lumen of mitochondria. In mitochondria, cleavage
of AM ester groups by esterases liberates the Ca2þ-sensitive form of Rhod-2 -
(bearing multiple nondelocalized negative charges), which is not membrane-per-
meant and thus trapped in the mitochondrial lumen. Therefore, Rhod-2 can be
used to monitor intramitochondrial Ca2þ signals (Babcock et al., 1997; Tsien and
Bacskai, 1995).
A. Estimating the Fraction of Intracellular Rhod-2 Indicator that Resides in Mitochondria
While Rhod-2 can be preferentially loaded into mitochondria, the discrimina-
tion against loading into other subcellular compartments is imperfect. Some
intracellular Rhod-2 is expected to reside in the cytosol and nonmitochondrial
organelles. A simple procedure based on diVerential permeabilization of cellular
membranes can be used to estimate the fraction of intracellular Rhod-2 that
actually resides in mitochondria. The procedure is a modification of the one
described in Section III.B.2. The procedure consists of monitoring total Rhod-
2 fluorescence from a cell or a group of cells bathed in low-Ca2þ medium and
treating the cells sequentially with (1) a Ca2þ ionophore (ionomycin or Br-
A23187), (2) the mild detergent digitonin to permeabilize the plasma membrane,
and (3) a strong detergent, for example, Triton X-100 or sodium dodecyl sulfate
(SDS), to permeabilize all membranes. Figure 16 shows the procedure being
applied to a vagal sensory neuron that had been incubated with 1 mM Rhod-
2 AM for 1 h at room temperature. Application of ionomycin abolishes significant
diVerences in [Ca2þ] between diVerent subcellular compartments. This ensures that
Rhod-2 in all compartments is at comparable levels of Ca2þ-binding, and thus
would contribute fluorescence intensity in proportion to their actual content in
each compartment. Once the fluorescence reaches a steady baseline after ionomy-
cin treatment, digitonin permeabilization of the plasma membrane allows cytosolic
Rhod-2 to escape,27 giving a decrement in total fluorescence (labeled ‘‘C’’ in
Fig. 16). Subsequent permeabilization of all cellular membranes by SDS allows
27 Digitonin treatment leads to release of Rhod-2 from the nucleus as well. The nuclear pores have a
size exclusion limit of 35–40 kDa; molecules with molecular mass less than the exclusion limit can freely
exchange between the nucleoplasm and cytosol. Therefore, with respect to low-molecular-mass solutes
such as simple ions (e.g., Ca2þ, Naþ, Cl�) and small organic molecules (e.g., glucose, ATP, fluorescent
indicators), the nucleo-cytoplasm functions as a single ‘‘cytosolic’’ compartment.
Iono
60 s
C
M + NM
Rho
d-2
fluor
esce
nce
(arb
itrar
y un
its)
DigitSDS
0-Ca/BAPTA
Fig. 16 A diVerential permeabilization experiment for estimating subcellular fractions of Rhod-2
indicator. A rabbit vagal sensory neuron was incubated with 1 mM Rhod-2 AM for 1 h at 23 �C and
then bathed in nominally Ca2þ-free physiological saline to which 2 mM Na4BAPTA was added (0-Ca/
BAPTA). The total Rhod-2 fluorescence from the cell was monitored. Ionomycin (Iono, 2 mM) was
applied to dissipate Ca2þ gradients between subcellular compartments. Digitonin (Digit, 20 mM)
permeabilized the plasma membrane selectively to release cytosolic Rhod-2 (intensity decrement
marked ‘‘C’’). Sodium dodecyl sulfate (SDS; 0.25%, w/v) permeabilized all cellular membranes to
release Rhod-2 from organellar compartments (decrement marked ‘‘MþNM’’). The durations of
reagent applications are indicated by the bars at the bottom.
144 Joseph P. Y. Kao et al.
Rhod-2 to escape from mitochondria as well as nonmitochondrial organelles; this
causes a further decrement in fluorescence (labeled ‘‘MþNM’’ in Fig. 16). For
four neurons tested, the ratio of the noncytosolic fraction, MþNM, to the cyto-
solic fraction, C, was
MþNM
C¼ 5:36
This provides one required algebraic condition; a second condition is
MþNM þ C ¼ 1
that is, intracellular Rhod-2 must be in the mitochondria, in nonmitochondrial
organelles, or in the cytosol. Since there are three variables, a third algebraic
condition is required, and this can be obtained by performing the permeabilization
experiment on cells whose incubation with Rhod-2 AM had been done in the
presence of a protonophore (e.g., CCCP, FCCP, 2,4-dinitrophenol),28 which
28 It is advisable to use oligomycin (e.g., 10 mM), a blocker of the mitochondrial F1F0-ATP synthase,
in conjunction with the protonophore. In discharging the mitochondrial membrane potential, the
protonophore eliminates the Hþ electrochemical gradient that is used by the ATP synthase to generate
ATP. This causes the ATP synthase to run in reverse—as an ATPase—and rapidly deplete cellular ATP.
Oligomycin, by blocking ATPase action, helps to preserve the cellular ATP pool.
5. Measuring [Ca2þ] with Fluorescent Indicators 145
abolishes the mitochondrial membrane potential and thus eliminates the driving
force for preferential partition of Rhod-2 AM into mitochondria. In these cells
with depolarized mitochondria, the fluorescence decrement caused by digitonin
still represents loss of cytosolic Rhod-2 (C), but the decrement caused by SDS may
be attributed to Rhod-2 loss from nonmitochondrial organelles (NM). In four
neurons loaded in the presence of 5 mM CCCP,
NM
C¼ 0:623
which provides the last algebraic condition required to solve for the three
unknowns, C, NM, and M. Using all the three conditions together yields
C¼0.156, NM¼0.097, and M¼0.747—about 75% of intracellular Rhod-2 reside
in mitochondria, with �15% in the cytosol and �10% in other organelles.29
B. Minimizing Rhod-2 Loading in the Cytosol
Having a few percent of Rhod-2 residing in nonmitochondrial organelles is likely
to be unimportant. The luminal [Ca2þ] in these organelles (e.g., lysosomes, ER, etc.)
is not expected to changemarkedly. Therefore, the Rhod-2 fluorescence signal from
these organelles should not change significantly during an experiment and thus can
be considered operationally to be part of the background fluorescence. In contrast,
Rhod-2 in the cytosol, although much less than that in the mitochondria, may
contribute a contaminating cytosolic Ca2þ signal when one attempts to measure
mitochondrial [Ca2þ]. The cytosolic fraction can beminimized by taking advantage
of the ubiquitous cellular transporters that extrude organic anions from the cytosol
into the extracellular space (this is the process discussed in Section III.C). After they
have been incubatedwithRhod-2AMat room temperature, cells can be transferred
into medium containing no AM ester and incubated for 20–30 min at 37 �C to
accelerate extrusion of Rhod-2 from the cytosol. Thereafter, the Ca2þ-responsiveRhod-2 signal should be predominantly mitochondrial.
29 This likely underestimates mitochondrial loading, because we assumed that in the presence of
protonophore, Rhod-2 loading into mitochondria was negligible. Without the benefit of the mitochon-
drial membrane potential, Rhod-2 AM can still diVuse into the depolarized mitochondria and be
processed by esterases therein. Therefore, the decrement caused by SDS should contain both contribu-
tions from nonmitochondrial organelles and from passively loaded, depolarized mitochondria. If we
assume that on a per-volume basis, the cytosol and mitochondria have comparable capacity to process
AM esters, then the Rhod-2 content in depolarizedmitochondria relative to that in the cytosol should reflect
the ratio of the mitochondrial and cytosolic volumes. Depending on cell type, mitochondria-to-cytosol
volume ratio can range from �1/5 to �1/3. Using the lower figure changes the third algebraic condition to
(C/5þNM)/C¼ 0.623, leading to themodified estimates:C¼ 0.156,NM¼ 0.066, and M¼ 0.778.Using the
higher figure of 1/3 leads to the condition, (C/3þNM)/C¼ 0.623, which yields C¼ 0.156, NM¼ 0.045, and
M ¼ 0.798. Comparing these numbers with those obtained in the main text shows that this additional
correction only changed the estimate by a few percent.
Caf
60 s
0-Ca
MitoCyto
ΔF/F
0=
0.5
Fig. 17 Simultaneous measurement of Ca2þ signals in the cytosol and in mitochondria. A rabbit vagal
sensory neuron was incubated with 1 mM each Rhod-2 AM and Fluo-3 AM for 1 h at 23 �C and then
superfused with nominally Ca2þ-free physiological saline (0-Ca). Rhod-2 and Fluo-3 fluorescence,
excited at 543 and 488 nm, respectively, were imaged simultaneously by laser-scanning confocal micros-
copy. Data are represented as fluorescence change relative to baseline (DF/F0). A 5-s pulse of caVeine
(Caf, 10 mM) was delivered by superfusion. The durations of reagent applications are indicated by the
bars at the bottom.
146 Joseph P. Y. Kao et al.
C. Monitoring Cytosolic and Mitochondrial [Ca2þ] Simultaneously
Because indicators whose AM esters are uncharged load primarily into the
cytosol, while Rhod-2 preferentially loads into mitochondria, one can monitor
Ca2þ signals in the two compartments simultaneously. For measuring cytosolic
[Ca2þ], one should select an indicator whose excitation and emission wavelengths
do not interfere with Rhod-2 measurement. Since Rhod-2 is a rhodamine-based
indicator, a fluorescein-based indicator would be suitable for the cytosolic mea-
surement (e.g., members of the Fluo family of indicators). Figure 17 shows an
experiment where the cytosolic and mitochondrial Ca2þ signals are monitored
simultaneously in a vagal sensory neuron being stimulated with a brief pulse of
caVeine (an agonist that activates ryanodine receptor Ca2þ channels to release
Ca2þ from Ca2þ stores in the ER). The cytosolic and mitochondrial Ca2þ
transients have very diVerent decay kinetics: the time for the Ca2þ signal to
decay by 80% was t80%¼7.7 s in the cytosol and t80%¼65.1 s in mitochondria.
VIII. Concluding Remarks
Fluorescent Ca2þ indicators have contributed enormously to our understanding
of intracellular calcium regulation. For those who are beginning to use these
indicators, the technical details can seem bewildering. This compendium of
5. Measuring [Ca2þ] with Fluorescent Indicators 147
common techniques has aimed to set in order the body of practical empirical
knowledge that underlies successful measurements of [Ca2þ] through the use of
fluorescent indicators.
Appendix 1. The fraction of a polybasic acid that exists in aparticular state of protonation
The native forms of the Ca2þ buVers and indicators discussed in this chapter are
polybasic acids; that is, they are species with multiple dissociable protons (e.g.,
H4EGTA). Almost invariably, the fully deprotonated species is the one that
actually binds Ca2þ with high aYnity; therefore it is useful to be able to estimate
the fraction of the indicator or buVer in solution that actually exists in the fully
deprotonated form. In general, deprotonation of a polybasic acid is characterized
by a sequence of dissociation equilibria. For the specific example of a tetrabasic
acid, H4A, the sequence of stepwise dissociation reactions and the corresponding
equilibrium constants are:
H4A > Hþ þH3A� K1 ¼ Hþ½ H3A
�½ H4A½
H3A� > Hþ þH2A
2� K2 ¼Hþ½ H2A
2�� �H3A
�½
H2A2� > Hþ þHA3� K3 ¼
Hþ½ HA3�� �
H2A2�� �
HA3� > Hþ þA4� K4 ¼Hþ½ A4�� �
HA3�� �
If we define C0 to be the total concentration of the acid, irrespective of the state
of protonation, then the fraction that is present as the fully deprotonated,
tetraanionic form, A4�, is
a4 ¼A4�� �C0
ðA1:1Þ
To derive a useful expression for a4, we first recognize that the total concentration,C0, encompasses the concentrations of all the possible protonated forms:
C0 ¼ H4A½ þ H3A�½ þ H2A
2�� �þ HA3�� �þ A4�� � ðA1:2ÞBecause all the concentrations are related to each other through the dissociation
equilibrium constants,K1 throughK4, the concentration of any particular protonated
species can be written in terms of the concentration of any other species. In the
present case, it is convenient to express all the concentrations in terms of [A4�]:
148 Joseph P. Y. Kao et al.
HA3�� � ¼ Hþ½ K4
A4�� � ðA1:3Þ
H2A2�� � ¼ Hþ½
K3
HA3�� � ¼ Hþ½ 2K3K4
A4�� � ðA1:4Þ
H3A�½ ¼ Hþ½
K2
H2A2�� � ¼ Hþ½ 3
K2K3K4
A4�� � ðA1:5Þ
H4A½ ¼ Hþ½ K1
H3A�½ ¼ Hþ½ 4
K1K2K3K4
A4�� � ðA1:6Þ
By using these four relations, the expression for the total concentration can be
written in terms of [A4�]:
C0 ¼ Hþ½ 4K1K2K3K4
A4�� �þ Hþ½ 3K2K3K4
A4�� �þ Hþ½ 2K3K4
A4�� �þ Hþ½ K4
A4�� �þ A4�� �
ðA1:7ÞDividing through by [A4�] leads to
C0
A4�� � ¼ 1
a4¼ Hþ½ 4
K1K2K3K4
þ Hþ½ 3K2K3K4
þ Hþ½ 2K3K4
þ Hþ½ K4
þ 1 ðA1:8Þ
Writing the right side of Eq. (A1.8) as a fraction with a common denominator and
then inverting the fraction gives the desired final expression
a4 ¼ K1K2K3K4
Hþ½ 4 þ Hþ½ 3K1 þ Hþ½ 2K1K2 þ Hþ½ K1K2K3 þK1K2K3K4
: ðA1:9Þ
An important feature of Eq. (A1.9) to notice is that each term in the expression
actually represents the contribution of a particular protonated form, thus:
K1K2K3K4 , A4�
Hþ½ K1K2K3 , HA3�
Hþ½ 2K1K2 , H2A2�
Hþ½ 3K1 , H3A�
Hþ½ 4 , H4A
This insight makes it easy to write the fraction of the polybasic acid that is in a
particular form: the term representing the particular protonated form appears in
the numerator, while the denominator is simply the sum of all the possible terms.
For example, the fraction existing as HA3� is
a3 ¼ Hþ½ K1K2K3
Hþ½ 4 þ Hþ½ 3K1 þ Hþ½ 2K1K2 þ Hþ½ K1K2K3 þK1K2K3K4
; ðA1:10Þ
and the fraction existing in the doubly deprotonated H2A2� form is
5. Measuring [Ca2þ] with Fluorescent Indicators 149
a2 ¼ Hþ½ 2K1K2
Hþ½ 4 þ Hþ½ 3K1 þ Hþ½ 2K1K2 þ Hþ½ K1K2K3 þK1K2K3K4
ðA1:11Þ
The plots shown in Fig. 7 were generated using the above expressions for a2, a3,and a4 in conjunction with the four stepwise dissociation constants for EGTA.
In footnote 14, it was stated that Kd represents the ‘‘absolute’’ or intrinsic
dissociation constant characterizing the fully deprotonated form of the chelator
(e.g., A4� in the case above). At any pH where not all of the chelator is in the fully
deprotonated form, Kd must be corrected for the weakening eVect of acidic pH; the
corrected, or ‘‘conditional,’’ dissociation constant is K0d. As one would expect, the
correction factor is the fraction of chelator that exists in the fully deprotonated
form at the desired pH (e.g., a4 in the case of a tetrabasic acid like EGTA). Thus,
for a tetrabasic chelator,
K0d ¼
Kd
a4ðA1:12Þ
The plots shown in Fig. 8 were generated using Eq. (A1.12).
Appendix 2. Deriving an expression for the amount offluorescence emitted by a solution of fluorescent indicator
Light absorption by a solution containing a light-absorbing molecule, such as a
colorimetric or fluorescent indicator, is described by the Beer–Lambert Law:
A ¼ � logI
I0¼ elc ðA2:1Þ
whereA is the absorbance (or ‘‘optical density’’) of the solution, I0 is the intensity of
a light beam impinging on the solution, I is the intensity after the beam has passed
through the solution (I0� I¼ Iabs is the amount of light absorbed), e is the molar
extinction coeYcient (also known as themolar absorptivity), l is the thickness of the
solution throughwhich the light beam passes, and c is the concentration of the light-
absorbing molecule. The equation can be rearranged to the exponential form:
I ¼ I0e�2:303elc ðA2:2Þ
By convention, e has units of M�1cm�1 (i.e., l mol�1cm�1), l is measured in cm,
and c is measured in units of molarity (M, or mol l�1). For typical imaging
experiments where fluorescent indicators are loaded into cells, numerical limits
may be defined for the three parameters of interest:
e < 50; 000M�1 cm�1
l < 50� 10�4 cm
c < 100� 10�6M
150 Joseph P. Y. Kao et al.
The basis for these limits is as follows. First, indicators are small molecules whose
extinction coeYcients almost never surpass 50,000. Second, with extremely rare
exceptions, cells do not exceed 50 mm in diameter. Third, incubation with AM
esters usually achieves intracellular indicator concentration of several tens of
micromolar, and when indicators are introduced through a whole-cell patch elec-
trode, the concentration is usually kept below 100 mM to ensure that excessive
Ca2þ buVering capacity is not introduced into the cell. Applying these limits gives
2:303elc < 0:0575 � 1
Knowing that for x�1, e�x�1�x, we can recast Eq. (A2.2) as
I ¼ I0 1� 2:303elcð Þ ¼ I0 � 2:303elcI0 ðA2:3Þwhich rearranges to a simple expression of the amount of light absorbed by the
sample:
I0 � I ¼ Iabs ¼ 2:303elcI0 ðA2:4ÞThis approximate expression is accurate to within �1% for absorbances less than
0.06 (i.e., elc<0.06).
After absorbing a photon, a fluorescent molecule may reemit the absorbed
quantum of energy as fluorescence. The probability that after absorbing a photon,
a molecule will emit a photon of fluorescence is known as the quantum eYciency of
fluorescence. Quantum eYciency is usually symbolized as Q or f (Greek phi). The
amount of fluorescence emission should be the amount of light absorbed multi-
plied by the quantum eYciency:
F ¼ 2:303QelcI0 ðA2:5ÞIn this expression, only Q and e are intrinsic molecular properties, and these can
diVer substantially between the Ca2þ-free and Ca2þ-bound forms of the indicator
(symbolized as In and CaIn, respectively). For typical nonratiometric indicators,
QCaInQIn, while eCaIn� eIn. The total concentration of indicator, CT, is the sum
of the concentrations of the Ca2þ-bound and Ca2þ-free forms:
CT ¼ CaIn½ þ In½ ðA2:6ÞMaking use of the dissociation equilibrium constant:
CaIn > Ca2þ þ In Kd ¼ Ca2þ� �
In½ CaIn½
we can deduce the fraction of indicator that is Ca2þ-bound, fCaIn, and the fraction
that is Ca2þfree, fIn:
fCaIn ¼ Ca2þ� �
Ca2þ� �þKd
and fIn ¼ Kd
Ca2þ� �þKd
ðA2:7Þ
5. Measuring [Ca2þ] with Fluorescent Indicators 151
and, by necessity, fCaInþ fIn¼1. Therefore,
CaIn½ ¼ fCaInCT and In½ ¼ fInCT ¼ 1� fCaInð ÞCT ðA2:8ÞThe total fluorescence, FT, from a solution of indicator contains contributions
from both CaIn and In forms:
FT ¼ FCaIn þ FIn ¼ 2:303QCaIneCaInl CaIn½ I0 þ 2:303QIneInl In½ I0¼ 2:303QCaIneCaInlfCaInCTI0 þ 2:303QIneInlfInCTI0
¼ 2:303lCTI0 QCaIneCaInfCaIn þQIneInfIn½ ðA2:9Þ
This shows that the total fluorescence depends on the intrinsic molecular
properties of In and CaIn and the relative abundance of the two forms:
FT / QCaIneCaInfCaIn þQIneInfIn ðA2:10ÞMoreover, because the product, Qe, is a composite measure of a molecule’s ability
to absorb light and then emit fluorescence, we may think of Qe as the ‘‘intrinsic
brightness’’ of a fluorescent molecule. The brightness contribution of each indica-
tor form to the total fluorescence is weighted by the relative abundance of each
form. Finally, because fCaInþ fIn¼1, we can write
FT / QCaIneCaInfCaIn þQIneIn 1� fCaInð Þ ðA2:11Þ
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CHAPTER 6
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
Genetically Encoded Probes forMeasurement of Intracellular Calcium
Michael WhitakerInstitute of Cell and Molecular BiosciencesMedical School, Newcastle University, Framlington PlaceNewcastle upon Tyne, United Kingdom
A
OGY,All rig
bstract
VOL. 99 0091hts reserved. 153 DOI: 10.1016/S0091
-679X-679X
I. In
troduction II. G enetically Encoded SensorsA.
The Cameleon Family B. Camgaroos C. Pericam G-CaMP FamilyIII. A
pplications of Genetically Encoded Sensors A. Targeting to Subcellular Locations B. Tissue-Specific ExpressionIV. U
se of Genetically Encoded Calcium Sensors V. C onclusionsR
eferencesAbstract
Small, fluorescent, calcium-sensing molecules have been enormously useful in
mapping intracellular calcium signals in time and space, as chapters in this volume
attest. Despite their widespread adoption and utility, they suVer some disadvan-
tages. Genetically encoded calcium sensors that can be expressed inside cells by
transfection or transgenesis are desirable. The last 10 years have been marked by a
rapid evolution in the laboratory of genetically encoded calcium sensors both
figuratively and literally, resulting in 11 distinct configurations of fluorescent
proteins and their attendant calcium sensor modules. Here, the design logic and
performance of this abundant collection of sensors and their in vitro and in vivo
use and performance are described. Genetically encoded calcium sensors have
proved valuable in the measurement of calcium concentration in cellular
/10 $35.00(10)99006-7
154 Michael Whitaker
organelles, for the most part in single cells in vitro. Their success as quantitative
calcium sensors in tissues in vitro and in vivo is qualified, but they have proved
valuable in imaging the pattern of calcium signals within tissues in whole animals.
Some branches of the calcium sensor evolutionary tree continue to evolve rapidly
and the steady progress in optimizing sensor parameters leads to the certain hope
that these drawbacks will eventually be overcome by further genetic engineering.
I. Introduction
Small, fluorescent, calcium-sensing molecules have been enormously useful
in mapping intracellular calcium signals in time and space, as chapters in this
volume attest. Despite their widespread adoption and utility, they suVer some
disadvantages.
All low molecular mass fluorescent cytoplasmic calcium sensors are highly
charged molecules, so cross the cell’s plasma membrane very poorly. They are
placed into the cytoplasm by microinjection using fine-tipped micropipette or a
patch clamp pipette in whole cell mode. This limits their utility. Cell-permeant
fluorescent calcium sensors can be made by masking the charged carboxylic groups
by forming acetoxymethyl (AM) esters. Once inside the cell, the ester bonds are
cleaved, trapping the sensor in the cell. It is straightforward to bathe cells in culture
with the aposensor at low concentration and these AM esters have been very
widely used. One major drawback of the method is that the calcium sensor finds
itself not only in the cytoplasm, but also in intracellular compartments such as the
endoplasmic reticulum (ER) (Silver et al., 1992). Calcium concentrations are
higher in the ER than in the cytoplasm, so this leads to a significant unwanted
fluorescence signal from sensor in the ER that makes interpretation of the true
cytoplasmic concentration changes diYcult. It is also very challenging to use low-
molecular-mass fluorescent calcium sensors in whole animals.
For these reasons, genetically encoded calcium sensors that can be expressed
inside cells by transfection or transgenesis are desirable. One such sensor is
aequorin, a calcium-sensing protein found in the jellyfish Aequoria victoria. Origi-
nally, aequorin was isolated as a protein from jellyfish and placed inside cells by
microinjection (Baker, 1978; Gilkey et al., 1978). More recently, a construct
encoding recombinant aequorin has been used to express the aequorin apoprotein
in cells directly (see Chapter 10). Aequorin is a luminescent molecule and at the
concentrations used inside cells emits relatively few photons compared to fluores-
cent molecules at appropriate excitation intensities (Varadi and Rutter, 2002b).
However, proteins that are fluorescent at the visible wavelengths best suited to
fluorescence imaging are relatively rare. As it happens, A. victoria also expresses
a fluorescent protein, green fluorescent protein (GFP), and it is the work that
has produced the variously colored versions of GFP that has improved our
knowledge of this fluorophore and led to recombinant fluorescent calcium sensors
(Tsien, 2010).
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 155
The first recombinant fluorescent calcium sensors were described by Tsien and
Persecchini in 1997 (Miyawaki et al., 1997; Persechini et al., 1997; Romoser et al.,
1997). They were based on a concatenation of a recombinant calcium-binding
domain with GFP-derived fluorescent protein pairs. This approach has bred a
family of these cameleon indicators, so called because they are based on a long
tongue-like interaction between calmodulin (CaM) and a binding peptide and
change color (Miyawaki et al., 1997). Later, when it was realized that the GFP
beta-can structure lent itself to circular permutation without loss of function
(Baird et al., 1999), insertion of a calcium-binding domain within the GFP
(Baird et al., 1999) or concatenated to new N- or C-terminals (Nakai et al., 2001)
led to a second family of calcium sensors based on the fluorescence of a single
GFP-derived molecule, the camgaroos, pericams and their relatives.
The last 10 years have been marked by a rapid evolution in the laboratory of
these two families and their relatives, both figuratively and literally, as random
mutagenesis and clonal selection in bacteria has on occasion been used to opti-
mize the properties of the sensors (Griesbeck et al., 2001). This rapid diversifica-
tion has generated not only continuing improvements in the performance of the
sensors, but also a plethora of choice. Reviews have been written to track progress
in the field (Barth, 2007; Demaurex and Frieden, 2003; Garaschuk et al., 2006;
Griesbeck, 2004; Hires et al., 2008; KotlikoV, 2007; Mank and Griesbeck, 2008;
Miyawaki, 2003a,b, 2005; Pozzan and Rudolf, 2009; Solovyova and Verkhratsky,
2002; Zacharias et al., 2000). Most of the new variants have first been tested by
their makers in living cells as proof of principle rather than to answer substantial
questions in biology. I shall first set out the evolution of this growing tribe of
genetically encoded calcium sensing probes, dealing with the two broad families
in turn and then describe their application and utility in various biological
settings.
II. Genetically Encoded Sensors
A. The Cameleon Family
1. Origins
The family founders were described in three papers that followed rapidly in
succession in 1997. Their conception was aided by previous work in which GFP
had been altered by directed mutagenesis to produce diVerent colored variants
with altered excitation and emission spectra (Heim et al., 1995). As an aside, these
diVerently colored variants are sometimes referred to collectively as GFPs, though
they are not green. Persechini’s group described a construct (FIP-CBsm) in which a
red-shifted excitation variant of GFP (RSGFP; Delagrave et al., 1995, hereafter
GFP) and blue fluorescent protein (BFP) are linked by a sequence that includes 17
amino acids from the calmodulin-binding domain of avian myosin light chain
kinase (MLCK). This novel protein indirectly senses calcium concentrations inside
156 Michael Whitaker
cells, as when calcium increases, endogenous calmodulin becomes activated and
binds to the MLCK calcium-binding domain. This in turn alters the disposition of
the attached GFPs and leads to changes in Forster resonance energy transfer
(FRET) between the blue and green proteins (Romoser et al., 1997).
FRET is the phenomenon on which the cameleon sensor family relies. It occurs
between closely apposed fluorophores that have overlapping emission and excita-
tion spectra (Jares-Erijman Elizabeth and Jovin Thomas, 2003). In this example,
the emission spectrum of BFP overlaps with the excitation spectrum of GFP. The
extent of FRET depends on the degree of overlap between the two spectra, the
orientation of the fluorescence dipoles and crucially, the distance between them.
There is a very steep sixth power relationship with distance, so the energy transfer
is very sensitive to distance between fluorophores over the range 1–10 nm (Jares-
Erijman and Jovin, 2003). Calmodulin binds to the helical MLCK sequence by
wrapping its two lobes around it (Ikura et al., 1992). In FIP-CBsm, the steric bulk
of the calmodulin molecule when it binds to the MLCK peptide linker forces the
BFP and RFP further apart and reduces FRET (Romoser et al., 1997). FRET can
be measured in a variety of ways (Jares-Erijman Elizabeth and Jovin Thomas,
2003; Visser et al., 2010), but conceptually the simplest method is to excite the
donor fluorophore, here BFP, and measure the emission of both the donor and the
acceptor, here GFP. FRET takes place by nonradiative energy transfer, so high
levels of FRET transfer energy from BFP to GFP, reducing BFP emission at
around 440 nm and increasing GFP emission at 510 nm. Calmodulin binding
reduces FRET, increasing emission at 440 nm and reducing emission at 510 nm.
These changes can be expressed as a ratio of emission at the two wavelengths, a
value independent of the concentration of the protein. In HEK-239 cells expressing
FIP-CBsm, ratio changes (F510/F440) of around three- to fourfold could be observed
after raising free intracellular calcium concentration with the calcium ionophore
ionomycin (Romoser et al., 1997).
FIP-CBsm relied on endogenous calmodulin to generate a calcium-sensitive
FRET signal between GFPs. Tsien’s construct concatenated Xenopus laevis cal-
modulin and an MLCK calmodulin-binding peptide, M13 (Ikura et al., 1992),
together between BFP and GFP and also in an analogous construct between two
other GFP variants, enhanced cyan fluorescent protein (ECFP) and enhanced
yellow fluorescent protein (EYFP). In this concatenated configuration, binding
of calcium to calmodulin causes it to loop back toward the M13 peptide (the
cameleon’s tongue) as it binds, reducing the distance between the two GFP
variants and enhancing FRET (Miyawaki et al., 1997). This study beautifully
exemplifies the power of the cameleon concept linked to selective mutagenesis:
the original BFP/GFP construct (cameleon-1) worked well in vitro, but did not
express suYciently in mammalian cells; the enhanced variant with mammalian
codon usage (EBFP/EGFP—cameleon-2) showed much improved expression, but
the best expression, brightness, and signal-to-noise data were seen with enhanced
cyan and yellow variants of GFP (ECFP/EYFP—yellow cameleon-2). These ben-
efits came, however, at the expense of a lower FRET change between calcium
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 157
bound and unbound forms (1.5 vs. 1.8 when expressed as a ratio of emission
wavelengths) and a greater pH sensitivity. Mutagenesis can also be applied to the
calcium-binding aYnity of the calmodulin moiety: calmodulin has two classes of
calcium-binding sites and site-directed mutations in either high-(K0d 70 nM) or low
(K0d 11 mM) aYnity sites give rise to constructs in which high-aYnity sites are
suppressed to give a monotonic binding curve (K0d 4.4 mM: cameleon-3) or low-
aYnity sites are altered to give an enhanced range over four orders of magnitude of
calcium concentration (K0ds of 83 nM and 700 mM: cameleon-4). The third dimen-
sion of modification adds signal tags to the constructs. Nuclear localization tags
gave cameleon-2nu and ER localization tags produced yellow cameleon-3er (K0d
4.4 mM) and cameleon-4er (K0ds of 83 nM and 700 mM).
Tsien’s seminal paper also exemplifies some challenges in the approach: on the
one hand, the complexities of permutation and combination of mutant variants
and their concomitant properties and on the other hand, the relatively low magni-
tude of FRET modulation by calcium over a very wide range of concentrations.
The subsequent proliferation of family members results from attempts to improve
brightness and dynamic range, but at the expense of adding to the combinatorial
complexity.
Persechini’s second sensor design also concatenated GFPs, MLCK peptide, and
calmodulin, though in diVerent order. A calmodulin whose EF hand calcium-
binding sites had been reversed in order (CN-CaM) was added to the FIP-CBsm
C-terminal to BFP to make FIP-CA (Persechini et al., 1997). This produced a
sensor with a monotonic FRET response and a K0d of 100 nM. Variants with lower
aYnities for calcium were obtained by mutating the MLCK calmodulin-binding
peptide sequence, rather than the calmodulin calcium-binding sites. As with FIP-
CBsm, calmodulin binding reduced FRET, the ratio (now expressed as F440/F510)
increasing approximately 1.7-fold over the calcium dynamic range. The interaction
was markedly pH sensitive in the range 6.5–7.5. This configuration of calmodulin
and calmodulin-binding peptide did not lead to later variants and appears to have
been an evolutionary dead end.
The cameleon family of calcium sensors is shown in Fig. 1.
2. Evolution
The EYFP in yellow cameleon-2 and-3 shows an apparent pKa of 6.9, so
contains a significant proportion of the protonated species at physiological pH
(Miyawaki et al., 1999). The protonated species does not participate in FRET
(Habuchi et al., 2002). As pH can vary by several tenths of a pH unit when cells are
stimulated; changes in pH would be read as changes in calcium ion concentration.
Two adjacent point mutations in EYFP (V68L and Q69k) lower the pKa to 6.1,
markedly reducing the pH sensitivity in the physiological range (Miyawaki et al.,
1999). Replacing EYFP with EYFP-V68L/Q69K abolished pH sensitivity above
pH 6.9 (Miyawaki et al., 1999). This substitution produces yellow cameleon-2.1
(YC2.1; K0ds for calcium: 100 nM and 4.3 mM) and yellow cameleon-3.1 (YC3.1;
Fig. 1 Schematic depiction of the diVerent classes of genetically encoded calcium sensors. EYFP and EGFP variants for individual
sensors are shown to the right, as are the identities of the red-emitting sensors.
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 159
K0d for calcium: 1.5 mM) with around a twofold diVerence in 528/476 nm emission
ratios in calcium-free and calcium-saturating conditions. Recalling that the calci-
um-dependent signal from FIP-CBsm relied on binding of endogenous calmodulin,
an obvious concern would be that YCs would be perturbed by such interactions
and also perhaps themselves perturb downstream calcium-signaling pathways.
In fact, EC50s for YC2.1 and YC3.1 stimulation of calmodulin-dependent phos-
phodiesterase were two to three orders of magnitude greater than for calmodulin
and the sensors were unperturbed by addition of 3 mM calmodulin. Of course, the
YC constructs will buVer calcium inside cells. This was tested by studying the
calcium oscillations induced in HeLa cells induced by addition of histamine. At a
YC3.1 concentration of 150 mM, calcium oscillations were evident whereas at
concentrations greater than 300 mM, oscillations were not seen, though the overall
magnitude of the response was little altered. The loss of oscillations suggests
calcium buVering. Below around 20 mM, the fluorescent signal was too faint to
give acceptable signal-to-noise ratios (Miyawaki et al., 1999). Thus, working YC
concentrations in the range 40–150 mM do not substantially perturb calcium-
dependent signaling mechanisms.
Yellow fluorescent proteins, besides being sensitive to pH, are more prone than
GFP to photobleaching and to quenching by biological anions such as chloride.
Because YFPs show such utility as one of the partners in the CFP/YFP FRET
couple, this defect is worth fixing. Mutagenesis by error-prone PCR and expression
in Escherichia coli uncovered a mutation to methionine in residue 69 that was much
more resistant to chloride quenching than EYFP-V68L/Q69K, twice as resistant to
photobleaching, with a pKa of 5.7 rather than 6.1 and of comparable spectral
properties including brightness (Griesbeck et al., 2001). This YFP is known as
citrine, and substituted for EYFP-V68L/Q69K as the FRET acceptor produced
the cameleons YC2.3 and YC3.3. These two cameleons express well at 37 �C, showa ratio change of around 1.5 to calcium over their dynamic range and are pH
insensitive down to around pH 6.5. To demonstrate the utility of YC3.3 in an
acidic compartment, it was targeted to the Golgi using an 81 residue N-terminal
construct from human galactosyl transferase type II. The sensor was saturated
when expressed in the Golgi, suggesting high resting levels of free calcium concen-
tration in this cellular compartment (Griesbeck et al., 2001).
The CFP/citrine couple was also used in an ER-targeted sensor, Cameleon
D1ER. Here, the rationale was to design a sensor based on the M13/CaM-biding
pair that would be insensitive to interaction with endogenous calmodulin (Palmer
et al., 2004), as had been reported (Hasan et al., 2004; Heim and Griesbeck, 2004).
The M13 and CaM were co-mutated to provide a binding pair that would not
interact strongly with endogenous calmodulin. Cameleon D1ER has a very wide
range of calcium sensitivity with K0ds of 0.81 and 60 mM, appropriate for ER
calcium sensing, and was successfully used in HeLa cells to monitor cytoplasmic
and ER calcium simultaneously in conjuction with Fura2 (Palmer et al., 2004).
The GFP family of proteins is remarkable in possessing a visible wavelength
fluorophore that is formed through an oxidation reaction involving adjacent
160 Michael Whitaker
amino acids (Tsien, 1998). Fluorescence develops relatively slowly when the pro-
tein is expressed in cells, the process of what is known as maturation taking tens of
minutes to hours; maturation is also temperature dependent, oxidation to form the
fluorophore being the rate-limiting step. Another potential diYculty with FRET-
based probes using the CFP/YFP partners is that maturation of YFP is substan-
tially slower than that of CFP, particularly at mammalian body temperatures
(Miyawaki et al., 1999), a very important consideration especially for expression
in transgenic mammals. If the YFP partner of the FRET couple matures more
slowly than the CFP partner, then the sensors dynamic range is compromised, as
mature CFP in a sensor that contains immature YFP will contribute to the 476 nm
emission in the absence of 528 nm emission from the same construct, so that the
overall population 528/476 emission ratio will be depressed as a function of the
proportion of disparately matured sensor constructs (as illustrated by the behavior
of YC6.1 discussed below; Evanko and Haydon, 2005). The F46L mutation in
YFP greatly accelerates oxidation to the mature fluorophore and four additional
point mutations contributed to create a construct that matured two orders of
magnitude faster than EYFP from a urea-denatured state (Nagai et al., 2002;
Rekas et al., 2002); because of its resulting brightness, this YFP construct was
given the name Venus. Venus also has a low pKa (6.0) and low sensitivity to
chloride, comparable to citrine in these respects (Griesbeck et al., 2001), though
it lacks citrine’s improved resistance to photobleaching. Substitution of Venus for
EYFP-V68L/Q69K resulted in a new rapidly maturing yellow cameleon (YC2.12).
Bright YC2.12 fluorescence was seen to develop rapidly after ballistic transfection
of Purkinje cells in cerebellar slices, though the fold ratio change after depolariza-
tion suggests that its dynamic range was not much altered from earlier family
members (Nagai et al., 2002).
The challenge of improving dynamic range was addressed systematically by
altering the orientation of the YFP fluorescence dipole relative to the CFP dipole
(Jares-Erijman and Jovin, 2003) to maximize FRET (Nagai et al., 2004). Changes
in orientation were achieved by circular permutation (see below, Section II.B.1) of
the Venus construct. The YC3.12-based construct with EYFP-V68L/Q69K sub-
stituted by circularly permutated Venus with a new N-terminal at Asp-173 (termed
YC3.60) showed the largest increase in fluorescence emission ratio dynamic range
between calcium free and calcium-bound forms in vitro: around 6.6-fold compared
to 2.1-fold for YC3.12. This large improvement in dynamic range was verified by
expression of each the two sensors in HeLa cells and challenge with ATP to raise
cytoplasmic free calcium levels (Nagai et al., 2004). This study also illustrates the
important point that altering the properties and conformation of the FRET
partners at the N- and C-terminals of the sensor can also alter the apparent calcium
activation characteristics of the calmodulin-M13 inner pair as measured by FRET.
YC3.60 showed a monotonic increase with calcium concentration, as would be
expected from a construct based on the monotonically increasing cameleon-3
(Miyawaki et al., 1997), but the apparent dissociation constant for YC3.60 is
0.25 mM, compared to 4.4 mM for cameleon-3. YC2.60, based on cameleon-2,
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 161
has a single high-aYnity K0d of 40 nM, compared to the two K0
ds of 70 nM and
11 mM of cameleon-2 (Miyawaki et al., 1997). YC4.6 has K0ds of 58 nM and
14.4 mM, compared to K0ds of 83 nM and 700 mM in cameleon-4. The YCX.60
series of cameleons show the rapid maturation and low pH and chloride sensitiv-
ities of their Venus forbears, YCX.12, and are the best performing native M13-
based cameleons to date.
Both citrine and the circularly permutated Venus (cpv) of the YCX.60 series were
used as alternative acceptors in a further series of cameleons based on Cameleon
D1ER (Palmer et al., 2006). Computational design of novel M13 and CaM-based
binding pairs led to Cameleons D2, D3, and D4 and D2cpv, D3cpv, and D4cpv,
oVering a wide range of calcium aYnities, good sensor dynamic range (the cpv series
comparable to YC3.60), and insensitivity to endogenous calmodulin. This cameleon
set showed good performance in reporting cytoplasmic and mitochondrial calcium
concentrations in HeLa cells and peri-plasmalemmal calcium concentrations in
hippocampal neurones when localized with the appropriate targeting sequences.
ECFP/EYFP-based cameleons require excitation at near-UV wavelengths.
It would be convenient to have FRET-based calcium sensors that can be excited
at visible wavelengths. One possibly solution is to use a FRET couple in which
GFP is paired with a red fluorescent protein. GFP-like red fluorescent proteins are
found in corals (Baird et al., 2000; Miyawaki et al., 2003b). However, they are less
tractable than GFP and its variants as they oligomerize, mature very slowly via a
green-emitting intermediate and in general, show low extinction coeYcients and
quantum yield (Miyawaki et al., 2003b). A GFP/RFP cameleon has been devel-
oped using a DsRed variant—a tandem dimer mutant (Yang et al., 2005). The
maturation rate is tens of hours and the emission ration change is less than 1.2-fold
when cells expressing the sensor are challenged with ionomycin (Yang et al., 2005).
3. Changing the Sensor Mechanism 1
Solution NMR showed that the calmodulin-binding peptide of calmodulin-
dependent kinase kinase (CKKp) has a diVerent relation to the two lobes of
calmodulin than M13 peptide (Truong et al., 2001). The structural modeling
suggested that the peptide might be concatenated in a recombinant construct
between the N- and C-terminal lobes of calmodulin. Calculations suggested that
if ECFP and EYFP-V68L/Q69K were attached to the N- and C-terminals of the
split calmodulin, then the distance between the fluorophores when calcium was
bound and the calmodulin interacting with its binding peptide might be less than
40 A, rather than the 50–60 A in M13-based YC2.1. Given the sixth power depen-
dency of FRET on distance between fluorescent dipoles (Jares-Erijman and Jovin,
2003), this approach promised an improvement of the dynamic range of the ratio
of fluorescence emission. The splitting of the N- and C-domains of calmodulin in
this construct (termed YC6.1) led to a monotonic calcium-binding curve with a K0d
of 110 nM, in some respects more suited to measurement of smaller changes in
intracellular free calcium concentration. While in the event, YC6.1 showed a more
162 Michael Whitaker
modest fold emission ratio change than predicted (2.1 vs. 1.4 for YC2.1 in parallel
experiments), the twofold change was expressed over a narrower range of calcium
concentrations (0.05–1 mM) in the physiologically relevant cytoplasmic range.
YC6.1 of course suVers from the pH and chloride sensitivity and the slow
maturation of its EYFP-V68L/Q69K fluorophore that we discussed above. Repla-
cing EYFP-V68L/Q69K with Venus (Evanko and Haydon, 2005) gives the sensor
VC6.1 (Venus cameleon 6.1: the nomenclature is confusing and unhelpful, given
that the Venus CaM–M13 cameleons are known as YC2.12 and YC3.12). VC6.1
shows a emission ratio change of around 2.1-fold between zero and saturating
calcium concentrations. Thus, as with substitution with Venus for EYFP-V68L/
Q69K to produce YC2.12 from YC2.1, dynamic range is not much altered, while
improvements in maturation and pH and chloride sensitivity are obtained. It
would be logical to develop a YC6 sensor that contains the circularly permutated
Venus used in YC2.6 and YC3.6 (Nagai et al., 2004); this would be predicted to
much improve the ratio dynamic range.
Small improvements in dynamic range for YC6.1 and VC6.1 can be obtained by
excluding from analysis cells that express a low resting YFP/CFP ratio (Evanko
and Haydon, 2005): the authors very reasonably suggest that this screens out cells
in which the YFP partner is less-mature relative to its CFP pair.
4. Changing the Sensor Mechanism 2
One potential disadvantage of calmodulin-based sensors is that calmodulin is a
near-ubiquitous protein with many binding partners. It is possible that calmodulin-
based sensors may suVer interference from binding partners when expressed in the
cytoplasm or other cellular compartments. While there is no direct evidence to
support this conjecture, it is nonetheless true that performance in vivo does not
always mirror the sensor properties demonstrated in vitro (Hasan et al., 2004; Heim
and Griesbeck, 2004). With this potential pitfall in mind, a sensor has been devel-
oped based on troponin C, a calcium-binding protein and close homologue of
calmodulin that is, however, expressed only in muscle. The approach was to
concatenate TnC with CFP and citrine (Heim and Griesbeck, 2004). While devel-
oping these CFP–TnC–citrine sensors, a variant strategy was pursued to concate-
nate TNI, a TnC-binding partner, alongside TnC by analogy with theM13 binding
partner of calmodulin in the classical cameleons; this was unsuccessful. The con-
structs showing the greatest change in FRET between calcium-free and calcium-
bound forms contained a chicken skeletal muscle TnC with an N-terminal 14
residue truncation, TN-L15, and a human cardiac TnC, TN-humTnC. TN-L15
showed a 140% change and TN-humTnC a 120% change, measured in the absence
of magnesium ion. At physiological (1 mM) magnesium concentrations, the
dynamic ranges were 100% and 70%, respectively. Apparent dissociation constants
were 470 nM for TN-L15 and 1.2 mMfor TN-humTnC. The TnCEF hand calcium-
binding sites in TN-L15 were mutated to give K0ds of 300 nM and 29 mM. The pH
sensitivities were similar to the other CFP/citrine-based sensors, with a reduction in
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 163
dynamic range below pH 6.8 and little eVect in the physiological range pH 6.8–7.3.
Calcium oV rates were similar to or slightly faster than that of YC2.3 (Heim and
Griesbeck, 2004). The TN-L15 sensor was targeted to the plasma membrane using
GAP43, Ras, or Synaptobrevin. In direct comparison with YC2.1 and YC 3.3, it
showed markedly greater sensitivity and no diminution of dynamic range.
Mutations to EF hands III and IV and substitution of citrine with a circularly
permutated variant, Citrine-cp174 produced a TnC-based sensor that showed no
magnesium dependence, a fourfold dynamic range and a K0d of 2.5 mM —TN-XL.
TN-XL has a very fast oV rate with a dominant component with a time constant of
142 ms (Mank et al., 2006). Expressed in Drosophila under a UAS/Gal4 neuronal
promoter, it showed response times to calcium signals at the neuromuscular junction
significantly faster thanother sensors—YC2.0,YC3.3, Inverse Pericam,G-CaMP1.3,
andG-CaMP1.6. Furthermutagenesis and rearrangement of the TnC domain gave a
higher aYnity variant, modestly named TN-XXL, that was capable of long-term
monitoring of individual neuronal responses in flies and mice (Mank et al., 2008).
B. Camgaroos
1. Circular Permutation of EYFP
Remarkably perhaps, the beta-can that surrounds the cyclized and oxidized
fluorophore is amenable to circular permutation, by which is meant the insertion
of a peptide linker between N- and C-terminals of the protein and the creation of a
newN- and C-terminal pair elsewhere in the sequence, in the loops that connect the
component beta-sheets and in the beta sheets themselves (Baird et al., 1999). As we
have seen, circular permutation of Venus led to YC2.60 and YC3.60, the two
cameleons with the largest emission ratio dynamic range (Nagai et al., 2004). The
discovery that N- and C-terminals of EYFP could be rearranged prompted the
discovery that a calcium sensor could be fashioned by insertion of calmodulin
within EYFP itself.Xenopus calmodulin was inserted between residues 144 and 146
of each of ECFP, EYFP, and EGFP. Each of these constructs was a calcium
sensor, with the EYFP insertion giving the largest calcium response. In calcium-
free conditions, the construct absorbs predominantly at 400 nm, while in calcium-
saturating conditions, the dominant absorption peak is at 490 nm. The 400-nm
absorption is due to the protonated form of EYFP and the 490-nm absorption to
the unprotonated form. As discussed in Section II.A.2, in EYFP, the protonated
species is not fluorescent (Habuchi et al., 2002), so the excitation spectrum shows a
single peak at 490 nm and both the excitation and emission spectra are strongly
dependent on calcium concentration, with around an eightfold increase in emission
intensity at saturating calcium concentrations. Calcium binding was monotonic
with an apparent dissociation constant of 7 mM. Calcium binding clearly shifts the
proportion of protonated and unprotonated forms at constant pH, so the pKa’s for
the two forms are diVerent: 10.1 and 8.9, respectively. Continuing the whimsical
tradition, this calcium sensor is termed Camgaroo-1, because it is yellowish, carries
164 Michael Whitaker
a smaller companion (the calmodulin) in a pouch, can bounce high in signal and
may spawn improved progeny (Baird et al., 1999) The increase in fluorescence
intensity after addition of histamine to Camgaroo-1 expressing HeLa cells was a
modest 40% and the characteristic calcium spiking activity was almost invisible, so
the sensor is not quite as bouncy as its name implies when sensing cytoplasmic free
calcium; however, addition of ionomycin caused an overall sevenfold increase in
fluorescence. The modest increase observed in response to histamine is almost
certainly due to the 7 mM K0d, high relative to the calcium increase from around
100 nM to 1 mM expected when histamine is added to HeLa cells.
Camgaroo-1 does not fold well at 37 �C and could not be targeted to intracellu-
lar organelles, for example, mitochondria (Baird et al., 1999). In an attempt to live
up to another of its attributes, the possibility that it may spawn improved progeny,
Camgaroo-1 was subjected to error-prone PCR mutagenesis (Griesbeck et al.,
2001); selection of the brightest clone after expression in E. coli revealed a point
mutation of residue 69 to methionine. This new sensor, Camgaroo-2, had
very similar calcium-binding properties and fluorescence dynamic range as
Camgaroo-1, but expressed far more brightly in HeLa cells grown at 37 �C. Theresponse to histamine a (5% fluorescence increase) was lower even than for Cam-
garoo-1, but targeting to mitochondria using the targeting sequence of subunit
VIII of cytochrome c oxidase was demonstrated. Mitochondrial calcium increases
that raised the resting fluorescence signal by about 70% were demonstrated in
response to histamine and subsequent addition of ionopmycin gave an overall 1.5-
fold increase in fluorescence signal (Griesbeck et al., 2001), lower than that ob-
served with cytoplasmic Camgaroo-2, perhaps because the resting mitochondrial
calcium concentration is higher than that of the cytoplasm.
Using a similar camgaroo-like strategy, the EF hand calcium-binding site was
introduced into EGFP between residues 144–145, 157–158, or 172–173 (Zou et al.,
2007). These Ca-G family sensors had extinction coeYcients and quantum yields
comparable to EGFP. They operate in the ratiometric mode and with excitation at
398 and 490 nm showed a sensor dynamic range of 1.8 at a 510-nm emission
wavelength. Comprising a single EF hand-binding site, the apparent dissociation
constants are in the millimolar range (0.4–2 mM) and are, therefore, suitable only
for monitoring high calcium environments such as the ER. They are markedly pH
sensitive, with a pKa of around 7.5. Expressed in the ER of HeLa and BHK-21cells,
they showed modest ratio changes in response to agonists (Zou et al., 2007).
C. Pericam G-CaMP Family
1. Pericams
In pursuit of the idea that the clefts introduced into the beta can structure by
circular permutation might make the fluorophore more accessible to solution
protons and so susceptible to structural changes brought about by reorientation
of concatenated peptides, Miyawaki’s group developed the pericam series of
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 165
sensors (Nagai et al., 2001). Circular permutation of EYFP-V68L/Q69K to give an
EYFP with Y145 as the N-terminal and N144 as the C-terminal (cpEYFP)
produced an EYFP variant that could be concatenated with M13 and calmodulin
(bearing the E104Q mutation that conferred a monophasic calcium-binding curve;
Miyawaki et al., 1997). The construct with calmodulin at the N-terminal (CaM–
cpEYFP–M13) showed no calcium-dependent properties, confirming the finding
reported for a cpGFP variant (Nakai et al., 2001), but the opposite concatenation
(M13–cpEYFP–CaM) gave a construct that showed threefold brighter 520 nm
fluorescence in high calcium media compared to calcium-free media when excited
at 485 nm. This construct was given the name pericam (from a circularly permuted
YFP and CaM—calmodulin). Pericam was the prototype from which three peri-
cams with enhanced features were developed. Flash pericam has three additional
point mutations that confer an eightfold increase in 520-nm fluorescence on
calcium binding. Flash Pericam is a single wavelength, nonratiometric indicator
with a K0d of 0.7 mM .Knowing that substitution of phenylalanine at residue 203 in
YFP conferred fluorescence on the protonated form, this mutation was introduced
into Flash Pericam. The result, Ratiometric Pericam, was a sensor whose emission
ratio at 520 nm when excited at 494 nm or 415 nm changes 10-fold between
calcium-free and calcium-saturating conditions with a K0d of 1.7 mM; this excita-
tion ratio sensor is functionally analogous to fura-2 (Grinkiewicz et al., 1985).
Further semirandommutagenesis of Ratiometric Pericam gave a single wavelength
construct whose fluorescence intensity at 513–515 nm decreased on calcium bind-
ing—Inverse Pericam (K0d; 0.2 mM). Two advantages of Inverse Pericam are that it
is bright and has excitation/emission characteristics similar to fluorescein; the latter
advantage it shares with Flash Pericam: these two YFP-based indicators are
functionally equivalent to the Fluo-3 and Fluo-4 single wavelength calcium sensors
(Gee et al., 2000; Kao et al., 1989; Minta et al., 1989). Expression in HeLa cells
showed that Ratiometric Pericam and Inverse Pericam expressed significantly
better at 37 �C than did Flash Pericam. Ratiometric Pericam gave a 2.5-fold
increase in excitation ratio emission after addition of histamine, while Flash and
Inverse Pericams oVer a �100% increase and decrease in signal, respectively, with
the same agonist. As might be expected from our earlier discussion of the camgar-
oos, the calcium-free and calcium-bound forms of all three pericams showed
diVerent pKa’s and all three have pH sensitive emissions in the physiological pH
range. Miyawaki showed a proof of principle that the excitation ratio-based
Ratiometric Pericam can be used in the context of confocal imaging (Shimozono
et al., 2002); recent confocal microscopes based on acousto-optical filters oVerturnkey solutions to excitation ratiometric imaging.
2. GCaMPs
Single wavelength nonratiometric sensors that use the same sensor strategy
as pericams but are based on circularly permutated GFP rather than EYFP
were developed at almost the same time as the pericams, their development
166 Michael Whitaker
preceding the pericams’ by a matter of months (Nakai et al., 2001). Both the CaM–
cpGFP–M13 and M13–GFP–CaM concatenations were tested: only the latter
showed significant calcium-sensing properties. Twenty-six variants of the M13–
N149cpGFPC144–CaM concatenate were tested and the variant that showed the
greatest fluorescence increase in HEK-239 cells after ATP addition was termed
G-CaMP (presumably for green fluorescent calmodulin protein). In HEK-239
cells, G-CaMP gave a 1.5-fold increase in fluorescence in response to ATP and a
fourfold increase in response to ionomycin. G-CaMP has very similar fluorescence
parameters to Flash Pericam, with an excitation maximum at 489 nm, an emission
maximum at 509 nm and a 4.5-fold increase in fluorescence on calcium binding
(cf. eightfold for Flash Pericam). The apparent dissociation curve was monotonic,
with a K0d of 0.24 mM. As with the camgaroos and pericams and for the same
reasons, the sensor signal is strongly pH dependent in the physiological range. The
association time constant for calcium binding was strongly calcium dependent and
varied from 250 ms at low calcium concentration to 2.5 ms at higher concentra-
tions; the dissociation time constant was 200 ms. G-CaMP expresses poorly at
37 �C. G-CaMP-expressing smooth muscle showed a response to rapid depolari-
zation of around 50%, with a time course comparable to that previously measured
with Fluo-3. Carbachol addition gave a 2.5-fold increase. pH was monitored in
these experiments and did not change (Nakai et al., 2001).
This first GCaMP family member, later designated GCaMP1, had very weak
fluorescence when expressed at physiological temperatures compared to GFP
itself. This was addressed by introducing two mutations V163A and S175G that
were known to improve the temperature-dependent maturation of GFP to give a
variant known as G-CaMP1.6 (Ohkura et al., 2005); this increased brightness
about 40-fold. However, these modifications did not lead to adequate maturation
above 30 �C. The G-CaMP construct was subjected to error-prone PCR mutagen-
esis and the clones fluorescing most brightly at 37 �C were selected (Tallini et al.,
2006). The two new mutations in the brightest clone were identified (D180Y and
V93I), but it also turned out that the RSET leader sequence that had been added to
facilitate purification of the expressed protein was essential for thermal stability at
37 �C. This construct, GCaMP2, is around 200 times brighter than G-CaMP1 at
37 �C (with an extinction coeYcient at 487 nm of 19,000 and a quantum yield of
0.93 with emission at 508 nm) and shows the same four- to fivefold increase in
fluorescence a saturating calcium concentrations when compared to calcium-free
conditions. Though not reported, it should be assumed that this sensor remains
pH-sensitive. GCaMP2 was expressed using tissue-specific promoters in transgenic
animals and calcium transients were detected in granule cells in cerebellar slices
(Diez-Garcia et al., 2005) and in isolated heart in vitro and in adult and embryonic
heart in vivo (Tallini et al., 2006). Some insight into the sensor mechanism of
GCaMP2 is aVorded by its crystal structure (Akerboom et al., 2009; Wang
et al., 2008).
Even so, in HEK293 cells, GCaMP2 fluorescence is still 100-fold lower than
GFP itself (Tian et al., 2009). HEK293 cell medium-throughput screening assays
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 167
were used to identify brighter GCaMP2 mutants; attention was also paid to
improving the sensitivity to small calcium changes through mutations of the
CaM EF hands and of the M13/CaM interaction domains. The upshot was
GCaMP3, with a dynamic range of 12, due to a twofold decrease in calcium-free
fluorescence and a 1.5-fold increase in calcium-saturated fluorescence relative to
GCaMP2, and a K0d of 0.66 mM (Tian et al., 2009).
3. Cases 12 and 16
The Case (presumably Calcium sensor) constructs were developed by analyzing
the linker sequences between M13 and cpEYFP/GFP and cpEYFP/GFP and
calmodulin and the three key residues 148, 145, and 203 in the pericams and
G-CaMPs (Souslova et al., 2007). Based on this analysis, constructs were made
containing the G-CaMP linker sequences and the cpEYFP derived from Ratio-
metric Pericam. Nine point mutants were made with alterations in both the linker
sequences and in the three key residues within cpEYFP. As expected, combinations
of Asp148 and Phe203 produced ratiometric indicators akin to Ratiometric Peri-
cam, while Asn or Glu at residue 148 combined with Phe203 had a single excitation
peak at 490 nm. The Glu148/Thr145 and Glu148/S145 variants showed a 14.5-fold
increase in 490-nm fluorescence between calcium-free and calcium-bound forms.
The E148/S145 variant of these pericam-G-CaMP hybrids was optimized for
folding at 37 �C using error-prone PCR, resulting in a variant with a 12-fold
dynamic range named Case12. Substituting Thr for Ser at the 145 position of
Case12 gave Case16, with a 16.5-fold dynamic range. The apparent dissociation
constant for both Cases 12 and 16 was 1 mM. Like the pericams and G-CaMP
sensors, the calcium-bound forms of Cases 12 and 16 (pKa 7.2)—and thus their
fluorescence—are aVected by any changes in pH within the physiological range.
III. Applications of Genetically Encoded Sensors
A. Targeting to Subcellular Locations
Low molecular mass fluorescent calcium sensors do make their way to intracel-
lular compartments (Silver et al., 1992) and can be used to measure calcium there,
but they are diYcult to target precisely (Varadi and Rutter, 2002b). One of the two
major advantages of genetically encoded calcium sensors is that chimeric con-
structs and signaling tags can target them specifically to subcellular locations.
Methods to achieve some of these specific localizations had already been developed
for GFP itself and for the calcium sensor aequorin (De Giorgi et al., 1996). The
ability to target cameleons YC-3er and YC-4er was demonstrated in the study in
which cameleons were first described (Miyawaki et al., 1997).
168 Michael Whitaker
1. Endoplasmic Reticulum
ER calcium concentrations have been measured using low molecular mass
calcium sensors and with aequorin (Solovyova and Verkhratsky, 2002), but it
seems fair to say that the cameleon-based sensors (YC-3er and YC-4er) have
given the best estimates of ER calcium concentration and turnover (Foyouzi-
Youssefi et al., 2000; Graves and Hinkle, 2003a,b; Varadi and Rutter, 2002a; Yu
and Hinkle, 2000). In summary, cameleon-based indicators have presented a
picture of the ER as an organelle with resting calcium concentrations in the
range 250–600 mM and a very active calcium turnover that depends very heavily
on the activity of the SERCA ATPase (Demaurex and Frieden, 2003). Transgenic
YC3.3er has been engineered to give tissue-specific expression in mouse pancreatic
beta cells (Hara et al., 2004). The interpretation of calcium changes in the ER
measured by cameleon indicators is tempered by the finding that pH changes
within the organelle may interfere with estimates of dynamic calcium concentra-
tion (Varadi and Rutter, 2004). Improved sensors for ER calcium are now avail-
able (Palmer Amy et al., 2004; Zou et al., 2007).
2. Mitochondria
Mitochondrial targeting of recombinant aequorin was achieved using the
N-terminal presequence of subunit VIII of cytochrome oxidase (Rizzuto et al.,
1992). The same targeting strategy was used to locate ratiometric pericam within
mitochondria (Robert et al., 2001) and to show that the pericam tracked beat to
beat calcium changes in cardiomyocytes, just as did aequorin. Cameleon probes
targeted to mitochondria were eVective only at very low expression levels
(Arnaudeau et al., 2001). In a comparison of mitochondrially targeted cameleon
(mtYC2), camgaroo-2, and Ratiometric Pericam (Nagai et al., 2001) in HeLa
cells, it was found that Ratiometric Pericam was the most reliable and faithful
of the sensors (Filippin et al., 2003). Mislocalization and poor expression of the
mitochondrially targeted YC2 sensor could be improved by inserting a tandem
repeat of the subunit VIII presequence as the targeting sequence (2mtYC2)
(Filippin et al., 2005). 2mtYC2 was used successfully to demonstrate calcium
handling by skeletal muscle mitochondria during contraction (Rudolf et al.,
2004). Insertion of a tandem targeting repeat was an ineVective strategy
for the preferred citrine or Venus variants (Filippin et al., 2005), but in contrast,
the D2cpv, D3cpv, and D4cpv cameleons (Palmer et al., 2006) functioned
well as mitochondrial calcium sensors when targeted with the cytochrome
oxidase tandem repeat (Palmer et al., 2006). These constructs are now the
recommended genetically encoded mitochondrial calcium sensors. An recent
overview of calcium sensor approaches in mitochondria is available (Pozzan
and Rudolf, 2009).
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 169
3. Peroxisome
Cameleon D3cpv was furnished with a modified peroxisome localization
sequence (D3cpv-KVK-SKL) to monitor calcium concentrations in this organelle
in HeLa cells in response to agonists or depolarization (Drago et al., 2008).
4. Golgi
The Citrine cameleon YC3.3 has been expressed in the Golgi using an 81 residue
N-terminal sequence from human galactosyl transferase type II (Griesbeck et al.,
2001); it was saturated, oVering no useful information but that the Golgi has a very
high resting calcium concentration.
5. Plasma Membrane
Sub-plasmalemmal calcium concentrations may diVer from those in bulk cyto-
plasm. Localized calcium concentrations around secretory vesicles were shown to
be higher than those in cytoplasm by using a phogrin chimera to target YC2 to
secretory vesicle membrane (Emmanouilidou et al., 1999). A number of targeting
strategies have proved successful in localizing sensors to the plasma membrane.
The cpVenus cameleon YC3.60 has been targeted using a Ki-Ras chimera (Nagai
et al., 2004). The TN-L15 sensor localized to the plasma membrane as GAP43,
Ras, or synaptobrevin chimeras (Heim and Griesbeck, 2004). Localization
can also be achieved with a myristoyl/palmitoyl N-terminal tag (Zacharias
et al., 2002), an approach that was used with the cameleon D series (Palmer
et al., 2006). A chimera of GCaMP2 and synaptotagmin (SyGGCamp2) has been
used to monitor synaptic calcium signals, in this case in vivo in zebrafish (Dreosti
et al., 2009).
B. Tissue-Specific Expression
The other major advantage of genetically encoded calcium sensors is tissue-
specific expression in intact organisms.
1. YC2.1
The first transgenic tissue-specific expression of genetically encoded calcium
sensors was demonstrated in plants. YC2.1 was expressed in Arabidopsis guard
cells of the leaf, first using a CaMV promoter (Allen et al., 1999) and then a guard
cell-specific det promoter (Allen et al., 2000), demonstrating that aspects of the
calcium-signaling response in guard cells were under diVerential genetic control.
YC3.1 was used in transgenic Aradidopsis plants to visualize calcium signals in the
pollen grain (Iwano et al., 2004).
170 Michael Whitaker
YC2.1 was expressed transgenically in Caenorhabditis elegans pharyngeal
muscle under the control of the pharyngeal-specific myo-2 promoter (Kerr
et al., 2000) and tracked calcium changes during pharyngeal pumping;
YC3.1 tracked temporal changes more faithfully than YC2.1, being the
faster sensor, but YC2.1 tracked calcium changes to basal level more faith-
fully than YC3.1, as might be expected from its lower K0d. Expression of
YC2.12 in C. elegans touch neurons under the control of the mec-4 promot-
er identified a role for specific ion channels in the touch response (Suzuki
et al., 2003).
The UAS/Gal4 tissue-specific expression system was used to express YC2.1 in a
subset of the antennal lobe projection neurones of Drosophila in order to study
odorant responses in the antennal lobe and mushroom body calyx in vivo
(Diegelmann et al., 2002; Fiala et al., 2002). Odorant-specific patterns of neuronal
excitation were seen in both the antennal lobe and the calyx. In the former, the
EYFP/ECFP emission ratio changes were 1.23�0.23% (mean and sem) and in the
latter 0.6�0.06%. In the antennal lobe, the changes in sensor signal were observed
in spatially restricted regions of around 10–30 mm diameter, the size of individual
glomeruli. These very small changes were nonetheless reproducible, with distinct
and reproducible patterns of activity from fly to fly associated with diVerentodorants.
The same UAS/Gal 4 technology was used to express YC2 in neurones of
larval Drosophila (ReiV et al., 2002) to the evolution of calcium signaling in
presynaptic terminals innervating larval muscle. A 28% emission ratio change
was measured in vivo during spike train stimulation of the neuromuscular junc-
tion and signals of this magnitude could be resolved in single synaptic boutons;
there were no detectable diVerences in neuromuscular junction physiology
between wild-type and transgenic larvae. This study illustrates the point that
targeted expression of genetically encoded sensors in individual neurones is for
some applications superior to the use of low molecular mass synthetic calcium
indicators, as the specificity of expression more than compensates for the loss of
brightness.
In a similarly mature use of YC2.1 sensor technology coupled to UAS/Gal4
transgenic expression, neuronal calcium measurement coupled with electrophysi-
ology was used to identify thermosensory neurones in the larval nervous system
in vivo (Liu et al., 2003). Changes in emission ratio of 10–50% were associated with
heating and cooling. A functional map of thermosensory neurones was generated
and it was found that neurones with diVerent temperature responses were anato-
mically segregated.
YC2.1 was also used in zebrafish to record the behavior of Rohon-Beard (RB)
neurones during the fish’s escape response (Higashijima et al., 2003). This careful
study started with transient expression of the YC2.1 transgene in the RB neurones to
show proof of principle before generating transgenic lines in which the calcium signals
in the RB neurons could be correlated with the escape response in conscious fish.
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 171
2. YC3.3er
YC3.3er (the citrine-based sensor) was expressed in the beta cells of transgenic
mice under the control of the mouse Insulin 1 promoter (Hara et al., 2004). The
sensor signal could be detected in isolated pancreatic islets and addition of thapsi-
gargin or carbachol gave the expected decrease in the 535/485 emission ratio.
3. Camgaroos and Inverse Pericam
UAS/Gal 4 expression was used to create transgenic Drosophila that expressed
camgaroos-1 and-2 in the mushroom bodies of adult brain (Yu et al., 2003).
Dissected fly brains were used. Camgaroo-2 fluorescence in the mushroom bodies
was much more intense than that of camgaroo-1, but the camgaroo-1 emission
ratio signal on potassium depolarization was more than double that of camgaroo-2
(38% vs. 14% in the mushroom body lobe and 83% vs. 28% in the mushroom body
itself ). It was shown that these increases were not due to changes in pH. Applica-
tion of the putative mushroom body transmitter, acetylcholine, causes ratio
changes of a few percent. In this setting, camgaroo-2, although brighter, showed
substantially lower ratio changes than camgaroo-1; it also underwent significantly
faster photobleaching.
Inverse pericam is an intensity-coded sensor that decreases its fluorescence as
calcium increases. Addition of DsRed2 to the C-terminal of inverse pericam
produces a ratiometric indicator whose 615/510 nm emission ratio increases as
calcium increases. This indicator (DsRed2-referenced inverse pericam (DRIP))
requires dual excitation and dual emission optics (Shimozono et al., 2004). The
DsRed2 fluorescence is a passive, calcium-independent signal that is proportional
to the concentration of the sensor and helps control for alterations in overall
fluorescence intensity due for example to movement artifacts. DRIP was expressed
transgenically in worms under the control of the myo 2 promoter that is specific for
pharyngeal muscle. Ratio changes of 30–40% were measured in worms undergoing
fast pharyngeal pumping.
After screening six sensors (flash pericam, inverse pericam,G-CaMP, camgaroo-2,
YC2.12, andYC3.12) for calcium sensitivity in stably transfected fibroblast cell lines,
the two with the greatest dynamic range (inverse pericam: �40% and camgaroo-2:
þ170%), together withYC3.12 that gave inconclusive results in the fibroblast expres-
sion screen but is optimized for expression at 37 �C, were used to generate transgenic
mice under the control of the TET expression system (Hasan et al., 2004); the TET
system allows tissue-specific expression by crossing the TET mice with mice expres-
sing the TET transactivator under tissue-specific control. TET sensor mice were
crossed with a line expressing the transactivator under the control of the alpha-
calmodulin/calcium dependent kinase II (aCamKII) promoter. All mice developed
normally. Five highly expressing lines were obtained out of 36 transgenic lines: two
YC3.12, two camgaroo-2, and one inverse pericam. Expression patterns in brain
slices and excised retina were analyzed by two-photonmicroscopy. They appeared to
172 Michael Whitaker
be mosaic, not mapping directly to the known patterns of aCamKII expression.
Neocortical expression could also be imaged through the thinned skull in anaesthe-
tized mice. Two photon fluoresence recovery after photobleaching suggested that as
much as half the fluorescence signal was immobile and this together with punctuate
staining patterns suggested that this immobile sensor fraction might be due to
interaction between the M13 and CaM moieties of the sensors and their normal
cellular targets. Cellular and synaptic stimulation of pyramidal neurones in
cortical slices using sharp and patch microelectroded gave 5–10% increases
in 535 nm fluorescence using wide field imaging and around 20–100% for cam-
garoo-2 and �30% for inverse pericam using two photon imaging. In the retina, a
ganglion cell subset was strongly labeled in YC3.1-expressing mice, but no light-
evoked responses were detected. In camgaroo-2 expressing lines, bleaching occurred
in the retina too quickly for measurements to be made. In one inverse pericam-
expressing mouse, 7 of 12 ganglion cells tested showed a transient decrease in
fluorescence attributable to a calcium increase in response to light. Sensors were
imaged in the olfactory bulb in vivo using wide fieldmicroscopy. Camgaroo-2 expres-
sing mice showed a 1–3% increase in response to odors, while inverse pericam gave
�8% decrease. Each distinct odor evoked a unique pattern of activity, similar odors
evoking similar patterns.
This thoughtful study established four main facts: around half of the transgeni-
cally expressed cameleon family sensor was immobile; this reduced sensitivity and
made quantitation of the calcium signals impossible; nonetheless, it was possible to
observe patterns of neuronal activity; YC3.12 was not an eVective transgenic
sensor. The study also reports unpublished experiments in which transgenic mice
expressing YC3.0 under the control of a b-actin promoter gave only 1–2% ratio
changes during wide filed imaging in cerebellar slices. The high proportion of
immobile sensor in transgenic animals remains for the moment inexplicable—it
was not seen in the stably transfected fibroblast lines.
4. GCaMP
G-CaMP (Nakai et al., 2001) was expressed in mice under the control of a
smooth muscle myosin heavy chain promoter and was expressed in vascular and
nonvascular smooth muscle (Ji et al., 2004). The signatures of inotropic (ion
channel) and metabotropic (InsP3-mediated) postsynaptic signaling could be dis-
tinguished in single excised smooth muscle cells.
In a set of experiments strikingly parallel to those with YC2.1 (Diegelmann
et al., 2002; Fiala et al., 2002), but using two photon imaging, G-CaMP was
expressed in a subset of projection neurones in Drosophila antennal lobe to
demonstrate that diVerent odorants activated specific patterns of glomeruli
(Wang et al., 2003). Individual glomeruli are diVerentially sensitive to a given
odorant and more are recruited as the odorant concentration is increased.
Increases of fluorescence of up to 50% (at 525 nm) were measured in responsive
glomeruli.
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 173
Transgenic expression of GCaMP2 has been achieved in mouse heart (Tallini
et al., 2006). The TET system was used: the GCAMP2 sequence was placed
downstream of a weakened a myosin heavy chain promoter (aMHC) and seven
tetO enhancer sequences to permit suppression of gene expression using doxycy-
cline. These mice were crossed with others with a hemizygous aMHC-tetracycline
transactivator allele. The doubly transgenic mice expressed GCaMP2 only in the
heart. Doxycycline suppression of the transgene was essential, as mice constitu-
tively expressing GCaMP2 from birth showed markedly enlarged hearts, a pheno-
type comparable to that seen in mice overexpressing calmodulin. This phenotype
was avoided entirely by administering doxycycline in utero and until 13–15 weeks
postpartum. Subsequent removal of doxycycline for up to 6 weeks led to no
detectable cardiomegaly. Robust GCaMP signals were present 4 weeks after
doxycycline removal.
Striking wide field fluorescence images of cardiac calcium transients in whole
mouse heart beating at up to 300 beats/min were obtained with anaesthetized,
ventilated open-chested mice, the first to be recorded under wholly physiological
conditions with the heart under normal load. As expected sympathetic stimulation
with isoproterenol markedly increased the calcium signal and also increased end
diastolic calcium concentration. Signal-to-noise ratios were good and it was possi-
ble to record very clean signals from a single pixel of the 100�100 pixel imaging
array (tens of microns). Using a photodiode array in isolated perfused heart,
signals from a membrane potential sensitive dye and from GCaMP2 were acquired
simultaneously. Association and dissociation kinetics of calcium were rapid (t¼14
and 75 ms, respectively) and unaltered in vivo. Comparison with a fast calcium dye
Rhod2 nonetheless showed that the rise and decay times of the GCaMP2 signal in
beating heart was around 45% slower, but with a three times greater dynamic
range. Calcium sparks could not be observed in isolated ventricular myocytes
expressing GCaMP2. GCaMP2 imaging in open-chested embros from embryonic
day 10 allowed the analysis of the development of the atrio-ventricular node
conduction pathway.
GCaMP2 fused to synaptotagmin localizes to synaptic boutons. It reports the
location of synapses in zebrafish in vivo and shows a linear response over a wide
range of action potential frequencies (Dreosti et al., 2009). It can report spiking
frequencies in optic tectum; it also reports activity in the graded synapses of retinal
bipolar cells. GCaMP2 has also been used to map functional connections in the C.
elegans nervous system (Guo et al., 2009). Connections can be mapped grossly, but
the sensor’s signals are too weak to distinguish direct from indirect connections.
5. TN-L15, TN-XL, and TN-XXL
Acerulean version of TN-L15, cerTN-L15, was used to create a transgenicmouse
line that expressed the sensor widely in brain, especially in the neocortex and
hippocampus (Heim et al., 2007). Calcium changes resulting from two to three
action potentials could be resolved and calcium responses in spiny dendrites of
174 Michael Whitaker
pyramidal cells could be detected after puYng on glutamate, an excitatory neuro-
transmitter (Garaschuk et al., 2007; Heim et al., 2007). TN-XLwas expressed using
the UAS/Gal4 tissue-specific expression system in Drosophila neuromuscular
junction (Mank et al., 2006). Its rapid oV rate for calciummade it significantly better
at tracking calcium changes than its counterparts. TN-XXL showed improved
sensitivity and long term-stability in sensing calcium signals from fly neurones; in
mice, tuning curves for orientation-specific neurones in visual cortex could be
monitored repeatedly over timescales of days or weeks (Mank et al., 2008).
6. Comparison of the Performance of Genetically Encoded Calcium Sensors
Though progress in the field has been periodically reviewed (Barth, 2007;
Garaschuk et al., 2007; Griesbeck, 2004; Mank and Griesbeck, 2008), few studies
have systematically compared the performance of diVerent genetically encoded
calcium sensors, except to demonstrate the superiority of a novel sensor over its
predecessors. I have discussed above (Section III.B.3) the systematic comparisons
of camgaroo-1 and-2 when expressed in Drosophila mushroom bodies (Yu et al.,
2003) and of inverse pericam, camgaroo-2 and YC3.1 when expressed in mouse
brain (Hasan et al., 2004).
The performance of GCaMP, inverse pericam, and camgaroo-2 was compared
with that of the low molecular mass synthetic indicators X-Rhod-5F and Fluo4-
FF in apical dendrites of pyramidal cells in hippocampal brain slices from 6- to 7-
day-old rats transfected using a biolistic approach and maintained at room
temperature (Pologruto et al., 2004). Images were obtained using two-photon
microscopy. Action potentials were triggered using current injection into the cell
body. Under these conditions, X-Rhod-5F and Fluo4-FF could detect calcium
changes (signal twice that of noise) in the dendrite due to voltage-dependent
calcium channel activation after single action potentials while with the same
criterion GCaMP required five action potentials, camgaroo-2, 33 action potentials,
and inverse pericam over 20. For comparison, the dynamic ranges (DF/F) for thethree sensors under these conditions in vitro was1.8, �2, and �0.25, so the
sensitivity of camgaroo-2 was poor despite its larger dynamic range. Power spec-
trum analysis was used to analyze the fluorescence response during action potential
trains at 20 Hz. Most of the power in the frequency analysis of X-Rhod-5F and
Fluo4-FF fluorescence was at the fundamental frequency, 20 Hz, indicating that
the fluorescence signal tracked each action potential. For the genetically encoded
sensors, no clear peak was observed at 20 Hz, indicating that the sensors were too
slow to track individual action potentials at this stimulation frequency.
It was possible to measure calcium activation curves in situ for the three sensors
and thus their apparent dissociation constants by simultaneously measuring
calcium concentration using a calibrated X-Rhod-5F signal and the fluorescence
signal from the sensor at various levels of stimulation. For inverse pericam (K0d
0.9 mM) and camgaroo-2 (K0d 8 mM), these were comparable to those previously
reported in vitro; however, GCaMP showed a K0d (1.7 mM) almost an order of
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 175
magnitude greater than that previously reported in vitro (Nakai et al., 2001).
Because the calcium concentration profile of dendritic action potentials is well
characterized (Pologruto et al., 2004), there seems little doubt that the calcium
dissociation characteristics of GCaMP vary markedly in vitro and in vivo. FRAP
studies in dendrites showed that the all three sensors were mobile, with mobilities
comparable to GFP itself. This result is quite firmly at odds with that reported in
mouse brain (Hasan et al., 2004) and discussed above (Section III.B.3). It may be
that dendrites, being relatively free of organelles, mirror better the behavior of the
sensors in cytoplasm than cell bodies; it should be noted that punctuate staining
was reported in mouse brain (Hasan et al., 2004). It should also be borne in mind
that though the observations on mouse brain slices were carried out at room
temperature, as were these experiments in rat brain slices, in the mouse study,
the sensors had been expressed at body temperature, whereas the biolistically
transfected rat brain slices were maintained throughout at room temperature.
These data, as the authors point out (Pologruto et al., 2004), demonstrate that
the genetically encoded sensors are better-suited to measuring summated neuronal
responses after multiple stimuli, not single action potentials, consistent with their
reported use to monitor patterns of neuronal activity (Fiala et al., 2002; Hasan
et al., 2004; Wang et al., 2003); as it happens, these three studies all described
odorant-specific patterns of neuronal signaling.
As an addendum to the study, Svoboda’s group also provided in vitro solution
X-ray scattering evidence that showed that the calcium-dependent fluorescent
signal of GCaMP, as theorized, depends on a coupled structural change in which
calcium binding to CaM is closely linked to binding of CaM to M13; in contrast,
the calcium-dependent fluorescence signal in camgaroo-2 is solely due to binding
to CaM, the M13 peptide paradoxically playing no part in the sensor response
(Pologruto et al., 2004).
A second comparative study was undertaken at the Drosophila larva neuromus-
cular junction (ReiV et al., 2005), using an approach previously reported (ReiVet al., 2002). The responses of 10 sensors from the three families to 40 and 80 Hz
stimulation of the synaptic bouton were compared. Camgaroos-1 and-2 and flash
pericam did not sense calcium changes in the bouton. YC2.0, 2.3, 3.3, TN-L15,
inverse pericam, and GCaMP1.3 and 1.6 all showed adequate responses (around
5% on average at 40 Hz and 10–15% at 80 Hz) to pulse train stimuli, but none
exhibited dynamic ranges anywhere near comparable to those measured in vitro
(ReiV et al., 2005). None was comparable in performance in this system when
compared to the later developed TN-XL (Mank et al., 2006). In an echo of the
work in rat brain slices, the performance of YC3.3, TN-L15, GCaMP1.6,
GCaMP2, YC2.60, YC3.60, cameleon D3, and TN-XL were compared one with
another and calibrated against a low molecular mass indicator, Oregon-Green-
BAPTA-1 (Hendel et al., 2008). The latter four sensors were around twofold more
responsive than their earlier counterparts. None of the sensors were seen to detect
single action potentials, though YC3.60 and cameleon D3 could detect two action
potentials in succession. None showed the fast temporal response of the low
176 Michael Whitaker
molecular mass indicator. A theoretical framework in which to consider the pros
and cons of calcium sensors in recording neuronal activity has been adumbrated
(Hires et al., 2008).
GCaMP1.6 and GCaMP2 were compared in pyramidal cells dendrites in
mammalian brain slices transfected ballistically or by electroporation (Mao
et al., 2008) under conditions that allowed comparison with first generation
sensors (Pologruto et al., 2004). Their performance was not significantly better
than GCaMP, even when localized using membrane and cytoskeletal targeting
chimeras (Mao et al., 2008). GCaMP3, however, showed substantial gains in
sensitivity and discrimination (Tian et al., 2009): overall, the signal-to-noise
ratio was much improved and responses in dendrites to single action potentials
could be reliably detected. Direct comparison with TN-XXL and cameleon D3
showed that, although brighter, the two FRET sensors gave smaller fluorescence
changes and less favorable signal-to-noise ratios. GCaMP3 was also more photo-
stable. After either adenoviral transfection or in utero electroporation, calcium
responses in pyramidal neurones could be observed in awake, behaving mice
(Tian et al., 2009). Parallel electrical recordings showed that detectable calcium
responses were associated with three or more action potentials. Calcium responses
were also readily observed in the glomeruli of Drosophila antennal lobe and in
sensory neurones of C. elegans, altogether a methodological tour de force (Tian
et al., 2009).
IV. Use of Genetically Encoded Calcium Sensors
For single cell applications, wide-field fluorescence imaging, spinning disk, or
confocal microscopy are appropriate methods. Dual excitation laser scanning
confocal imging is achievable (Shimozono et al., 2002). For whole animal applica-
tions, particularly in intact brain or brain slices two photon microscopy is recom-
mended, as it reduces tissue damage and oVers improved imaging within tissue (see
Chapter 9; Fan et al., 1999).
Expression of sensors in cells and tissues, as we have seen, can be achieved by
transfection and transgenesis. One advantage of transgenic approaches is that
expression can be confined to a specific tissue or cell type, an advantage even if it
is excised for imaging. Random expression in a subset of cells can more simply be
achieved by using biolistic transfection of excised tissue.
Ratiometric sensors (in this context the FRET-based sensors, ratiometric peri-
cam and DRIP) oVer the advantage that the quantitative signal is in theory
independent of variations in sensor distribution and concentration within cells
(Silver et al., 1992). This allows reliable calibration of the signal in terms of calcium
concentration (see chapter 1). Nonratiometric sensors (e.g., GCaMP3) are ade-
quate for determining changes in calcium concentration, for example, when mea-
suring overall spatial and temporal patterns of calcium signaling. Even in these
circumstances, caution should be exercised in case the responses are nonlinear,
6. Genetically Encoded Probes for Measurement of Intracellular Calcium 177
especially at low calcium concentration, so that a subset of smaller signals is
overlooked (ReiV et al., 2005).
In general, genetically encoded calcium sensors are not available commercially,
though Invitrogen oVers YC3.60 (http://probes.invitrogen.com/media/pis/
mp36207.pdf). Some can be obtained for noncommercial use from their creators
(http://www.tsienlab.ucsd.edu/ and http://cfds.brain.riken.jp/). Or you can make
your own using the handbook (Miyawaki et al., 2003a, 2005).
V. Conclusions
Genetically encoded calcium sensors have proved valuable in the measurement
of calcium concentration in cellular organelles, for the most part in single cells
in vitro. Their success as sensors in tissues in vitro and in vivo is qualified. They have
also proved valuable in imaging the pattern of calcium signals within tissues,
particularly in the poikilotherms, C. elegans, Drosophila, and zebrafish. In home-
otherms, the record is largely disappointing, even when tissue is excised and
monitored at room temperature (Pologruto et al., 2004). Striking exceptions are
the use of GCaMP2 to image calcium-signaling patterns in mouse heart (Tallini
et al., 2006) and pyramidal neurones (Tian et al., 2009). For the most part, sensors
are still not capable of sensing individual calcium events in single cells when these
cells are part of tissue, though single cell responses can be monitored in disaggre-
gated cells (KotlikoV, 2007). Some branches of the calcium sensor evolutionary
tree continue to evolve rapidly and the steady progress in optimizing sensor
parameters leads to the certain hope that these drawbacks will eventually be
overcome by further genetic engineering.
Acknowledgments
I thank Jill McKenna for helping with this chapter. Our work is supported by grants from the
Wellcome Trust.
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CHAPTER 7
METHODS IN CELL BIOL
Patch Clamp Methods for StudyingCalcium Channels
David L. Armstrong, Christian Erxleben, and Jody A. WhiteMembrane Signaling GroupLaboratory of NeurobiologyNational Institute of Environmental Health SciencesNIH, Durham, North Carolina, USA
A
OGY,
bstract
VOL. 99 0091183 DOI: 10.1016/S0091
-679X-679X
I. I
ntroduction II. R ationale III. M ethodsA.
Assembling the Patch Clamp Rig B. Making Pipettes C. Making Seals D. Making RecordingsIV. M
aterials V. D iscussion VI. S ummaryR
eferencesAbstract
The patch clamp technique, which was introduced by Neher and Sakmann and
their colleagues in 1981, has allowed electrophysiologists to record ion channel
activity from most mammalian cell types. When well-established precautions are
taken to minimize electrical and mechanical fluctuations, current transients as small
as 0.5 pA and as brief as 0.5 ms can be measured reliably in cell-attached patches of
plasmamembranewith apolishedglass pipettewhen it forms a giga-ohmsealwith the
membrane. Inmany cases, this is suYcient to watch individual channel proteins open
and close repeatedly in real time on metabolically intact cells. No other technique
currently provides a more precise or detailed view of the function and regulation of
calcium channel gating. If antibiotics are added to the pipette to permeabilize the
/10 $35.00(10)99007-9
184 David L. Armstrong et al.
membrane underneath to small monovalent cations, thereby allowing the entire cell
to be voltage-clampedwithout disrupting its contents, the integrated activity of all the
calcium channels in the surface membrane can be measured.
I. Introduction
Calcium ions trigger many fundamental cellular processes by binding to pro-
teins, usually with dissociation constants around 1 mM calcium. However, unlike
other second messengers such as cyclic nucleotides, calcium is neither created nor
destroyed by biological processes. Therefore, regulating calcium-dependent pro-
cesses requires moving calcium ions into and out of cellular compartments. Trans-
port proteins that hydrolyze ATP move calcium ions from lower to higher
concentrations. They maintain resting cytoplasmic calcium at 100–200 nM by
pumping calcium out of the cytosol into the endoplasmic reticulum, the mitochon-
dria, or out across the plasma membrane. By contrast, ion channel proteins, when
the channels are open, allow calcium ions to diVuse rapidly down their electro-
chemical gradient back into the cytosol. Many channels are permeable to calcium
because they pass all cations, including calcium, up to a certain size, but most of
the current is carried by more prevalent ions such as sodium or potassium. In
contrast, some ‘‘calcium-selective’’ channels pass calcium almost exclusively, even
in the presence of other cations.
There are currently three classes of calcium-selective channel proteins (Table I):
three families of voltage-gated calcium channels in the plasma membrane (CaV1-3);
two families of calcium-selective channels in the endoplasmic reticulum membrane;
Table ICalcium-selective channels
Voltage-activated calcium channels in the surface membrane
L type CaV1.1 CACNA1S Skeletal muscle
CaV1.2 CACNA1C Cardiovascular muscle
CaV1.3 CACNA1D Endocrine cells, neurons
CaV1.4 CACNA1F Retina
P/Q type CaV2.1 CACNA1A Nervous system
N type CaV2.2 CACNA1B Nervous system
R type CaV2.3 CACNA1E Nervous system
T type CaV3.1 CACNA1G Brain, heart
CaV3.2 CACNA1H Brain, endocrine cells, heart
CaV3.3 CACNA1I Brain
Calcium release-activated channel (CRAC) in the surface membrane
STIM-gated ORAI1-3?
Ligand-gated calcium channel in the endoplasmic reticulum
IP3-gated ITPR1-3
Ryanodine-gated RYR1-3
7. Patch Clamp Methods 185
and calcium release-activated channels (CRAC) that mediates store-operated calci-
um entry (SOCE) across the plasma membrane (Hille, 2001; Hogan et al., 2010). All
of these proteins provide channels that allow calcium to diVuse into the cytosol whenthe channel opens. Each open channel protein has a unique unitary conductance for
calcium, ranging from approximately 0.1 to several hundred picosiemens (pS), but
the proteins spontaneously cycle between open and closed conformations on a time
scale of milliseconds. It is the rates of these transitions rather than the conductances
which are regulated by physiological events to control the amplitude of calcium
fluxes. Such unitary currents are often diYcult tomeasure because they are small and
individual openings last less than a millisecond. However, the total amount of
current crossing the entire surface membrane of a cell at any time is the product of
the number of channels (N), the fraction of time they spend in the open state (Po),
their unitary conductance (g), and the electrochemical driving force (DV) measured
as the diVerence between the voltage across the membrane (Vm) and the ion’s Nernst
potential.
Even small currents produce physiologically significant increases in intracellular
calcium. For example, a current of only 0.1 pA (pA¼10�12 A), which corresponds
to 0.1 pC of charge per second, or �300,000 divalent ions per second, will transfer
300 calcium ions each millisecond the channel is open. Such a current, 0.1 pA
lasting 1 ms, is just below the current technology of detection with the patch clamp
technique. Nevertheless, it would produce a physiologically significant change in
intracellular calcium. The calcium ions cannot diVuse on average much farther
than a micrometer in a millisecond, so the concentration under the membrane will
rise transiently to 0.5 mM, more than double the resting level of calcium. If there
was only one such channel that opened for 1 ms in every square micrometer of
membrane of a spherical cell with a diameter of 10 mm (volume�0.5 pL; area -
�300 mm2), then the resulting 30 pA current would almost double intracellular
calcium concentration throughout the cell. Action potentials that depolarize cells
for tens of milliseconds will have correspondingly larger eVects. Thus, millisecond
diVerences in calcium channel kinetics have profound consequences for cell physi-
ology and human health (Erxleben et al., 2006).
This calculation also illustrates the danger of expressing recombinant channels
in mammalian fibroblasts. Investigators routinely report currents of a few
nanoamperes, which even inexperienced investigators can measure with the
patch clamp technique. However, in the scenario outlined above, a 3-nA current
would represent a 100� higher density of channels and produce 100� larger
increases in calcium, which might lead to cytotoxic reactions. In most cases, such
recordings are made with exogenous calcium buVers in the cytosol, which not only
prevent cytotoxicity but also preclude analysis of physiological regulation of
calcium channels by calcium-dependent signaling. In addition, because most calci-
um channels have a low probability of opening (Po) less than 0.1, the larger current
density reflects at least 1000 channel proteins per square micrometer, or more than
10% of the space available with close packing. At this density, there might not be
room for each channel protein to be associated with its normal penumbra of
186 David L. Armstrong et al.
regulatory proteins, even if their expression was upregulated by the cell to com-
pensate for the increased expression of the channels.
Recording the activity of recombinant channels from cells that are dialyzed with
simple salt solutions using the conventional ‘‘whole-cell’’ configuration of the
patch clamp is also dangerous because in the absence of normal metabolic activity,
many calcium channels have conformations from which it is diYcult to elicit
robust activity with physiological stimuli. Nevertheless, if the plasmid drives the
expression of 100–10,000 times more channels than are normally present, even
channels with very little activity might generate quite large currents. We would
argue, however, that identifying the mechanisms that halve or double the activity
of recombinant channels, which only open for 1 or 2 ms every second, might not be
relevant to their physiological functions. Thus, this chapter focuses on the more
diYcult and consequently less frequently used techniques of single-channel record-
ing from cell-attached patches and the perforated patch technique for voltage
clamping metabolically intact cells.
II. Rationale
Calcium signaling can be investigated at many levels. Although the patch clamp
method is a very quantitative technique for measuring ion fluxes across the plasma
membrane, internal stores of calcium are important sources that the patch clamp
technique cannot access. To study calcium release from internal stores through
calcium-selective channels, one must use fluorescent calcium indicators or recon-
stitute the channels from organelles in bilayers. This chapter provides an overview
of the patch clamp method for measuring voltage-activated calcium currents
across the plasma membrane. We have applied this technique primarily to dis-
sociated mammalian cells in vitro culture, and it has also been adapted to studying
neurons in brain slices (Sakmann and Neher, 1995). Very basically, the technique
involves the use of an operational amplifier circuit to clamp voltage changes
between a wire in the patch pipette and a wire in the bath and the measurement
of how much current it takes to hold the voltage constant. When a giga-ohm
(GO¼109 ohms) seal is formed between the glass patch pipette and the cell
membrane, background current fluctuations can be reduced suYciently to detect
picoampere currents. Gigaohm seals are most eVective for stable, low-noise re-
cording when they are in the 10–100 GO range. However, in our experience, most
investigators routinely settle for seals in the 1–10 GO range. The methods de-
scribed below allow us to routinely obtain seals around 50 GO. Unfortunately,
they are all necessary for success.
When the patch clamp technique was introduced almost 30 years ago (Hamill
et al., 1981), it was the only quantitative method available to obtain reliable
physiological information about calcium signals in mammalian cells and the
proteins that mediate them. Now, however, there are increasingly sophisticated
calcium indicators that can be targeted genetically to specific compartments in
7. Patch Clamp Methods 187
specific cells. In addition, genome cloning, fluorescence confocal microscopy, and
structural studies of membrane proteins have advanced to the point where they are
better suited than the patch clamp technique for identifying which channel proteins
are expressed by specific cell types, how their distribution on and traYcking to and
from the cell surface is regulated, and their three-dimensional molecular structures.
Nevertheless, the patch clamp remains the technique of choice for investigating the
channels’ physiological function and regulation at the molecular level in real time.
In the future, genetic manipulation of channel sequence and expression in model
organisms and mass spectrometry of posttranslational modifications might begin
to supplant even these applications of the patch clamp technique.
Three volumes of Methods in Enzymology have already been devoted to the
patch clamp method, so it would be impossible to provide a suYciently detailed
introduction here that would allow a novice to make recordings without consulting
other sources. Scientists with little training in physics and/or membrane physiology
should start with the ion channel primer by Aidley and Stanfield (1996) and the
technical primers by Molleman (2003) and Ogden (1987). More experienced phy-
siologists could move directly to the compendia by Sakmann and Neher (1995) and
by Rudy and Iverson (1992). More specialized topics are covered in subsequent
volumes edited by Conn (1998, 1999). If you are creating your own rig from
scratch, chapter by Jim Rae and Rick Levis in volume 207 is essential reading.
III. Methods
A. Assembling the Patch Clamp Rig
To localize contributions to noise, it is important to assemble the rig slowly,
piece by piece. There are two general classes of noise to eliminate initially. Sinu-
soidal fluctuations arise from electronic interference by power sources and from
mechanical vibrations. In practice, they are often diYcult to distinguish because
they produce oscillations of similar frequency, but both should be eliminated
completely. Later, after giga-ohm seal formation, any remaining noise should
have much higher frequencies. Some of this noise arises from capacitative coupling
between the glass wall of the pipette and the salt solution surrounding it in the
bath, which can be minimized experimentally. Remaining contributions to high
frequency noise can only be reduced by filtering the signal, which limits how brief
an event of a given amplitude one can detect.
To begin, run the calibration tests that are specified by the manufacturer on the
amplifier while it is sitting on a desk. Then assemble the air table, the faraday cage,
the microscope, and the manipulator. They should share a common ground. The
lamp of the microscope will require a DC power supply and each of the manip-
ulator’s motors must be grounded separately. Surprisingly, despite all their metal
parts, many microscopes are not isopotential and need to be grounded at several
points. With the input of the amplifier’s headstage still open and the output filtered
Table IIChecklist for reducing noise
Electrical sources
Faraday cage grounded
Microscope and (all three) manipulator motors grounded
DC supply for microscope lamp or LED
Check for ground loops
Pipette holder clean and dry
Mechanical sources
Table floating freely
Microscope and manipulator tightly secured to table
Electrical lines and perfusion tubing lashed tightly to microscope base or table
No air drafts blowing on rig
Rubber gasket that holds glass capillary in pipette holder in new condition
188 David L. Armstrong et al.
to 100 Hz, confirm that there are no sinusoidal fluctuations at 10� the highest gain
you plan to use. Table II contains a checklist for major sources of noise.
Finally, test the pipette holder, metal electrodes, and perfusion system by
inserting into the recording chamber a cover slip that has a hemisphere of solidified
sylgard elastomer (184, Dow Corning). Pressing a patch pipette into the sylgard
allows you to obtain seals of up to 100 GO. Silver wires can be chlorided by
dipping them in chlorine bleach for several minutes, then rinsed thoroughly and
air dried. Because silver is toxic to cells, the ground wire is connected to the bath
through an agar salt bridge.
B. Making Pipettes
Commercial programmable pipette pullers are now available, but some experi-
mentation is required inevitably to find the settings that produce pipettes with the
desired overall shape: long and narrow tapers for cell-attached patch recordings to
minimize capacitance with the bath; and short stubby tapers for perforated patch
whole-cell recordings to speed antibiotic diVusion to the tip and minimize access
resistance. Handling the capillaries with wet or greasy hands on the portion that is
heated contributes to variability, but before pulling the pipettes, both ends of the
capillaries should be fire polished gently to increase the longevity of the silver
chloride coating on the wire in the pipette holder. Otherwise, the sharp glass edge
at the back of the pipette scrapes oV a little silver chloride every time you insert a
new pipette. Also make sure the pipette holders and their internal rubber gaskets
are designed to hold capillaries of the same outer diameter as your pipettes, or they
will not fit snugly.
For most low-noise applications, the pipette must be coated with ‘‘Sylgard’’
(Corning 184), an elastomer that is cured with heat, which insulates the pipette
walls from the bath solution. The reported curing time for Sylgard is 10 min at
7. Patch Clamp Methods 189
150 �C; however, a brief exposure (<1 min) to a heating element followed by 10–
20 min at room temperature should be adequate for curing before moving on to
shaping of the tip. The Sylgard is most eVective when it is applied thickly, but
prevents giga-ohm seal formation if it gets on the tip. Our method of applying
Sylgard is illustrated in Fig. 1. We use gravity by mounting the pipette vertically
A
C E
FD
B
Fig. 1 Application of Sylgard. Sylgard (Corning 184), a heat-cured polymer, is applied to patch
pipettes after pulling but before polishing to reduce the capacitance between the glass walls and the bath.
(A) Sylgard is stored in small 1-ml plastic tubes in the freezer until it is ready for use. Room temperature
Sylgard is applied using a 25-gauge hypodermic needle that is bent for ease of application. The needle
hub is aYxed to a plastic tube for ease of handling. (B) The pipettes are placed into a hand-made holder
that holds the pipette upright and allows it to be positioned easily so that the heating element made of
tungsten wire surrounds the tip. (C) The pipette is positioned so that the area from the first narrowing of
the glass to the tip is placed above the heating element. (D) Sylgard is applied in consecutive rings or
‘‘donuts’’ starting at the area where the glass first narrows. This lower ring helps protect against noise if
your bath height changes between perfusions. (E) Between applications of consecutive rings, apply heat
to cure each ring. Remember that heat goes up, so the hottest area near the heating wire is above it. (F)
The final ring should also coat the taper of the pipette and can be placed as close as 50–100 nm from the
tip. Be careful not to get Sylgard on the tip or on the heating element!
190 David L. Armstrong et al.
inside a loop of tungsten wire across the prongs of an outlet plug. We control the
heat transiently with a rheostat and observe the pipette through an inexpensive,
low power, dissecting scope mounted horizontally on a boom arm. The Sylgard is
precured to a viscous consistency at room temperature when it is first made and
then stored in small 1 ml aliquots in the freezer. We apply it with a bent hypoder-
mic needle by taking up a dollop onto the needle, touching the dollop to the
pipette, and slowly winding it on to the glass by turning the pipette from below.
Care must be taken to avoid touching Sylgard to the heating/curing wire, or the
Sylgard vapors will quickly coat the tip. You will know when you get Sylgard too
close to the tip because when you try to polish it, the Sylgard contracting around
the thin glass walls at the tip literally wrinkles the glass.
Everybody has their own favorite glass that they believe forms the tightest seals,
but their relative intrinsic noise can be tested empirically using the Sylgard hemi-
spheres. The lowest noise is produced by quartz glass capillaries (Levis and Rae,
1993), but they require a special laser puller to create the pipettes. On real cells, the
size and shape of the tip also influence success. It is extremely diYcult to make
pipettes that will routinely make >10 GO seals when you cannot clearly see the tip
while you polish it. An extra long working distance (ELWD) objective with at least
40� magnification is essential but they are expensive, so they must be mounted
below the polishing element that is heated to prevent cracking the lens from
repeated heating. Our polishing strategy is illustrated in Fig. 2. We find that it is
also critical to melt a small bead of the pipette glass onto the apex of the heating
element, usually a small loop of platinum wire, presumably because it prevents the
tip from being coated with metal. We find that pipettes with bullet-shaped tips and
initial resistances between 3 and 5 MO make the best seals.
Filling the pipettes also requires some attention to detail. All pipette solutions
should be filtered through 0.2 or 0.45-mm filter disks but do not use filter disks
prepared with wetting agents. To avoid bubbles and washing the dirt inside the
capillaries down into the tip, both of which reduce seal success, one must first fill
the tip separately by immersing it in a small vial of pipette solution and allowing
the first 20–50 mm to fill by capillary action. Then the rest can be backfilled.
Usually bubbles are visible to the naked eye, and they can be removed by gently
flicking the pipette while it is held between the thumb and the forefinger.
C. Making Seals
To reliably get seals over 10 GO, all the precautions in Table III are important. In
addition you have to be fast. It should take only 3–5 min from filling the pipette to
touching down onto the cell, including mounting the pipette in the holder, manip-
ulating it into the chamber, zeroing out the oVset, measuring the resistance, finding
it in the microscope, and manipulating it just above the cell without touching the
bottom or another cell. This takes practice. It also helps to run a little solution
through the chamber before lowering the pipette into the bath to clear any debris
that accumulates on the surface. To minimize debris from collecting on the patch
A
B C
D E
Fig. 2 Fire-polishing pipettes. Fire polishing the tip allows the user to narrow the tip opening to gain
the desired shape and resistance. (A) The pulled pipettes are placed horizontally in a 2D holder mounted
on the stage of an inverted microscope with a long working distance objective �40�. On the opposite
side of the stage, a 3D manipulator is placed with a small platinum wire loop on which a bead of pipette
glass has been melted. (B) The unpolished pipette tip is brought into the same plane of view as the glass
bead, but at opposite sides of the periphery. (C) When current is passed through the wire, the bead gets
hot and expands into the center of the field. The tip of the pipette is transiently manipulated closer to the
bead until the opening at the tip starts to close. (D)When the tip narrows and attains rounded edges, the
pipette is withdrawn and the heat is turned oV. (E) A finished, polished pipette tip.
Table IIIChecklist for obtaining giga-ohm seals
Cells are healthy
CO2 regulation is accurate
No trypsin for 24 h and no transfection reagents in past 12 h
Maintained in salt solution at room temperature for less than 15 min
Pipettes are functioning properly
Pipettes are stored in dust-free box for �4 h after polishing
Pipette solution filtered through �0.45 mm mesh
Tip filled separately by capillary action before back filling; no bubbles
Pipette holder gasket sized to glass and fits snugly (but do not over tighten)
Suction line intact and dry
No mechanical vibrations transmitted to tip (see Table II)
Solutions
Bath solution has more solutes than pipette solution
Neither solution has proteins or ATP
At least one of the solutions has �0.1 mM Ca2þ
7. Patch Clamp Methods 191
192 David L. Armstrong et al.
pipette after it is lowered in the bath, gentle positive pressure is applied on the
suction line which is then pinched off until contact with the cell. In addition, bath
solutions are designed to be 10% hyperosmotic to the pipette solution (usually by
adding glucose), so water will stream out of the pipette tip. Axial approaches are
more eVective than lowering the angled pipette straight down on to the cell. Some
manipulators program a fourth axis or you can mount the manipulator at an angle
so one axis is parallel to the pipette. We prefer to monitor the approach of the
pipette to the cell surface by looking at the pipette current trace in response to a
small voltage step. As the pipette touches the cell, the amplitude of the current
diminishes. When it is reduced to 33–50%, we release the positive pressure on the
pipette, and then, if necessary, we apply additional suction. To make these manip-
ulations without disturbing the tip, the small tube connected to the pipette holder
must be firmly anchored to the headstage and the microscope stage to prevent
vibrations. Most of us prefer to apply the suction directly by mouth because the
suction is controlled more easily and changes more evenly. Small flat cells are
obviously harder to patch than large round ones. Visualization of the cell surface
is improved by interference contrast optics, but this can be implemented on very
simple microscopes for the price of two small pieces of black tape (Axelrod, 1981).
Most people we have trained find the rat pituitaryGH cell lines to be the easiest cells
with which to learn patching. They can be obtained from the ATCC.
D. Making Recordings
1. Cell-Attached Patch Recordings
The two most important things to remember about the cell-attached configura-
tion are that the voltage polarities are reversed relative to traditional whole-cell
recording and that there are two membranes in the current path between your
electrodes. The amplifier sets the voltage between the pipette wire and the ground
electrode in the bath, but, by convention, the cell interior is negative to the outside,
so depolarizing the membrane of a cell-attached patch means making the pipette
more negative. However, the cell membrane also contributes a voltage which is not
clamped in the cell-attached configuration, so to accurately determine the voltage
across the patch, the cell’s membrane potential must be set to zero by bathing the
cell in an equimolar potassium solution of 140–150 mM. Unfortunately, that
means all the voltage-gated channels, including the calcium channels, in the
membrane outside the patch will be activated. Therefore, it is essential to reduce
extracellular calcium to avoid flooding the cell with calcium. Some investigators
use calcium buVers, but the bath is infinite relative to the cell volume and most cells
are rapidly depleted of calcium. A more physiological solution is to reduce extra-
cellular calcium to 0.1 mM, which is still 100�more than resting calcium inside the
cell, but produces negligible currents in the presence of 2–5 mM magnesium.
For cell-attached recordings, the primary goal is increasing the signal and
reducing noise. Increasing the signal is usually achieved by increasing the
7. Patch Clamp Methods 193
concentration of the divalent ion in the pipette solution on the outside of the patch.
Practically, 90 mM calcium is as high as one can achieve without making the
pipette solution hyperosmotic to the bath. However, such large currents also invite
unphysiological, or even toxic, calcium responses in the cytoplasm, so most people
use 90 mM barium. The conductance of voltage-gated channels to barium is often
higher than to calcium, and barium blocks many potassium channels, but high
concentrations of divalent ions alter the surface potential of the membrane and
shift voltage-activation and inactivation curves. If all the precautions that are
described here are taken to reduce noise, it is possible to record unitary currents
with physiological concentrations of calcium (Josephson et al., 2010).
Patch pipettes with initial resistances greater than 10 MO rarely make high
resistance seals, so several square micrometers of membrane are usually drawn
into wider, lower resistance pipettes by the suction. Such large areas of membrane
usually contain many diVerent types of channels, many of which have much larger
unitary conductance than calcium channels. Therefore, sodium, potassium, and
chloride must be replaced with impermeant ions and sometimes, ion channel
blockers must be added too. Cesium does not permeate potassium selective chan-
nels, but it does go through many nonselective cation channels, so N-methyl-d-glucamine (NMDG) is a safer substitute for sodium. Chloride can be replaced with
methanesulfonic acid, although a few millimolar chloride must be left in to allow
reversible current movement between the wire and the solution.
To reduce background noise further, coating the pipette with Sylgard and
obtaining higher resistance seals are the two most practical steps. Lowering the
bath also helps although no perfusion system is perfect, so the bath cannot be too
low. Stable recordings also require a drift-free manipulator or lifting the cell oV the
bottom of the chamber, which is easier when calcium is removed from the bath
solution. If you can lift the cell oV the substrate, the lowest noise recordings can be
obtained by putting a layer of inert oil over the surface (Rae and Levis, 1992), but
then the chamber must be designed with the perfusion inlet and outlet at the
bottom.
2. Perforated Patch Recordings
Many people report that they have tried perforated patch recording but could
not get it to work. In our experience, there are two critical steps that befuddle most
beginners until we show them the ‘‘secrets.’’ The two essential steps to successful
perforated patch recording are eYcient solubilization of the antibiotic and optimal
loading of the pipette (Horn and Marty, 1988). We learned how to sonicate from
Robert Rosenberg, a calcium channel researcher at UNC Chapel Hill for many
years, who also used bilayers. He taught us that cylindrical devices are most
eVective, and they must be filled to the height where the water surface is most
agitated. Then placing a covered, round bottom, cylindrical glass tube with less
than 1 ml of solution into the vortex for less than a minute is suYcient to disperse
the nystatin or amphotericin or gramicidin, but this only lasts for an hour or two.
194 David L. Armstrong et al.
The second secret is empirically determining exactly how far to fill the tip with
antibiotic-free solution. If the antibiotic is too close to the tip, it can permeabilize
the entire cell membrane and prevents giga-ohm seal formation. If it is too far
back, it takes forever for the perforation to proceed. Therefore, each person must
take the time to figure out exactly how long to dip their pipettes in the antibiotic-
free solution by painstakingly examining and measuring how far up the solution
goes for a given count; ‘‘one one thousand, two one thousand, etc.’’ With those
preliminaries, we routinely get access resistances below 20 MO in 5 min and often it
goes down closer to 10 MO.Finally, there is one mistake that even experienced electrophysiologists make
that is much easier to avoid. Most patch clampers almost exclusively use the
‘‘whole cell’’ configuration, in which the membrane underneath the pipette is
disrupted by suction, and the cell is dialyzed with the pipette solution. To achieve
this configuration, they routinely add calcium chelators, such as EGTA, to the
pipette solution. Chelating calcium destabilizes the patch membrane and prevents
cytoplasmic calcium from rising to toxic levels. However, if EGTA is not removed
for perforated patch recordings, suction often leads to cell dialysis with the antibi-
otic, and the current through its channels dwarfs the calcium current.
3. Recordings of Calcium Release-Activated Currents, Icrac
The conductance of individual Icrac channels is too low to measure with the
patch clamp technique although it has been estimated by fluctuation analysis
(Prakriya and Lewis, 2006). While Icrac resulting from overexpression of STIM/
Orai in recombinant systems can be as large as 100 pA/pF at �100 mV, which
translates into 1 nA whole-cell current in an average HEK293 cell, endogenous
Icrac currents are only a few pA/pF or about 10 pA for the average cell. In order to
measure whole-cell currents in the 1–10 pA/pF range, low-noise techniques that
are usually only used for high-resolution single-channel current measurements
need to be employed. Specifically, the whole-cell recording electrodes should be
coated with an elastomer-like Sylgard 184 and, of course, the initial seal resistance
prior to establishing whole-cell configuration should be as high as possible. With
Sylgard-coated and subsequently fire polished electrodes, one should routinely
obtain seal resistances of �50 GO in the cell-attached mode on HEK cells. At
least for HEK293 cells, if calcium is buVered to 100 nM in the pipette, the whole-
cell configuration can then be obtained easily by a single, brief, and gentle suction
pulse.
In classical voltage jump protocols that are used to elicit whole-cell calcium
currents, the eVect of carefully coating the pipette with Sylgard can be readily
observed as a reduction of the fast capacitive transients at the beginning and end
of the step. For Icrac measurements, however, investigators routinely use fast
voltage-ramp protocols (typically �100 mV/s) to measure the quasi steady-state
IV relationship of Icrac. Under those conditions, the contribution of capacitative
current to the total current is less visible since it remains constant during the ramp,
7. Patch Clamp Methods 195
and small changes in the bath level that are almost inevitable during perfusion
changes will change the amplitude of the capacitative current. This can produce a
shift of the instantaneous current–voltage relationship along the current axis,
which can easily be misinterpreted as a shift in the reversal potential of Icrac.
IV. Materials
Although all patch clampers have their favorite vendors, as government-
employed scientists, we cannot reveal our specific preferences here (see
Table IV). If there is someone else at your institution who is patch clamping
successfully, you should consult them first anyway and consider buying what
they have after you try it because they will already be familiar with its operation.
V. Discussion
The patch clamp methods described here are not easy. They have daunting
initial costs in instrumentation, almost $100,000 at current list prices, and a steep
learning curve for both the technical aspects and conceptual foundations of
electrophysiology. Even for experienced investigators, the patch clamp technique
is a laborious method that requires constant trouble shooting. Nevertheless, like
any physical activity, patch clamp performance is always enhanced mysteriously
by a belief in success. Thus, a common refrain around our laboratory is ‘‘If Neher
or Sakmann could do it, then you can do it too.’’
Table IVComponents of a patch clamp rig
Tungsten wire for sylgarding and polishing
Silver wire and pellets
Pipette glass
Pipette puller
Dissecting microscope on swing arm for sylgard coating
Pipette polisher with
Inverted microscope with �40� LWD objective
Coarse manipulator for adjusting heating element
Fluorescent microscope with diVerential interference contrast optics
LED illumination is cheaper, cooler, and electrically quieter
3D micromanipulator with submicron resolution and no drift
Patch clamp amplifier, computer interface, and software
Computer according to the software manufacturer’s specifications
Physiological chamber and perfusion apparatus
Pipette holder and agar bridge ground electrode
Small cylindrical sonicator
196 David L. Armstrong et al.
VI. Summary
The patch clamp technique provides quantitative records of calcium channel
activity in the plasma membrane at the molecular level in real time in situ.
However, studying calcium channel function and regulation under physiologi-
cal conditions in metabolically intact cells requires more demanding approaches
than the conventional whole-cell recording through ruptured patches on dia-
lyzed cells. Whole-cell recordings through antibiotic-perforated patches and
single-channel recordings from cell-attached patches require additional eVortand attention to detail, but are currently unrivaled in their precision and
detailed view of calcium channel function and physiological regulation of
gating.
Acknowledgments
Our studies have been supported by the NIH intramural program at NIEHS through Grant Z01-
ES080043. We also thank Sue Edelstein for drawing Figures 1 & 2.
References
Aidley, D. J., and Stanfield, P. R. (1996). ‘‘Ion Channels: Molecules in Action.’’ 1st edn. Cambridge
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Axelrod, D. (1981). Zero-cost modification of bright field microscopes for imaging phase gradient on
cells: Schlieren optics. Cell Biophys. 3, 167–173.
Conn, P. M. (1998). ‘‘Methods in Enzymology: Ion Channels Part B.’’ Vol. 293. Academic Press,
New York, NY.
Conn, P. M. (1999). ‘‘Methods in Enzymology: Ion Channels Part C.’’ Vol. 294. Academic Press,
New York, NY.
Erxleben, C., Liao, Y., Gentile, S., Chin, D., Gomez-Alegria, C., Mori, Y., Birnbaumer, L., and
Armstrong, D. L. (2006). Cyclosporin and Timothy syndrome increase mode 2 gating of CaV1.2
calcium channels through aberrant phosphorylation of S6 helices. Proc. Natl. Acad. Sci. USA 103,
3932–3937.
Hamill, O. P., Marty, A., Neher, E., Sakmann, B., and Sigworth, F. J. (1981). Improved patch-clamp
techniques for high-resolution current recording from cells and cell-free membrane patches. Pflugers
Arch. 391, 85–100.
Hille,B. (2001). ‘‘IonChannels ofExcitableMembranes.’’ 3rd edn. SinauerAssociates, Inc., Sunderland,MA.
Hogan, P. G., Lewis, R. S., and Rao, A. (2010). Molecular basis of calcium signaling in lymphocytes:
STIM and ORAI. Annu. Rev. Immunol. 28, 491–533.
Horn, R., and Marty, A. (1988). Muscarinic activation of ionic currents measured by a new whole-cell
recording method. J. Gen. Physiol. 92, 145–159.
Josephson, I. R., Guia, A., Lakatta, E. G., Lederer, W. J., and Stern, M. D. (2010). Ca(2þ)-dependent
components of inactivation of unitary cardiac L-type Ca(2þ) channels. J. Physiol. 588, 213–223.
Levis, R. A., and Rae, J. L. (1993). The use of quartz patch pipettes for low noise single channel
recording. Biophys. J. 65, 1666–1677.
Molleman,A. (2003). ‘‘PatchClamping,An IntroductoryGuide toPatchClampElectrophysiology.’’ John
Wiley & Sons, Chichester, England.
Ogden, D. (1987). ‘‘Microelectrode Techniques: The Plymouth Workshop.’’ 2nd edn. The Company of
Biologists Limited, Cambridge, UK.
7. Patch Clamp Methods 197
Prakriya, M., and Lewis, R. S. (2006). Regulation of CRAC channel activity by recruitment of silent
channels to a high open-probability gating mode. J. Gen. Physiol. 128, 373–386.
Rae, J. L., and Levis, R. A. (1992). A method for exceptionally low noise single channel recordings.
Pflugers Arch. 420, 618–620.
Rudy, R., and Iverson, L. E. (1992). ‘‘Methods in Enzymology: Ion Channels.’’ Vol. 207. Academic
Press, New York, NY.
Sakmann, B., andNeher, E. (1995). ‘‘Single-Channel Recording.’’ 2nd edn. PlenumPress, NewYork,NY.
CHAPTER 8
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
Nuclear Patch-Clamp Recording fromInositol 1,4,5-Trisphosphate Receptors
Taufiq Rahman and Colin W. TaylorDepartment of PharmacologyTennis Court Road, University of CambridgeCambridge, United Kingdom
A
OGY,All rig
bstract
VOL. 99 0091hts reserved. 199 DOI: 10.1016/S0091
-679X-679X
I. In
troduction II. N uclear Patch-Clamp Recording III. C hoice of Cells for Analyses of IP3R IV. M ethodsA.
Culture of DT40 Cells B. Isolation of Nuclei C. Solutions for Patch-Clamp Recording D. Patch-Clamp Recording E. Analysis of Single-Channel RecordsV. C
oncluding Remarks R eferencesAbstract
Inositol 1,4,5-trisphosphate receptors (IP3R) are ubiquitous intracellular Ca2þ
channels. They are regulated by IP3 and Ca2þ and can thereby both initiate local
Ca2þ release events and regeneratively propagate Ca2þ signals evoked by receptors
that stimulate IP3 formation. Local signaling by small numbers of IP3R underpins
the utility of IP3-evoked Ca2þ signals as a ubiquitous signaling pathway. The
physiological impact of Ca2þ release by very small numbers of IP3R underscores
the necessity to understand the behavior of IP3R at the single-channel level.
In addition, and in common with analyses of every other ion channel, single-
channel analyses have the potential to define the steps linking IP3 binding to
channel opening. Patch-clamp recording, by resolving the openings and closings
of single channels with exquisite temporal resolution, is the most powerful
/10 $35.00(10)99008-0
200 Taufiq Rahman and Colin W. Taylor
technique for analysis of single-channel events. It has contributed enormously to
the understanding of gating and desensitization/inactivation of numerous ion
channels. However, most IP3R reside within intracellular membranes, where they
are inaccessible to conventional patch-clamp recording methods. Here, we describe
the application of nuclear patch-clamp methods to single-channel analyses of
native and recombinant IP3R.
I. Introduction
Inositol 1,4,5-trisphosphate receptors (IP3R) comprise a family of tetrameric
intracellular channels that mediate the release of Ca2þ from the intracellular stores
of almost all animal cells (Foskett et al., 2007; Taylor et al., 1999). Three genes
encode homologous subunits of vertebrate IP3R, and a single gene encodes inver-
tebrate IP3R. The key structural determinants of IP3R activation, although pres-
ently poorly understood, are likely to be similar for all IP3R. Activation is initiated
by binding of IP3 to a conserved IP3-binding core toward the N-terminal of each
subunit (Bosanac et al., 2002), conformational changes then pass via the
N-terminal suppressor domain (Bosanac et al., 2005) to the pore, which is formed
by transmembrane regions lying toward the C-terminus (Boehning and Joseph,
2000; Foskett et al., 2007; Rossi et al., 2009; Taylor et al., 2004). Most IP3R in most
cells are expressed within membranes of the endoplasmic reticulum (ER). DiVerentIP3R subtypes may, however, diVer in their subcellular distributions (Taylor et al.,
1999) and in their modulation by various additional signals and associated pro-
teins (Betzenhauser et al., 2008b; Choe and Ehrlich, 2006; Mackrill et al., 1997;
Patterson et al., 2004; Wojcikiewicz and Luo, 1998). Resolving the roles of diVer-ent IP3R subtypes in the genesis of the complex Ca2þ signals that regulate cellular
activity is an important issue (Futatsugi et al., 2005; Miyakawa et al., 1999;
Sugawara et al., 1997; Wang et al., 2001).
Opening of the intrinsic pore of all IP3R requires binding of IP3 and Ca2þ
(Adkins and Taylor, 1999; Marchant and Taylor, 1997). IP3R can, therefore,
both initiate the Ca2þ signals evoked by receptors that stimulate IP3 formation
and then regeneratively propagate them by Ca2þ-induced Ca2þ release. This dual
regulation of IP3R allows a hierarchical recruitment of Ca2þ release events as the
stimulus intensity increases (Bootman et al., 1997; Marchant and Parker, 2001).
Single IP3R respond first, then several IP3R within a cluster open together to give
larger local events (‘‘puVs’’), and as puVs become more frequent, they ignite
regenerative Ca2þ waves (Bootman and Berridge, 1995; Marchant et al., 1999).
This hierarchy of events allows Ca2þ to function as a local or global messenger, a
feature that underlies its versatility (Berridge et al., 2000). A key point for the
present discussion is that local events involving very few IP3R underlie the Ca2þ
signals that regulate cellular activity. By contrast, for most ion channels, it is the
collective behavior of large numbers of channels, the macroscopic current, that
determines the physiological response, a change in membrane potential, or
8. Patch-Clamp Recording of IP3 Receptors 201
transcellular ion flux, for example. The distinction highlights the particular impor-
tance of single-channel recording in the analysis of IP3R. As with all such analyses,
they provide the highest resolution insight into the opening and closing of single
channels and can thereby reveal details of gating mechanisms (Colquhoun, 2007;
Sivilotti, 2010). But for IP3R and other Ca2þ channels too, openings of individual
channels are the physiologically significant behavior.
The patch-clamp technique, developed originally by Neher and Sakmann (1976)
with subsequent improvements (Hamill et al., 1981), is the most powerful means of
studying the behavior of ion channels in their native environment. It involves
recording currents passing through an electrically isolated, small area (‘‘patch’’)
of biological membrane in response to an applied voltage or ionic gradient
(Fig. 1A and B). Isolation of the patch is achieved by pressing a polished glass
pipette tip of �1 mm diameter (containing electrolyte solution) against the cell-
surface and applying gentle suction to form a very high-resistance ‘‘giga-Ohm’’
(GO) seal (Hamill et al., 1981). The tight seal is crucial because it isolates the patch
both electrically and physically, so reducing background noise and allowing single-
channel events to be resolved (Hamill et al., 1981). Because of the electrical
isolation and low resistance of the patch-pipette relative to the membrane, a
patch can be voltage-clamped by simply applying a potential to the pipette.
These patch-clamp recordings allow the openings and closings of individual chan-
nels to be resolved with submillisecond temporal resolution under optimal condi-
tions (Fig. 1C). The amplitudes of these tiny currents and their dependence on
applied potential and ion concentrations allow the ion selectivity and rates of ion
permeation to be determined. Lurking within the pattern of stochastic openings
and closings is the information from which the sequence of events that leads to
channel gating and desensitization/inactivation can be reconstructed. Comprehen-
sive descriptions of the patch-clamp technique are available from the original
articles (Hamill et al., 1981; Neher and Sakmann, 1976) and subsequent reviews
(Ogden, 1994; Sakmann and Neher, 1995). However, most IP3R are expressed in
membranes of the ER, where they are inaccessible to conventional patch-clamp
techniques. Alternative approaches are therefore needed.
II. Nuclear Patch-Clamp Recording
It is impracticable, despite one heroic success recording from IP3R within the
ER of an intact cell (Jonas et al., 1997), to use patch-clamp techniques routinely to
record the behavior of single channels within the membranes of intracellular
organelles in situ. A more promising approach for single-channel recordings
in situ is the ‘‘optical patch-clamp,’’ where high-resolution optical microscopy in
combination with fluorescent Ca2þ indicators is used to measure the Ca2þ signals
evoked by opening of single or small clusters of IP3R (Demuro and Parker, 2007;
Smith and Parker, 2009). Presently, however, these optical methods can be used
only to measure fluxes through Ca2þ channels, and they lack the temporal
C
O1
O2
20pA
500 msK+
Cell-attached
Inside-out Outside-out
Whole-cell
A
Referenceelectrode
Patch pipette
Ionchannel
Bath
Patch-clampamplifier
B
CIP3
Fig. 1 Conventional patch-clamp recording. (A) A polished glass pipette forms a tight seal against a
biological membrane isolating an area across which the tiny currents passing through small numbers of
open channels can be recorded. (B) DiVerent configurations of conventional patch-clamp recording.
Beginning with a cell-attached patch, an inside-out excised patch can be produced by pulling the patch-
pipette away from the cell, while suction or a strong brief voltage-pulse ruptures the underlying
membrane to give the whole-cell configuration. An outside-out patch can then be produced by pulling
the patch-pipette away from the whole-cell configuration. (C) Typical whole-cell recordings of IP3R3
expressed in the plasma membrane of DT40-KO cells expressing rat IP3R3. PS included IP3 (10 mM),
ATP (5 mM), and a free [Ca2þ] of �200 nM; Kþ was the charge carrier and the holding potential was
�100 mV. C, O1, and O2 show the closed state and the openings of 1 and 2 IP3R, respectively.
202 Taufiq Rahman and Colin W. Taylor
8. Patch-Clamp Recording of IP3 Receptors 203
resolution of conventional patch-clamp recording. Measuring the electrical activity
of intracellular channels, therefore, presently relies upon redirecting channels to the
plasma membrane, where they then become accessible to conventional patch-clamp
techniques (Xu et al., 2007); reconstituting the channel into an artificial membrane;
or isolating organelles that express the channel and adapting the patch-clamp
technique to record from these membranes. The latter has been used, for example,
to resolve the behavior of the mitochondrial Ca2þ uniporter from mitochondria
stripped of their outer membrane (‘‘mitoplasts’’) (Kirichok et al., 2004) and for
single-channel recordings of the endolysosomal protein, TRPML1, from artificially
enlarged lysosomes (Dong et al., 2008). All three methods have been used to record
single-channel behavior of IP3R.
We observed that DT40 cells express very small numbers of functional
IP3R within the plasma membrane (Dellis et al., 2006) and because DT40 cells
lacking native IP3R are available (Section III), conventional whole-cell patch-
clamp recording has been used by us (Dellis et al., 2008) and others
(Betzenhauser et al., 2008b, 2009a) to examine the behavior of recombinant and
mutant IP3R. Typical recordings from IP3R in the plasma membrane of DT40 cells
are shown in Fig. 1C. A limitation of this approach is that excised patch-clamp
recording, where the ‘‘intracellular’’ composition can be precisely controlled, is
impracticable (because plasma membrane IP3R are too scarce), and with whole-
cell recording, it is diYcult to define reliably the exact concentration of IP3 bathing
the IP3R (Dellis et al., 2006). Detailed descriptions of the methods used for whole-
cell recording of IP3R expressed in the plasma membrane of DT40 cells have been
published (Dellis et al., 2006; Taylor et al., 2009b). We prefer nuclear patch-
clamping (see below) to whole-cell recording because the nuclear envelope is
continuous with the ER (Fig. 2A), wherein reside most IP3R, and it is practicable
to work with excised patches that provide better control of media bathing both
sides of the membrane.
The first electrical recordings from IP3R were made by incorporating native or
purified IP3R into artificial lipid bilayers (Bezprozvanny et al., 1991; Ehrlich and
Watras, 1988; Maeda et al., 1991; Mayrleitner et al., 1991). As with all reconsti-
tuted systems, the lipid composition of the bilayer and the steps involved in
isolating, purifying, and reconstituting IP3R into the bilayer may aVect normal
function of the channel, not least its regulation by accessory proteins (Boehning
et al., 2001a; Foskett et al., 2007; Patterson et al., 2004). Anecdotally, and it
equates with our experience, it seems to be more diYcult to obtain bilayer record-
ings from IP3R than from its close relatives, the ryanodine receptors (Williams,
1995).
Many of the problems with bilayer recording are resolved by using nuclear patch-
clamp recording (Fig. 2). This technique was first introduced in the early 1990s
(Matzke et al., 1990; Mazzanti et al., 1990, 2001; Tabares et al., 1991) and subse-
quently, applied by the laboratories of Clapham (Stehno-Bittel et al., 1995) and
Foskett (Mak and Foskett, 1994) to record single-channel events from native
IP3R in nuclei from Xenopus oocytes. It has, subsequently, been successfully applied
Cytoplasm-out
A B
C
On-nucleus
ONMINM
Lumen-out
Cell
Nucleus
ONMINM
Nucleoplasm
10mm
C
10 pA
500 ms K+
IP3E
Nucleus
Debris
D
K+
Cytosol
Lumen
PS
+40 mV
BS
Fig. 2 Nuclear patch-clamp recording of IP3R. (A) The nuclear envelope comprises an inner (INM)
and outer (ONM) membrane surrounding a luminal space that is continuous with the lumen of the ER.
The continuity of the ONMwith the ERmembrane allows some ER proteins to invade the ONM, where
they become accessible to nuclear patch-clamp recording. (B) Phase-contrast image of a DT40 nuclear
preparation showing nuclei, one of which has debris attached, and an intact cell. (C) Three recording
configurations are used for nuclear patch-clamp recording. The on-nucleus, lumen-out, and cytosol-out
excised patch configurations are analogous to the cell-attached, inside-out, and outside-out configura-
tions of conventional patch-clamp recording (Fig. 1B). (D) An excised lumen-out nuclear patch
illustrating the convention used to report membrane potential. (E) Typical recording from a single
IP3R3 recorded in the lumen-out configuration from the nucleus of a DT40-KO cell stably expressing
rat IP3R3. PS included IP3 (10 mM), ATP (5 mM), and a free [Ca2þ] of �200 nM; Kþ was the charge
carrier and the holding potential was þ40 mV. C denotes the closed state.
204 Taufiq Rahman and Colin W. Taylor
to analyses of diVerent IP3R subtypes expressed in diVerent cells, including COS-7
cells (Boehning et al., 2001a), insect Sf9 cells (Ionescu et al., 2006), smooth muscle
(Kusnier et al., 2006), DT40 cells (Betzenhauser et al., 2009b; Dellis et al., 2006;
8. Patch-Clamp Recording of IP3 Receptors 205
Rahman et al., 2009), human B lymphoblasts (Cheung et al., 2008), cerebellar
Purkinje cells (Marchenko et al., 2005), and embryonic cortical neurons and fibro-
blasts (Cheung et al., 2008).
The utility of nuclear patch-clamp recording derives from the fact that the outer
nuclear envelope is continuous with the ER membrane (Dingwall and Laskey,
1992) (Fig. 2A). Channels that are normally expressed within ER membranes can,
therefore, pass into the outer nuclear envelope, allowing their activity to be
recorded from patches of nuclear membrane in a near-physiological setting
(Fig. 2). Abundant nuclear pore complexes, each with a large central conduit
linking cytoplasm and nucleoplasm (Mazzanti et al., 2001), might have been
expected to compromise formation of the tight seals required for patch-clamp
recording or at least pollute recordings from conventional channels with lesser
conductances. In practice nuclear pores appear not to cause problems. It seems
unlikely, though it remains possible, that this results from patching onto ER
overlying the nuclear envelope, rather than the envelope itself. It is perhaps more
likely that patches that include nuclear pores are rejected because they fail to form
giga-Ohm seals, or the high-Kþ medium used for nuclear patch-clamp recording
favors closure of nuclear pores (Bustamante and Varanda, 1998).
III. Choice of Cells for Analyses of IP3R
Almost all animal cells express IP3R, most express more than one of the three
vertebrate gene products, and substantial alternative splicing and posttranslational
modifications add further to the diversity of subunits from which IP3R are assem-
bled (Foskett et al., 2007). Assembly of these subunits into homo- and heterote-
trameric structures increases the diversity of functional IP3R enormously (Joseph
et al., 1995, 2000; Wojcikiewicz and He, 1995). Despite this complexity, there have
been many valuable studies of the single-channel behavior of native IP3R in, for
example, Xenopus oocytes, nuclei from oocytes, insect Sf9 cells, and cerebellar
Purkinje neurons, and of native IP3R reconstituted into lipid bilayers (Section II).
But the limitations of such studies are obvious when it to comes to exploring the
structural basis of IP3R activation. This demands a more homogenous population
of IP3R with a defined structure and ideally expressed in a native membrane.
At present, only one expression system provides the ‘‘null background’’ that allows
these demanding criteria to be satisfied: DT40 cells (Fig. 3).
DT40 cells originate from an avian leukosis virus-transformed bursal B cell
(Baba et al., 1985). The uniquely valuable feature of these cells is the unusually
high frequency with which they integrate targeted DNA constructs into their
genome (Buerstedde and Takeda, 1991). This feature, together with the shorter
introns of avian genes, allows targeted disruption of specific genes and has ensured
widespread use of DT40 cells for ‘‘gene-knockouts.’’ In a monumental eVort,Kurosaki and his colleagues used targeted gene disruption to inactivate both
copies of all three IP3R genes in DT40 cells and thereby to generate the first cell
DT40-KO
A B
−10 −8 −6 −4
0
50
100DT40-R3
Log {[IP3], M}
Ca2+
rel
ease
(%
)
- CDT40-KO
- C
DT40-R3
10 pA
1 sK+
IP3
Fig. 3 DT40-KO cells provide a null background for expression of functional IP3R. (A) IP3-evoked
Ca2þ release from permeabilized DT40 cells assessed using a luminal Ca2þ indicator (Tovey et al., 2006).
Permeabilized DT40-KO cells stably expressing rat IP3R3 (DT40-R3) release Ca2þ when stimulated
with IP3, whereas DT40-KO cells are unresponsive. (B) Currents recorded from lumen-out patches from
DT40-KO and DT40-R3 cells. PS included IP3 (10 mM), ATP (5 mM), and a free [Ca2þ] of �200 nM;
Kþ was the charge carrier and the holding potential was þ40 mV. C denotes the closed state.
206 Taufiq Rahman and Colin W. Taylor
line lacking functional IP3R (Sugawara et al., 1997). These DT40-KO cells, which
Kurosaki (RIKEN, Japan) has made widely available, provide the only null
background for functional expression of IP3R (Fig. 3). They have been extensively
used by many groups to express each of the three IP3R subtypes and define their
functional properties, and to explore the role of IP3R in many higher order
processes (e.g., Joseph and Hajnoczky, 2007; Miyakawa et al., 1999). A recent
review provides further details of the use of DT40 cells for analyses of Ca2þ
signaling pathways (Taylor et al., 2009b). Here, we describe our use of DT40
cells expressing mammalian IP3R for nuclear patch-clamp recording.
IV. Methods
A. Culture of DT40 Cells
Wild-type DT40 cells (Cell Bank number RCB1464) and DT40-KO cells
(RCB1467) are available from Riken Bioresource Center Cell Bank, Japan
(http://www.brc.riken.jp/lab/cell/). Cells are grown in RPMI 1640 medium (Invi-
trogen) supplemented with 10% fetal bovine serum (FBS, Sigma), 2 mM l-gluta-mine, 1% chicken serum (Sigma), and 50 mM b-mercaptoethanol (Invitrogen) in a
humidified atmosphere containing 5% CO2, ideally at 39–41 �C (matching the
increased body temperature of birds). It is, however, acceptable and more conve-
nient when incubators are shared with mammalian cells to culture DT40 cells at
37 �C without loss of viability. The only obvious eVect is a slowing of growth rate;
the doubling time has been reported to increase from about 10 h at 39–41 �C to
18 h at 37 �C (Mak et al., 2006). The chicken serum must be heat-inactivated
8. Patch-Clamp Recording of IP3 Receptors 207
(56 �C for 30 min). We avoid antibiotics, but addition of penicillin (10,000 units/
ml) and streptomycin (10 mg/ml) to culture media is optional (Winding and
Berchtold, 2001). The cells grow in suspension in flasks, Petri dishes, or multiwell
plates (Greiner Bio-one). Cells are passaged by 20-fold dilution every 2–3 days
when they reach a density of about 2�106 cells/ml. Avoid growing cells beyond a
density of 2.5�106 cells/ml. Cells (2�106ml�1) in either culture medium or FBS
supplemented with 10% dimethylsulfoxide (DMSO, Sigma) can be frozen and then
stored in liquid nitrogen following standard procedures. We routinely culture cells
for 30–35 passages, before thawing a new frozen stock. For the latter, 1 ml of cells
is dispensed into 20 ml of medium. We find it unnecessary to remove residual
DMSO at this stage. Cells are then passaged after 24 h.
DT40 cells are not easy to transfect with IP3R expression constructs, we there-
fore use cell lines stably expressing rat IP3R1 (GenBank accession number of
GQ233032.1), mouse IP3R2 (AB182290), and rat IP3R3 (GQ233031.1). Details
of the methods and sources of the original clones are provided in previous pub-
lications (Dellis et al., 2006; Rahman et al., 2009; Rossi et al., 2009; Tovey et al.,
2010). Briefly, DT40-KO cells are transfected by nucleofection with linearized
constructs of pcDNA3.2-IP3R using solution T and program B23 (Amaxa) using
5 mg DNA/106 cells. G418 (Invitrogen, 2 mg/ml) is used for selection. Expression
of IP3R in each cell line is quantified by immunoblotting using custom-made anti-
peptide antisera (Cardy et al., 1997; Dellis et al., 2006; Rossi et al., 2009; Tovey
et al., 2010) and, where needed, by3
H-IP3 binding. Functional expression of
IP3R in each DT40 cell line is verified by comparison with DT40-KO cells using
a luminal Ca2þ indicator and a high-throughput assay for IP3-evoked Ca2þ release
(Laude et al., 2005; Tovey et al., 2006) (Fig. 3). Only cell lines shown to express
functional IP3R are used for nuclear patch-clamp recording.
B. Isolation of Nuclei
Several methods have been described for isolation of nuclei; most rely on a
combination of osmotic and mechanical lysis of cells (Boehning et al., 2001a;
Bustamante, 1994; Franco-Obregon et al., 2000; Marchenko et al., 2005). Our
protocol is adapted from that of Boehning et al. (2001a). DT40 cells expressing a
recombinant IP3R (DT40-IP3R cells, 1.5–2�106 cells/ml) are centrifuged (500�g
for 2 min at 4 �C), washed once with ice-cold phosphate-buVered saline (PBS), and
then once with cold nuclear isolation medium (NIM). PBS has the following
composition: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2PO4, KH2PO4, pH 7.4,
with NaOH. NIM comprises: 250 mM sucrose, 150 mM KCl, 3 mM b-mercap-
toethanol, 10 mM Tris–HCl, 1 mM phenylmethanesulfonyl fluoride (PMSF,
Sigma), pH 7.5. Cell pellets are resuspended in NIM supplemented with complete
protease inhibitor cocktail (Roche, 1 mini-tablet/20 ml) and stored on ice for up to
4–5 h. For isolation of nuclei, 1 ml of the cell suspension is homogenized with 3–4
strokes of a Dounce homogenizer (Wheaton Industries, Inc.), which lyses about
5–10% of cells, assessed by staining with Trypan Blue (0.001%). This crude lysate
208 Taufiq Rahman and Colin W. Taylor
containing some isolated nuclei is stored in NIM on ice and used within 1 h for
patch-clamp experiments. Because the activity of nuclear IP3R has been reported
to decrease after �40 min at 20 �C (Boehning et al., 2001a), we routinely prepare
fresh nuclei at hourly intervals. DT40 cells are not much larger than their nuclei
(Fig. 2B). The inexperienced eye may therefore find it diYcult to distinguish nuclei
from cells. But nuclei rarely have the smooth surface of intact cells and they often
only partially protrude from broken cells, where the relatively clean exposed
surface allows formation of a giga-Ohm seal. The yield of nuclei can be substan-
tially increased to 50–60% using methods that require incubation in hypo-osmolar
media (Franco-Obregon et al., 2000), but we rarely detect active IP3R after such
isolation procedures. A nuclear isolation kit (Sigma Nuclei EZ Prep) also provides
nuclei in high yield (�85%), but we rarely succeed in forming giga-Ohm seals with
these nuclei. In practice, the low yield of nuclei with our protocol is not a limitation
for nuclear patch-clamp recording.
IP3R have also been reported to be expressed within the inner nuclear membrane
(Humbert et al., 1996; Marchenko et al., 2005). The citrate treatment used to
remove the outer nuclear membrane and so allow patch-clamp recording of
IP3R within the inner membrane (Marchenko et al., 2005) appears not, at least
in our experience, to be readily applicable to DT40 cell nuclei.
C. Solutions for Patch-Clamp Recording
For most recordings, we use Kþ as the charge-carrier. This eliminates the
complexity of having Ca2þ passing through the IP3R regulate its activity, and it
provides larger single-channel currents than with bivalent cations (Rahman et al.,
2009). The bath solution (BS), which bathes the luminal surface of the nuclear
envelope, typically contains 140 mM KCl, 10 mM HEPES, 100 mM 1,2-bis(2-
aminophenoxy)ethane-N,N,N0,N0-tetraacetic acid (BAPTA, tetra potassium salt,
Calbiochem), and a free [Ca2þ] of �200 nM (total CaCl2, 51 mM) adjusted to pH
7.1 with KOH. The usual pipette solution (PS), which bathes the cytosolic surface
of the membrane, contains 140 mM KCl, 10 mM HEPES, 500 mM BAPTA,
Na2ATP (0.5 mM), IP3 (American Radiolabeled Chemicals, Inc.), and a free
[Ca2þ] of �200 nM (total CaCl2, �254 mM) adjusted to pH 7.1 with KOH.
IP3, Ca2þ, and ATP are the three ligands of IP3R whose concentrations must be
adjusted to obtain optimal IP3R activity in patch-clamp recording (Foskett et al.,
2007). The concentration of IP3 in PS can be varied between experiments, depend-
ing on the aim of the analysis; 10 mMwill usually be suYcient to saturate responses
to IP3 (Foskett et al., 2007; Rahman et al., 2009). The potentiating eVects of ATP
diVer between IP3R subtypes with higher concentrations required optimally to
activate IP3R3 (Betzenhauser et al., 2008a; Miyakawa et al., 1999). The pH of PS
must be readjusted after addition of ATP, and its eVects on free [Ca2þ] also need to
be considered. Finally, because Mg2þ aVects the conductance of IP3R (Mak and
Foskett, 1998; Rahman and Taylor, 2009), it is advisable to use ATP of the highest
8. Patch-Clamp Recording of IP3 Receptors 209
purity. Freshly prepared dilutions of ATP and IP3 (from frozen stocks) are added
to PS as required.
EVective buVering of the free [Ca2þ], which might reasonably be varied between
nanomolar and several micromolar, requires buVers with appropriate aYnities for
Ca2þ (Patton et al., 2004). For free [Ca2þ] less than �1 mM, BAPTA
(KDCa�192 nM at pH 7.4) is preferable to EGTA because it has faster Ca2þ-
binding kinetics and lesser pH-dependence. Where the free [Ca2þ] of PS is
1–100 mM, we use 5,50-dibromo BAPTA (KDCa�1.83 mM, Fluka), EGTA
(KDCa�67 nM), and/or N-(2-hydroxyethyl)ethylenediamine-N,N0,N0-triacetic
acid (HEDTA, KDCa�2.2 mM, Sigma), alone or in appropriate combinations
(Bers et al., 1994). We initially estimate the amount of CaCl2 required to achieve
the desired free [Ca2þ] using WinMaxC software (http://www.stanford.edu/�-
cpatton/maxc.html) and then measure the free [Ca2þ] of the final media (supple-
mented with ATP, IP3, etc.) directly using either a fluorescent Ca2þ indicator
(Fluo-3, KDCa¼325 nM, Invitrogen) or a Ca2þ-sensitive electrode (Mettler
Toledo Ingold, Fisher Scientific) for higher free [Ca2þ] (Dellis et al., 2006;
Rahman et al., 2009).
The osmolarities of all solutions are adjusted to �290–310 mOsm kg�1 using
glucose and mannitol, and verified using a vapor pressure osmometer (Wescor,
Inc.). This is more important for recordings in the on-nucleus configuration than
for recordings from excised patches (Fig. 2C). PS is prepared to be slightly (�10%)
hypo-osmolar to BS to aid formation of giga-Ohm seals (Hamill et al., 1981). All
recording solutions are filtered using detergent-free 0.2-mm filters (AcrodiscÒ
syringe filters, Pall Corporation) (Ogden, 1994). Fresh recording solutions (with-
out added IP3 or ATP) are prepared monthly and stored at 4 �C.The presence within the nuclear envelope of other large-conductance cation and
Cl� channels (Franco-Obregon et al., 2000; Marchenko et al., 2005; Mazzanti
et al., 2001; Tabares et al., 1991) might potentially contaminate recordings of
nuclear IP3R. In practice, this appears not to be a significant problem. If such
problems should arise, they can be mitigated by replacing KCl in BS and PS with
cesium methanesulfonate (CsCH3SO3): Csþ permeates IP3R but not Kþ channels
(Tovey et al., 2010), while most anion channels are impermeable to CH3SO3�.
D. Patch-Clamp Recording
The equipment required for nuclear patch-clamp recording is the same as that
used for conventional patch-clamp recording (Fig. 1A). The basic rig includes an
amplifier, headstage, electrode holder, micromanipulator, AgCl bath electrode,
data acquisition system (i.e., analog-to-digital converter, computer, and software),
inverted microscope, air table, and a Faraday cage. In addition, a pipette puller
and fire-polisher or microforge are required to fabricate electrodes. Optional
extras include systems for exchange of solutions, an oscilloscope, and a low-pass
8-pole Bessel filter; the latter extends the filtering range down to 0.1 Hz from the
1 to 100 kHz provided by the inbuilt filter. Comprehensive descriptions of the
210 Taufiq Rahman and Colin W. Taylor
equipment used for patch-clamp recording are presented in relevant chapters of
Sakmann and Neher (1995). Among the many steps taken to minimize electrical
noise, the following are particularly important: appropriate grounding of equip-
ment, use of thick-walled glass capillaries, filling pipettes with PS to the minimal
level required to contact the recording electrode, and minimizing immersion of the
pipette in BS (Rae and Levis, 1992).
Pipettes are pulled from filamented, thick-walled borosilicate glass capillaries
(GC150-10F, Clark Electromedical Instruments) using a Flaming/Brown P-87
horizontal micropipette puller (Sutter Instruments), and then fire-polished to a
tip diameter of �1 mm using a microforge (MF-830, Narishige). It is advisable to
melt a small bead of glass onto the wire of the forge to prevent platinum vapor
from reaching the pipette tip. With monovalent cations as charge carriers, the
single-channel conductance (g) of IP3R is large enough (�360 pS, Section IV.E) to
achieve good signal-to-noise ratios without hydrophobic coating of the patch-
pipette (Penner, 1995). But when g of IP3R is reduced, with pore mutants or with
Ca2þ or Ba2þ as charge carriers, for example, it may be necessary to coat pipette
tips with SylgardTM
(Dellis et al., 2006, 2008). When filled with PS, the pipette
resistance typically remains within the range of 15–20 MO. Pipettes are best
prepared a few hours before experiments. Unused pipettes can, however, be stored
in an air-tight container and used later, but it is advisable to repolish them lightly
before use to remove any impurities accumulated during storage.
Petri dishes are precoated with poly-l-ornithine or poly-l-lysine (0.01%, Sigma)
for 1–2 h, then rinsed twice with deionized water and air-dried. The nuclear
preparation (15 ml) is added to a Petri dish containing BS (1.5 ml) and the cells/
nuclei are allowed to adhere. The dish is then mounted on the stage of an inverted
microscope (Zeiss Axiovert 100) coupled to an assembly of headstage (CV 203 BU,
Molecular Devices) and micromanipulator (PCS-1000, Burleigh Instruments).
Recordings are made at room temperature (�20 �C) in the on-nucleus or excised
configuration (Fig. 2C). The latter is preferable because it allows control of the
medium on both sides of the membrane and eVective control of the voltage acrossthe patch.
A nucleus largely free from debris is first identified (Fig. 2B) and the patch-
pipette is positioned, using the micromanipulator, with its tip just above the
nucleus. A slight positive pressure is applied to the inside of the patch-pipette
before dipping it into the BS to avoid dirt accumulating at the pipette tip and to
prevent backflow of BS into the PS (Hamill et al., 1981). After dipping the pipette
into BS, the pipette capacitance is compensated using the specific oVset on the
amplifier and the pipette resistance (typically �10–15 MO) is noted. As the pipette
tip approaches the nucleus, the positive pressure is relieved. Taking care not to
puncture the nuclear membrane, the pipette is lowered until it contacts the mem-
brane, which should increase the pipette resistance by at least 2 MO. A giga-Ohm
seal (�5 GO) usually forms within a few seconds of applying slight negative
pressure, by suction, to the inside of the pipette; this is usually controlled by an
attached 50-ml syringe or by mouth. Seal formation can sometimes be facilitated
8. Patch-Clamp Recording of IP3 Receptors 211
by applying a holding potential of about �40 mV once a high-resistance contact is
established (Ogden, 1994), and occasionally giga-Ohm seals form spontaneously.
Seals >5 GO can be routinely obtained with mild suction providing the nucleus is
immobile and free of debris.
To formanexcised patch, our preferred recording configuration, the patch is pulled
from the nucleus after forming the giga-Ohm seal (Fig. 2C). To prevent formation of
closedvesicles at the tipof thepatch-pipette, excisedpatchesarebriefly (1–2 s) exposed
to air and then reimmersed in BS (Hamill et al., 1981). Prewritten protocols are then
used to record currents through the excised patch at diVerent holding potentials. Thebath electrode is grounded (i.e., 0 mV) and for convenience, the potential across a
nuclear patch (whether attached or excised) is defined as the pipette potential minus
thebathpotential. That is,with symmetricalmedia, a positiveholdingpotentialwould
favor movement of cations from PS (the cytosolic surface) into BS (the luminal
surface) producing an outward current and an upward deflection on the channel
record (Fig. 2D and E) (Franco-Obregon et al., 2000; Mak and Foskett, 1994;
Rahman et al., 2009). For determination of current–voltage (I–V) relationships, and
thereby the single-channel conductance (g) of the channel (Section IV.E), the voltage
across the excised patch can be stepped from�60 toþ60 mV in increments of 20 mV
fromaholding potential of 0 mV.Applyingmore extreme voltages aVects the stabilityof thenuclear patch.For all other experiments, includingkinetic analyses, currents are
typically recorded atþ40 mV for between 1 and 10 min.
Currents are amplified with an Axopatch 200B amplifier in its voltage-clamp
mode, filtered at 1 kHz with a low-pass 4-pole Bessel filter (built into the amplifier),
and digitized at 10 kHz with a Digidata 1322A interface using the PC-based
acquisition software package pClamp 9.2 (Molecular Devices) (Colquhoun,
1994). This filtering, while it inevitably causes some loss of information, has the
eVect of rejecting signals (background noise) that are too brief to reflect the gating
of IP3R. If the filtering frequency is set too low, it will reject events that do reflect
gating of channels, and if set too high, background noise will obscure the openings.
The optimal filtering frequency is, therefore, a compromise that depends upon the
noise and time-course of the channel events; it needs to be optimized empirically.
The sampling rate must, of course, exceed the filter frequency if further valuable
information is not to be lost as the signals are digitized. In practice, digitization
should be 10–20 times faster than the cutoV or ‘‘corner’’ frequency of the filter
(Colquhoun, 1994). Most nuclear patch-clamp studies of IP3R have used 1-kHz
filtering (Dellis et al., 2006; Ionescu et al., 2006; Mak and Foskett, 1997; Rahman
et al., 2009). For presentation, traces can be further filtered oZine using a Gaussian
filter (built within ClampFit).
For the determination of relative permeabilities to cations, asymmetric record-
ing solutions are used. For example, the normal BS can be replaced by a Ba2þ-richBS (50 mM BaCl2, 30 mM KCl, 10 mM HEPES, adjusted to pH 7.1 with KOH),
while the PS remains unchanged (Boehning et al., 2001a; Dellis et al., 2006). The
liquid junction potential (LJP) under this asymmetric condition can be predicted
(5.2 mV at 20 �C), using the ‘‘junction potential calculator’’ (JPCalc, within
212 Taufiq Rahman and Colin W. Taylor
pClamp 9.2), which uses the generalized Henderson equation (Barry, 1994). The
calculated LJP is then subtracted from the observed reversal potential (Erev)
obtained from the current–voltage (I–V) plot. We return in Section IV.E, after
considering analysis of the raw traces, to describe how Erev allows the relative
permeability of IP3R to diVerent cations to be calculated.
In both the on-nucleus and excised patch configurations described above, the
cytoplasmic surface of the IP3R lies within the patch-pipette (Fig. 2C); it is, therefore,
diYcult to change the IP3 concentration once the giga-Ohm seal has formed. It is
possible, though diYcult, to perfuse a patch-pipette and thereby to vary the composi-
tion of the ‘‘cytosolic’’ medium while recording channel activity (Hering et al., 1987;
Maathuis et al., 1997), but this technique has not yet been applied to IP3R. Other
options include the cytoplasm-out configuration of nuclear patch-clamp recording
(Fig. 2C), which has been successfully applied to analyses of IP3R in Sf9 cells (Mak
et al., 2007). Alternatively, flash-photolysis of caged-IP3 within the patch-pipette in
either the on-nucleus or excised nuclear patch configuration can be used rapidly to
increase the IP3 concentration bathing the cytosolic surface of the IP3R once the
recording is underway (Rahman et al., 2009). For these flash-photolysis experiments,
pipettes are prepared from thin-walled, nonfilamented borosilicate glass capillaries
(Harvard Instruments) and PS includes d-myo-inositol 1,4,5-trisphosphate, (4,5)-1-
(2-nitrophenyl) ethyl ester (caged-IP3, �100 mM, Calbiochem). After recording for
30–60 s, IP3 can thenbe released intoPSbyphotolysis of caged-IP3usinga singlehigh-
intensity flash (1 ms) from a Xe-flash lamp (XF-10, Hi-Tech Scientific; 240 J with the
capacitor charged to 385 V) passed through a filter (300–350 nM) (Walker et al.,
1987). A problemwith this approach is the diYculty in assessing the concentration of
IP3 to which the IP3R are exposed after flash-photolysis of caged-IP3.
E. Analysis of Single-Channel Records
Two sorts of information can be extracted from single-channel records: the
properties of the open channel (its ability to conduct diVerent ions); and the
sequence of stable states through which the channel passes as it moves between
closed, open, and desensitized conditions. Here, we provide only a brief introduc-
tory summary of the methods used to extract this information from the openings
and closings of channels resolved by patch-clamp recording. The reader interested
in more rigorous and detailed descriptions is advised to begin with two excellent
books (Ogden, 1994; Sakmann and Neher, 1995).
Several software packages are available for analysis of electrophysiological
records. These include ClampFit, which includes the pClamp suite (Molecular
Devices), and DC-soft, which includes SCAN, EKDIST, and HJCFIT (http://
www.ucl.ac.uk/Pharmacology/dcpr95.html); QuB (www.qub.buValo.edu), Pulse/
Patchmaster (HEKA Elecktronik), Tac (Bruxton Inc.), and the Strathclyde Electro-
physiology Software (http://spider.science.strath.ac.uk/sipbs/software_ses.htm).
We use ClampFit and QuB for analyzing records (Dellis et al., 2006; Rahman
et al., 2009).
0.0
0.0
Cou
nt/to
tal
D
A
B
8. Patch-Clamp Recording of IP3 Receptors 213
Drifting baseline in current traces is first checked and corrected manually using
ClampFit. Current-amplitude histograms of recordings with no obvious sub-con-
ductance states are measured with a half-amplitude threshold-crossing criterion
using ClampFit (Sachs et al., 1982; Colquhoun, 1994). Only events lasting longer
than twice the filter rise time (tr¼0.3321/fc, where fc is the cutoV frequency of the
filter) can reach their full amplitude and so be reliably measured in a threshold-
crossing-based idealization procedure (Colquhoun, 1994) (Fig. 4A and B).
In recordings with 1-kHz filtering, the predicted filter rise time (assuming the filter
behaves as a Gaussian filter) is 332 ms. Events lasting<1 ms are, therefore, omitted
from the amplitude histograms. The peaks of the binned current amplitude histo-
grams are fitted in ClampFit by sums of the appropriate number of Gaussian
Log (duration, ms)
0
3
6to= 10 ms
Cou
nt/to
tal
−1 0 1 2
Log (duration, ms)
1 0 1 20
0.02
0.04
0.06 tc1= 1.14 ms(88%)
tc2= 92 ms(12%)
C
0 5 100
500
1000
Num
ber
of e
vent
s
Amplitude (pA)
C
O
C
O
10 pA
100 ms
Fig. 4 Analysis of the behavior of single IP3R from nuclear patch-clamping recording. (A) Fragment
of a raw record from an excised lumen-out nuclear patch with a single functional IP3R3. The recording
conditions were identical to those shown in Fig. 3B. (B) The idealized record of the same trace produced
as described in the text (Section IV.E). This idealized record is used for all subsequent analyses. (C) All-
points current amplitude histogram, showing two peaks, one at 0 pA (the closed state) and a second at
�10 pA (the single open state of one IP3R). (D) Distribution of the open (top) and closed (bottom) life-
times shown as Sigworth–Sine plots (Sigworth and Sine, 1987). The plots suggest a single open state with
to of 10 ms, and two closed states with tc of 1 and 92 ms.
214 Taufiq Rahman and Colin W. Taylor
probability density functions (pdfs) (Fig. 4C). Mean current amplitudes are plotted
against the corresponding applied potentials to create I–V curves (see Fig. 5).
The unitary conductance (g) and reversal potentials (Erev) are derived by
linear least-square regression analysis using statistical package such as Prism 5
(GraphPad Software, Inc.) or Origin Pro 7.5 (OriginLab Corporation).
Because g is a fundamental property of any ion channel, reflecting interactions
between permeating ions and the residues that form the pore and lead to it, I–V
relationships are often used as ‘‘fingerprints’’ to help identify channels. With Kþ as
charge carrier, the I–V relationships of mammalian IP3R in excised nuclear patches
from DT40-KO cells are linear across a range of applied potentials (�60 to
þ60 mV) (Fig. 5). From the slopes of these I–V plots, we have consistently
observed two populations of IP3R with unitary Kþ conductances (gK) of either�120 or �200 pS (Dellis et al., 2006; Rahman and Taylor, 2009; Rahman et al.,
2009) (Fig. 5B). Neither current was detected in the nuclear envelope of DT40-KO
cells or from DT40-KO cells stably expressing IP3R in the absence of IP3, or with
IP3 in the presence of a competitive antagonist. These values of gK are lower than
reported (�320–360 pS) for IP3R in the nuclear envelope of mammalian cells
(Foskett et al., 2007), but there is wide variation in published values (from �9 to
�480 pS) (Cheung et al., 2010; Rahman and Taylor, 2009). The reason for these
disparities is unresolved, but it may reflect variable amounts of free Mg2þ in PS
causing a reduction in gK (Mak and Foskett, 1998; Rahman and Taylor, 2009).
It is, however, clear from analyses of I–V relationships that all IP3R have a large gfor monovalent cations and lesser g for bivalent cations (Dellis et al., 2006; Foskett
et al., 2007) (Fig. 5B).
C
500 ms
20 pA
C
C
C
C
−80 −40 40 80
−15
−10
−5
5
10
15
V (mV)
I (pA)
BA
+40 mV
+60 mV
0 mV
−40 mV
−60 mV
Fig. 5 Current–voltage relationship for nuclear IP3R. (A) Currents were recorded from lumen-out
patches excised from the nucleus of DT40-KO cells stably expressing IP3R3. PS included IP3 (10 mM),
ATP (5 mM), and a free [Ca2þ] of �200 nM. Kþ was the charge-carrier and the holding potential was
varied between þ60 and �60 mV as shown. C denotes the closed state. (B) From the slope of the
current–voltage (I–V) relationship, g was 208 pS. Results are means�SEM, n¼4.
8. Patch-Clamp Recording of IP3 Receptors 215
Mutations within the putative pore region of IP3R that change g provide direct
evidence that the residues within the P-loop linking the last pair of transmembrane
domains are likely to contribute to the ion-permeation pathway (Boehning et al.,
2001b; Schug et al., 2008). Future work along similar lines is likely to define more
precisely the structural determinants of ion permeation. In addition, our demon-
stration that similar point mutations aVected g of the IP3-activated currents
detected in the plasma membrane of DT40 cells expressing mutant
IP3R provided definitive evidence that the currents were carried directly by IP3R,
rather than by another plasma membrane channel with which IP3R in the ER
might have associated (Dellis et al., 2006).
Resolving the unitary current events associated with opening of individual
IP3R also allows functional IP3R to be counted. The number of active IP3R in a
patch can be estimated from the maximal number of simultaneous openings to the
unitary current level (Horn, 1991) (Figs. 1C and 4C). The likelihood of several
channels opening simultaneously depends upon their Po and the number of chan-
nels (N). We would, for example, need to wait much longer, on average, for six
IP3R with low Po to open simultaneously than for the simultaneous opening of two
IP3R with high Po. We can be confident (p<0.01) that we have detected the entire
complement of active IP3R within a patch, when the recording period is longer
than 5(sNþ1) (Ionescu et al., 2006), where
sN ¼ toN Poð ÞN
" #exp
NtDto
� �ð1Þ
and sN is the mean interval between successive simultaneous openings of all N
IP3R; tD the minimum duration of an open event detectable after filtering
(200 ms in our experiments); and to is the mean channel open time. Confidently,
estimating the number of active IP3R within a patch is important, not the least
because there has been a suggestion that increasing concentrations of IP3 cause
increases in both Po (making it easier to detect simultaneous openings) and the
number of active IP3R (Ionescu et al., 2006). This interesting and unprecedented
behavior, which we fail to see (Rahman et al., 2009), has been invoked to explain
the unusual pattern of quantal Ca2þ release observed for IP3R (Taylor, 1992).
By varying the concentrations of cations on either side of the membrane
(Section IV.D and Fig. 6A), the relative permeability (PBa/PK) can be calculated
using a modified version of the Goldman–Hodgkin–Katz (GHK) equation
(Bezprozvanny and Ehrlich, 1994; Fatt and Ginsborg, 1958):
Erev ¼ RT
2Fln4PBa Ba2þ
� �o
PK Kþ½ ið2Þ
where PBa/PK is the relative permeability to Ba2þ and Kþ, [Kþ]i the [Kþ] in PS,
[Ba2þ]o the [Ba2þ] in BS, Erev the reversal potential (corrected for the LJP, see
Section IV.D), R the universal gas constant, F the Faraday constant, and T is the
C
C
5 pA
500 ms+40 mV
+60 mV
C−60 mV
A B
−80 −40 40 80
−5
5
V (mV)
I (pA)
Fig. 6 Determining the cation-selectivityof IP3R fromnuclear patch-clamprecording. (A)Currentswere
recorded in the same way as described in Fig. 5A, but with the usual PS changed to include Ba2þ (50 mM)
rather thanKþ. The currents recorded at diVerent holding potentials are shown. C denotes the closed state.
(B) I–V relationship showing a reversal potential (Erev) of �23.8�1.4 mV after correction for the liquid
junction potential (Section IV.D). From themodifiedGHKequation, this suggests that the permeability to
Ba2þ relative to Kþ (PBa/PK) is 4.7. The unitary conductance (g) from the slope of the plot is 45�4 pS.
216 Taufiq Rahman and Colin W. Taylor
absolute temperature (K). These analyses have established that IP3R (Dellis et al.,
2006; Foskett et al., 2007), like ryanodine receptors (Williams, 2002), are far less
selective (PBa/PK�7) than Ca2þ channels in the plasma membrane (Fig. 6). The
distinction is important because channels within the plasma membrane must be
able to discriminate between the many ions with an electrochemical gradient across
the membrane, whereas Ca2þ is probably the only cation with an appreciable
gradient across the ER membrane (Somlyo et al., 1977).
In addition to revealing the properties of the open pore, single-channel analyses
can also shed light on the steps that lead to its opening. The kinetic analyses of single-
channel records described here require that channel behavior has attained a steady-
state. This is most easily assessed from a stability plot of single-channel open
probability (Po) versus time (Colquhoun, 1994; Weiss and Magleby, 1989). Only
records or parts thereof with an overall steady-state Po should be used for kinetic
analysis. Files with stable baselines are exported asQuB-supported file formats (.ldt).
In QuB, the files are further examined and sections of data with spurious noise are
excluded using the preprocessing module (‘‘Pre’’). Current traces are then idealized
into noise-free, open, and closed transitions using the segmental k-means (SKM)
algorithm in the QuB suite. This uses a hidden Markov model (HMM) to decide
whether each excursion in the record should be classified as an open or closed state
based upon its amplitude (Qin, 2004) (Fig. 4B). The output at this stage is a categori-
zation of every transition into a switch between current amplitudes: a single closed
current amplitude (baseline noise) and one or several amplitudes of the open channel
(s).Where several evenly spaced current amplitudes are detected, it can be diYcult to
resolve whether they arise from openings of several channels or switches between
8. Patch-Clamp Recording of IP3 Receptors 217
equally spaced sub-conductance states of a single channel (Rahman and Taylor,
2009). For IP3R, sub-conductance states are rare (Rahman and Taylor, 2009), allow-
ing the simplest possible scheme, a switch between a single closed (C) and open (O)
state (C$O) with arbitrarily chosen rate constants (e.g., 100 s�1), to be used for the
initial idealization (Qin, 2004). Beginning with this simple scheme does not compro-
mise lateranalyses thatmight revealmorecomplex relationshipsbetween several open
and closed states. Direct comparison of raw traces with their idealized versions is
essential at this stage to confirm the fidelity of the idealization procedure.
Hitherto, the analysis, has considered only the amplitudes of the currents, the
next step considers the durations of these events in records from single channels.
This provides the opportunity to resolve diVerent open and closed states and
possible relationships between them, leading to plausible gating schemes. The
distribution of lifetimes of a single state of a channel is described by a single
exponential (Colquhoun, 1994). The analysis attempts iteratively to establish, for
each potential gating scheme (beginning with the simplest, C$O), the number of
exponential functions required to describe the closed and open lifetimes derived
from the idealization procedure. A maximum interval likelihood method (MIL) is
used to fit the lifetimes with pdfs (Qin et al., 1996, 1997, 2000). During this fitting
process, a dead-time of 200 ms (twice the sampling interval) is retrospectively
imposed for the correction of missed events (Sivilotti, 2010).
Dwell-time histograms are generated and displayed with logarithmic abscissa
and square root ordinate (Fig. 4D) (Sigworth and Sine, 1987) and fitted by a
mixture of exponential pdfs, defined in the function f(t) as
fðtÞ ¼Xni¼1
aiti
exp �t=tið Þ ð3Þ
where ai is the fractional area occupied by the ith component in the distribution, such
that the areas corresponding to all components sum to unity, and ti is the time
constant for the ith component. The mean life-time (t) is given by the following
equation:
t ¼Xni¼1
aitið Þ ð4Þ
The Sigworth–Sine transformation (Fig. 4D) allows a single plot clearly to display
dwell-times spanning several orders of magnitude. Individual exponential compo-
nents of the distribution can be directly identified from the peaks of the distribution.
After iterative exploration of alternative gating schemes, the log likelihood ratio
(Colquhoun, 1994) is used to identify the scheme that best fits the data. The chosen
scheme is then used to reidealize the raw data to provide the final gating para-
meters (mean life-times and Po). Although these are the methods we have used to
address the gating of IP3R (Rahman et al., 2009), more sophisticated approaches
exploit the additional information that lurks in the correlations that exist between
transitions (McManus et al., 1985).
218 Taufiq Rahman and Colin W. Taylor
Analyses like these identify the numbers of stable open and closed states and
plausible relationships between them. They lead thereby to models of the steps
through which the IP3R passes between its inactive and open states. Such analyses
have so far been rather limited for IP3R, but they clearly suggest the existence of a
single open state and several closed states (Ionescu et al., 2007; Rahman et al., 2009)
(Fig. 4D).
Extending the analysis to patches, in which we detected several IP3R, allowed us
to demonstrate that IP3 causes IP3R to form small clusters of �4–5 channels
within which to is reduced from �10 to �5 ms (Rahman et al., 2009). These
observations lead us to suggest that IP3 contribute to the evolution of elementary
Ca2þ signals by both regulating IP3R activity and by assembling IP3R into clusters,
within which regulation of IP3R by Ca2þ and IP3 is retuned (Rahman and Taylor,
2009; Rahman et al., 2009; Taylor et al., 2009a).
For most channels, including IP3R, single-channel open probability (Po) (rather
than g or the number of active channels) is the behavior that changes as the
stimulus intensity varies. Increasing IP3 or Ca2þ increases Po of IP3R because
both ligands shorten the duration of the closed times, without aVecting to; hence,the probability of finding the channel open (Po) is increased (Foskett et al., 2007;
Rahman et al., 2009). Po is calculated from the fitted amplitude histograms of the
current traces (typically lasting �1 min for IP3R) (Ding and Sachs, 1999):
Po ¼ Ao
Ao þ Ac
ð5Þ
where Ao and Ac are the areas under the curves corresponding to the open and
closed states in the current amplitude histogram, respectively.
When IP3R activity is low, it becomes very diYcult to know how many channels
are contributing because it is unlikely that all will open simultaneously. Under
these conditions, the overall activity is better expressed as NPo which is defined as
(Ching et al., 1999; Rahman et al., 2009):
NPo ¼PNn¼1
ntnð ÞT
ð6Þ
where tn is the total time for which n IP3R are simultaneously open and T is the
duration of the recording.
V. Concluding Remarks
Patch-clamp recording of IP3R expressed within the nuclear envelope allows
single-channel analyses of these otherwise inaccessible intracellular Ca2þ channels
(Figs. 1 and 2). DT40-KO cells provide a null background (Fig. 3) for expression of
recombinant and mutant IP3R allowing functional analysis of IP3R with defined
8. Patch-Clamp Recording of IP3 Receptors 219
composition (Taylor et al., 2009b; Tovey et al., 2006). Nuclear patch-clamp
recording of DT40 cells heterologously expressing mammalian IP3R, therefore,
allows single-channel recording with its exquisite temporal resolution to be com-
bined with opportunities to manipulate systematically the structure of the
expressed IP3R. The stability of these patch-clamp recordings in a native mem-
brane and the opportunity to apply them in various configurations (Fig. 2) aVordvaluable opportunities to examine the behavior of small numbers of IP3R directly
(Rahman et al., 2009) and as a means to address the mechanisms underlying
IP3R activation (Rossi et al., 2009).
In the short period during which nuclear patch-clamp analyses have been
applied to IP3R, they have succeeded in confirming that IP3R are large conduc-
tance, relatively nonselective cation channels, and revealed the durations of the
channel openings and closing (Dellis et al., 2006; Foskett et al., 2007; Rahman
et al., 2009) (Figs. 4–6). Together, these insights allow estimates of the likely Ca2þ
fluxes through individual IP3R for comparison with optical measurements of the
elementary Ca2þ signals evoked by IP3 in situ (Shuai et al., 2007, 2008). Combining
site-directed mutagenesis with nuclear patch-clamp recording has provided direct
evidence that the pore of IP3R is formed by residues within the ‘‘P-loop’’ linking
the final pair of transmembrane domains of each IP3R subunit (Boehning et al.,
2001b; Dellis et al., 2006, 2008; Schug et al., 2008). The eVects of a novel family of
synthetic partial agonists on normal and mutant IP3R analyzed by nuclear patch-
clamp recording have shed light on the first stages of IP3R activation by showing
that the initial conformation changes evoked by IP3 binding to the IP3-binding
core pass onward toward the pore entirely via the N-terminal suppressor domain
(Rossi et al., 2009). Similar analyses have revealed the means, whereby ATP
(Betzenhauser et al., 2008b, 2009b), cyclic AMP-dependent protein kinase
(Betzenhauser et al., 2009a), cyclic AMP (Tovey et al., 2010), and various accesso-
ry proteins (Cheung et al., 2008, 2010; Li et al., 2007) modulate IP3R behavior.
Future application of the nuclear patch-clamp technique to IP3R is certain to add
further to our understanding of the stochastic behavior of single and clustered
IP3R and to resolving the structural basis of IP3R activation.
Acknowledgments
This work was supported by grants from the Wellcome Trust, and the Biotechnology and Biological
Sciences Research Council (UK). T. R. is a Drapers’ Company Research Fellow at Pembroke College,
Cambridge.
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CHAPTER 9
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
Confocal and Multiphoton Imaging ofIntracellular Ca2þ
Godfrey Smith,* Martyn Reynolds,† Francis Burton,* andOle Johan Kemi**School of Life SciencesUniversity of GlasgowUnited Kingdom
†Cairn Research LimitedFaversham, KentUnited Kingdom
A
OGY, VOll rights rA
Les
bstract
. 99 0091-67erved. 225 DOI: 10.1016/S0091-67
I.
W hy Study Ca2þ Signaling with Confocal and Multiphoton Microscopy II. C onfocal Microscopy III. L imitations in Speed of Confocal Imaging IV. L aser Scanning Confocal Microscopy V. T otal Internal Reflection Fluorescence Microscopy VI. F orster Resonance Energy Transfer Microscopy VII. P arallel Scanning Confocal Systems VIII. S pinning Disk Confocal Microscopy IX. P rogrammable Matrix Microscopy X. A dvantages and Disadvantages of Confocal Microscopy XI. M ultiphoton Excitation Laser Scanning Microscopy XII. C a2þ Indicators for Use in Confocal and Multiphoton Microscopy XIII. U se of Dyes for Single-Photon Confocal Microscopy XIV. U se of Dyes for 2P Excitation Microscopy XV. Is It Worth Converting the Intracellular Fluorescence Signal to [Ca2þ]? XVI. C alibration of Single Wavelength Dyes XVII. E stimation of Fmax Values X VIII. E stimation of Fmin or the Dynamic Range of the Dye XIX. C onsequence of Errors in Estimation of Intrinsic and Dye Fluorescence XX. M ultimodal and Multiple Fluorophore Confocal and Multiphoton MicroscopyR
eferences9X/10 $35.009X(10)99009-2
226 Godfrey Smith et al.
Abstract
This chapter compares the imaging capabilities of a range of systems including
multiphoton microscopy in regard to measurements of intracellular Ca2þ within
living cells. In particular, the excitation spectra of popular fluorescent Ca2þ
indicators are shown during 1P and 2P excitation. The strengths and limitations
of the current indicators are discussed along with error analysis which highlights
the value of matching the Ca2þ aYnity of the dye to a particular aspect of Ca2þ
signaling. Finally, the combined emission spectra of Ca2þ and voltage sensitive
dyes are compared to allow the choice of the optimum combination to allow
simultaneous intracellular Ca2þ and membrane voltage measurement.
I. Why Study Ca2þ Signaling with Confocal and MultiphotonMicroscopy
Ca2þ is a ubiquitous intracellular messenger that controls a large number of
cellular processes, such as gene transcription, excitation, contraction, apoptosis,
cellular respiration, and the activity levels of many cell-signaling messenger cas-
cades. Inside the cell, Ca2þmay, under various conditions, sequester into the sarco/
endoplasmic reticulum, mitochondria, and the nucleus, or exist in the cytosol
either in its free form or as bound to buVers. Typically, a large Ca2þ concentration
gradient is maintained across the plasmamembrane of the cell. Because of diVerentCa2þ channels, pumps, and exchangers on the membranes of the cell or organelles,
Ca2þ fluxes may be created at multiple locations in the cell. Therefore, Ca2þ
concentration and signal may be specific with respect to both location and time.
Moreover, Ca2þ may concentrate in distinct cytoplasmic regions because of tight
physical loci not enclosed by membranes, for example, the dyadic area between the
transverse tubule and the sarco/endoplasmic reticulum of muscle cells. Given the
large number of Ca2þ channels feeding it with Ca2þ from both the extracellular
space (transverse tubule) and the sarco/endoplasmic reticulum, such that the dyad
may transiently have very diVerent localized Ca2þ concentrations compared to the
rest of the cytosol which may be only nanometers away. Thus, Ca2þ localizes in the
cytosol as well as within organellar compartments, and these Ca2þ signals may last
for very short timeframes (ns) or for substantially longer periods of time (min).
Ca2þ signaling per se is not within the remit of this chapter and will not be covered
in any detail, but the interested reader is referred to other sources, for example,
Bootman et al. (2001).
Nonetheless, for the purpose of this chapter, it is important to acknowledge that
the average Ca2þ concentration in any given cell usually ranges 0.01–1 mM, but
that the Ca2þ almost never exists uniformly across the cell, and that local Ca2þ
events may occur with very fast time courses. This therefore requires Ca2þ imaging
of live specimens with high spatial and temporal resolution. Thus, one would
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 227
ideally want to distinguish Ca2þ in distinct areas that may be within a nanometer
distance from each other, and to record localized Ca2þ events that may last only a
millisecond. Although such requirements tax any given microscopy system, confo-
cal and multiphoton microscopy systems oVer a range of imaging capabilities that
fulfill these criteria.
II. Confocal Microscopy
Optical sectioning by confocal microscopy adds several benefits to Ca2þ imaging.
Since its early development, confocal microscopy has fundamentally transformed
optical imaging to now provide a valuable addition that allows unprecedented
imaging ofminute optical sections within live specimens in close to real-time speeds.
In terms of Ca2þ imaging, this has opened up new fields of study, given that Ca2þ
signaling in many, if not all, biological systems is compartmentalized within small
sections of the cell and occurs often at very high velocities. Confocal microscopy
allows the study of these events within discrete depths of cell or tissue by blocking
light originating outside the plane of focus. This is achieved by the addition of
confocal apertures in front of the illumination source and in the image plane
directly in front of the signal detection system (see Fig. 1); usually, a photomulti-
plier tube (PMT) that rejects out-of-focus light originating from fluorescence
outwith the area of interest (the focal plane), and only allowing in-focus light
through to the PMT (Webb, 1999) (Fig. 1). This is in contrast to regular epifluor-
escence microscopy, in which the majority of the fluorescence is out-of-focus light
that generally reduces the contrast of the in-focus light, and also dramatically
compromises in-focus detail. This occurs since the emitted fluorescence cannot
be discriminated along the Z-axis (top to bottom), and also less along the X- and
Y-axes (although this has also to do with excitation light sources; see later) in
conventional epifluorescence microscopy (Lichtman and Conchello, 2005)
(See also later). Thus, although confocal microscopy also excites the specimen
along the entire Z-axis in line with conventional epifluorescence microscopy, only
in-focus light is allowed to pass the pinhole of the confocal aperture to enter the
signal detector.
Importantly, confocal imaging may be performed on live specimens residing
under physiologic conditions and that are electrically, chemically, mechanically,
and otherwise active and healthy. Specimens may also be electrically and mechani-
cally stimulated and superfused by any given solutions that would not interfere
with the confocal imaging.
The diVerence between regular epifluorescence and confocal microscopy light
capture abilities can be illustrated by the following examples. Considering that the
depth of focus of a high numerical aperture (NA>1.3) objective is restricted to
�0.3 mm,whereas the depth of a fluorescent cell may be�5–25 mm, it becomes clear
that the depth of focus will only constitute�1–5% of the full depth of the cell. Since
epifluorescence microscopy captures light along the entire Z-axis, 95–99% of the
Lightdetector
Dichroicbeamsplitter
Focal plane
Objective
Confocal aperturewith pinhole
Laser
Fig. 1 Overview of the optical pathway of a confocal microscope. The blue path illustrates the
excitation light, whereas the green path illustrates the emitted fluorescence light. Note that the confocal
aperture with the pinhole in front of the light detector (usually, a photomultiplier tube (PMT)) blocks
out-of-focus light.
228 Godfrey Smith et al.
cell volume will contribute to unwanted out-of-focus background signal, or noise
(Lichtman and Conchello, 2005), and this cannot be distinguished from in-focus
fluorescence. Thus,most of the cell will be out of focus, andwith it, the vastmajority
of the signal will come from out-of-focus areas. In contrast, setting the pinhole of
the confocal aperture to 1 airy unit to achieve true confocality will provide an
optical section or Z-resolution of 0.5–1 mm with the same high NA objective as
described above. This will only allow a minimum of out-of-focus light to reach the
signal detector, without any other interference to the optical pathway or any
secondary digital processing of the signal at the time of recording apart from the
scanning and building of the image itself, which otherwise would have further
compromised the scanning speed.
The full 3D XYZ-resolution, or the ability to discern two points from each
other, will, however, be diVraction-limited as determined by the point spread
function (PSF) set by the optical performance of the microscope. With high NA
objectives (>1.2), this is typically �0.3�0.3�0.6 mm (Cox and Sheppard, 2004).
Although Z-resolution is dramatically diVerent between conventional epifluores-
cence and confocal microscopes, the 2D spatial resolution in the XY-field is
not, though factors, such as excitation wavelength, objectives and the optical
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 229
pathway, and diVerent media and surfaces, will also aVect this. However, several
factors associated with the confocal principle allow for improving also XY-
resolution, as compared to widefield epifluorescence microscopy. First, spatial
resolution may be improved by further reducing the pinhole diameter in the
confocal aperture to a size smaller than the width of the central disk of the airy
unit pattern, though this also dramatically reduces light transmission. This
principle works because the pinhole is aligned with the center of the airy unit
pattern of the illuminating beam, which means that any emission originating
from any fluorescent molecules excited by the outer airy rings of the illuminating
beam will be blocked by the confocal aperture; like all light, the illumination
beam also presents with a airy wave pattern consisting of a central bright spot
and outer ring waves that in comparison are more faint. In other words, the
resultant fluorescence emission may be experimentally manipulated to originate
from an area smaller than the airy unit, which cannot be achieved by conven-
tional widefield epifluorescence microscopy. Moreover, because the PSF of the
confocal microscope is narrower at normalized light intensities relative to that of
the conventional widefield microscope, it means the XY spatial resolution will be
�1.4�greater with a confocal microscope than a widefield microscope
(Conchello and Lichtman, 2005). Finally, some of the in-focus light will scatter
on its way through the specimen, due to diVraction, reflection, and refraction as
cell structures interfere with the light path. This also compromises fluorescence,
but not confocal microscopy, as the confocal aperture also blocks scattered light
from reaching the signal detector.
In addition, combining confocal microscopy with total internal reflection fluo-
rescence (TIRF) or Forster resonance energy transfer (FRET) microscopy techni-
ques has the capacity to increase resolution to only a few tens of nanometers
(see below for more detailed information). Other techniques such as narrowing
the boundaries of the PSF by suppressing (de-exciting) the fluorescence from the
edge of the center spot of the airy pattern by stimulated emission depletion
(STED), and other nonlinear optical masking techniques, have further enhanced
optical resolution of confocal microscopes (Bullen, 2008; Willig et al., 2006),
though these techniques are not yet compatible with fast scanning of Ca2þ events
that take place over fast timescales, and will therefore not be discussed here.
Finally, secondary signal processing or deconvolution (computationally reverse
optical distortion to enhance resolution) of the recorded images also serve to
enhance spatial resolution of both confocal and epifluorescence microscopy by a
factor of 2–3.
Because light scattering increases proportionally to increasing thickness of the
specimen, this becomes more of an issue with deeper imaging of thicker speci-
mens. Therefore, appropriately setting the pinhole not only allows for imaging of
thin optical sections, but also aVects the signal-to-noise (SNR) ratio. Whereas a
case made be made that reducing the pinhole diameter increases XYZ-resolution
(especially Z-, but also XY-resolution; see above), opening the pinhole to ap-
proximately match the projected image of the diVraction-limited spot (1.22l/NA,
230 Godfrey Smith et al.
where l is the illumination wavelength) will substantially increase the SNR with
only minimal reduction in the Z-resolution (Conchello et al., 1994). Thus, this
would increase the quality of the signal with little degradation of the depth
discrimination.
III. Limitations in Speed of Confocal Imaging
The removal of out-of-focus light allows for relatively fast imaging of �0.6 mmthin sections either in spot, line, or frame modes that either may be repeated
sequentially, or combined with stepwise up- or down-focusing through the speci-
men in order to generate 3D reconstructions. 3D sectioning does not allow for
recording of cellular events in real time, but repeated 1D line imaging or even
repeated 2D imaging with reduced frame sizes or restricted pixel numbers in
contrast do allow for relatively fast recording with a temporal resolution of
approaching a microsecond scale. Although not as fast as regular widefield imag-
ing, even 2D frame imaging may still be acquired on fast time scales, usually within
hundreds of millisecond, though there will be a trade-oV between temporal and
spatial resolution. The reason for the lower temporal resolution compared to
widefield imaging is that conventional confocal imaging requires some form of
scanning, that is, sequential pixel sampling, in order to ‘‘build’’ an image, which
thus happens pixel-by-pixel. In contrast, the whole field during widefield imaging is
captured simultaneously either by PMTs or charge-coupled device (CCD) cameras
(Ogden, 1994). For some Ca2þ events, imaging with a temporal resolution in the
order of milliseconds may be satisfactory, but other events may occur considerably
faster than this. Likewise, some events allow the microscopist to sample images
with a low spatial resolution, whereas others require the opposite. Thus, the speed
of confocal image acquisition depends on the mode and the settings of the scanning
and how many pixels are scanned before returning to the same pixel again.
This will be discussed later.
IV. Laser Scanning Confocal Microscopy
Out-of-focus light rejection and image acquisition through a confocal aperture
with a pinhole is the common principle that constitutes confocal microscopes, but
the illumination and excitation principles may diVer between various systems.
First, confocal microscopy by laser scanning the specimen (laser scanning confocal
microscopy, often abbreviated to LSM or LSCM) is the most widely used illumi-
nation and excitation method today.
During LSCM, a laser beam is directed on to the specimen, whereupon it scans
the designated field, which may be a single spot (in reality rarely used for biological
imaging apart from fluorescence recovery after photobleaching (FRAP) applica-
tions), a 1D line, or a 2D frame. The laser is controlled by the use of two oscillating
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 231
mirrors in the scanhead that deflect the beam along a fast and slow axes perpen-
dicular to one another. Thus, during a 2D frame scan, the beam is first directed
along the horizontal axis, after which it ‘‘jumps’’ down one pixel and scans the next
line, and this process continues until a full frame has been scanned. This may then
be repeated for serial frame scanning, or the plane of focus may be moved along
the Z-axis for 3D imaging. The fluorescence emission returns along the same light
path (descanning), but is deflected by a dichroic mirror splitting the excitation and
emission lights, such that the emission light only is directed onto the confocal
aperture, where the in-focus light penetrates though the pinhole to reach the signal
detector.
Unfortunately, scanning is a rate-limiting step for gaining fast images, especially
in a 2D frame scan mode, because of the mechanical characteristics of the mirrors.
A typical 512�512 pixel 2D frame may be scanned in �1 s, depending on the
settings under which the scan is performed. When scanspeed is of concern, several
approaches may be taken to increase this, for example, by linescanning instead of
framescanning. In this configuration, the same line is scanned sequentially for a
given period of time to allow detection of one or more events occurring within the
time frame of the scan. Although this provides a high temporal resolution, the
acquired information is limited to a one-pixel wide area of the cell, such that
information about events occurring elsewhere in the cell is missed. Furthermore,
reducing the pixel dwell time (the period over which each pixel is scanned) also
increases scanspeed, but this also reduces the SNR. Reduced SNRmay partially be
compensated for by increasing the laser power, but this may present other problems
such as photodamage to the specimen; especially in live cells, and photobleaching.
Finally, the length of the line or the size of the frame to be scanned may also be
reduced, or fewer pixelsmay be scanned during 2D frame scanning to yield the same
eVect, and the scan may also be run in a bidirectional mode instead of unidirection-
al, though using two lines running in opposite directions may cause a slight oVsetbetween them, which tends to blur the signal. More recently, several approaches
have been taken to increase scanspeeds, in particular for 2D frame imaging, such as
utilizing resonant oscillating mirrors in the scanhead instead of the more conven-
tional galvanometer-driven mechanical mirrors. Other options include arranging
prisms and acousto-optical deflectors into the excitation light path to illuminate the
entire line simultaneously, instead of a pixel-by-pixel illumination applied by the
conventional laser scanning microscopes described above. This means that the
scanning in a 2D frame mode would only involve movement along one dimension
(X), since the other dimension (Y) would all be scanned at once (simultaneously),
and therefore, 2D frame scanning may be performed at linescan speeds or at speeds
approaching video rates, if the emitted fluorescence is deflected onto a linear CCD
camera, though PMTarraysmay also be used. Single PMTswould, however, not be
able to construct the images if lines instead of single pixels are scanned. The caveat
with these approaches is that true confocality will be lost because the pinhole of the
confocal aperture must be replaced by one or more longitudinal slit openings to
accommodate the simultaneous scanning of lines (hence, this is also called slit
232 Godfrey Smith et al.
scanning). Also, series of holographic or curved mirrors in the scanhead have also
been utilized to scan more than one pixel at a time, but this has remained more of a
rarity compared to the abovementionedmicroscopymodifications (Callamaras and
Parker, 1999; Tsien and Bacskai, 1995).
The aforementioned confocal scanning approaches depend on the use of single-
photon lasers as the source of illumination and excitation. These are based on the
principle that a single photon provides enough energy to excite a single fluorescent
molecule, that is, to ‘‘lift’’ it from a ground state to the ‘‘excited’’ state. The phase
where the fluorophore is lifted to the excited state lasts for femtoseconds (10�15 s),
whereas the fluorophore remains in the higher-energy excited state for picoseconds
(10�12 s) where it undergoes internal conversion and starts to vibrate, which eVec-tively leads to dissipation of energy, such that it drops back to the ground state;
measurable to a time scale of nanoseconds (10�9 s). When this happens, the
fluorophore releases a photon that due of the loss of energy has a longer wavelength
(less energy), and this is what creates the fluorescence emission that may be
measured by signal detectors such as PMTs or CCD cameras. The diVerencebetween the excitation and emission spectra (emission wavelengths being longer
than excitation wavelengths) is called the Stokes shift. The process of excitation and
subsequent relaxation with photon release and fluorescence emission can be illu-
strated by a Jablonski diagram (Fig. 2), and is not restricted to confocal microsco-
py, but is in fact the basis for all fluorescence techniques including epifluorescence
microscopy and spectroscopy. As detailed above, it is the volume of the recorded
fluorescent emission that diVers between confocal and epifluorescence microscopy
modalities, though the volume of excitation may also diVer, but this has to do with
how much of the specimen is subjected to the illumination light. However,
although the laser excites fluorophores along the entire Z-axis of the specimen
(see also above and Fig. 3), peak excitation and as such peak brightness occurs at
the focal plane, whereas out-of-focus excitation decreases with the square of the
distance from the focal plane. This is because the laser excitation beam presents
with an hourglass shape, with the ‘‘waist’’ of the hourglass coinciding exactly with
the focal plane.
Several laser lines have been developed that allow single-photon excitation
of fluorescent Ca2þ indicators (fluorophores), in particular, the multiline
argon ion (Ar-ion) laser that provides high-intensity light from the ultraviolet
(UV) to the green spectrum (�250–514 nm wavelengths), the single-line helium–
neon (He–Ne) lasers that extend the covered spectrum to �633 nm, and argon–
krypton (Ar–Kr) lasers that providehigh-intensity light fromblue to redwavelengths.
Thus, these lasers are well suited for exciting the common Ca2þ indicators dyes and
are also as such much used in Ca2þ signaling research. Recent developments in solid
state and diode lasers have also added more choices for the microscopist. However,
lasers will not be covered in detail here, but the interested reader will find a wealth of
literature on this topic by searching the appropriate literature databases ormicrosco-
py textbooks, literature that covers the topic from both physics and biology
perspectives.
Internal conversion, vibration state(loss of energy) (ps)
High energyexcited states
Absorption(excitation)(fs)
Absorption(excitation)
Wavelength
Rel
ativ
e in
tens
ity
Excitation light
B
A
Low energyground state
Stokes shift
Emission
Emission(ns)
Emission light(longer wavelength)
Fig. 2 (A) A Jablonski diagram showing the energy states of a given fluorophore, including the
process of absorption and emission of longer wavelength light upon excitation. The duration of each
state is also indicated. (B) The absorption and emission spectra of a given fluorophore including the
Stoke’s shift due to emission of a longer wavelength photon upon excitation of the fluorophore.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 233
V. Total Internal Reflection Fluorescence Microscopy
Confocal microscopy coupled to TIRF provides a very thin optical section
of fluorescence excitation that allows imaging with low background noise and
minimal out-of-focus fluorescence. This is because total internal reflection can
only occur when the excitation light beam in a medium of high refractive index
reaches an interface of a medium with a lower refractive index at an angle of
incidence that is greater than the specific critical angle y. When the light is
totally internally reflected, none of it penetrates the medium with the lower
refractive index, and ideally, there is no net energy flux escaping the glass.
Incident light
Incident light
Range of incidenceangles greater thanthe critical angle q
Specimen, low refractive index
Glass, high refractive index
Reflected light
Evanescent field<100 nm
Reflected light
Objective
SpecimenGlass
A
B
Fig. 3 Total internal reflection fluorescence (TIRF) microscopy. (A) Overview including the inci-
dent and reflected laser light paths within the objective. (B) Once the incident light reaches a medium
with a lower refractive index at an angle greater than the critical angle y, the incident light does not
penetrate the specimen, but an electromagnetic field is created that penetrates up to ~100 nm above
the surface, called an evanescent wave. This may excite fluorophores within the range of the evanes-
cent wave.
234 Godfrey Smith et al.
However, the reflected light generates an electromagnetic field that penetrates
beyond the interface and into the lower refractive index medium as an evanes-
cent wave. This wave, with a wavelength similar to the excitation light beam,
decreases exponentially with the distance into the medium. The penetration
depth of the evanescent wave may be manipulated by changing the angle of
incidence beyond the critical angle, which may be calculated accurately by
knowing the angle of incidence, but will typically be limited to �100 nm or
less (Cleemann et al., 1997; Mashanov et al., 2003). This allows for imaging of
Ca2þ events occurring in close proximity to the plasma membrane of live cells.
However, this does require that the cell is positioned within the evanescent wave
and not above it on the glass surface, which may well be the case for the
majority of the cell if the cell is of a certain size. In our experience, it requires
an experimental eVort to physically position large cells within the evanescent
wave and yet still maintain physiological conditions for the cell. This is required
because the evanescent wave is generated from the glass surface and not from
the boundary of the cell.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 235
VI. Forster Resonance Energy Transfer Microscopy
Confocal microscopy coupled with FRET imaging has experienced something
of a surge of interest recently fromCa2þ researchers, because of the development of
Ca2þ-sensitive cameleons. The FRET process is based upon nonradiative energy
transfer from a fluorophore in an excited state (‘‘donor’’) to another chromophore
(‘‘acceptor’’) that usually is, but does not have to be, a diVerent fluorescent
molecule within a range of 10–100 Angstroms (1–10 nm) (fluorophore) (Jares-
Erijman and Jovin, 2003). Thus, when FRET occurs, it is not only the acceptor
emission that will be recordable, but the measurable emission from the donor will
also be greatly reduced because of the energy transfer. Also, the intensity of the two
emission bands will depend on the distance between the two donor and acceptor
fluorescent proteins or molecules even within the FRET distance. The closer the
distance between the donor and acceptor, the greater the longer-wavelength emis-
sion (from the acceptor) and the less the shorter-wavelength emission (from the
donor). The fluorescence emission from the acceptor is discernible from the donor
emission due to the spectral redshift, such that the instrumental requirement is that
the two emission bands must be separately recordable. The presence of the longer-
wavelength acceptor emission will confirm that the acceptor molecule is within
the FRET distance of the donor, that is, within a distance of �10 nm. Likewise,
the absence of it suggests that the physical distance between the donor and
acceptor is larger than that. Traditionally, FRET imaging has been used to study
protein–protein interactions by tagging diVerent proteins with fluorescent probes
(e.g., yellow or green fluorescent proteins (YFP or GFP)) and thereby study
whether they exist within or outwith the FRET distance.
However, it is the recent development of cameleons that has moved FRET
imaging to also become a sought-after technique with respect to studies of Ca2þ.Cameleons are genetically encoded fluorescent proteins that are sensitive to Ca2þ,in which a blue- or cyan-mutant of GFP (serving as the donor), calmodulin that
can bind to Ca2þ, a calmodulin-binding peptide such as the calmodulin-binding
domain of skeletal muscle myosin light chain kinase (the M13 peptide), and a
long-wavelength yellow mutant or normal GFP (serving as the acceptor) are
tandemly fused (Miyawaki et al., 1997). This configuration forms a stable and
compact complex that remains intact once transfected into the target cell, and
subsequent development has also improved the spectral properties and rendered
the cameleons less susceptible to changes in pH (enhanced GFPs). In the absence
of Ca2þ, the cameleon remains in its linear tandem configuration, whereby the
two GFP mutants (donor and acceptor) at the two flanks of the tandem are too
far apart to create FRET. However, when the concentration of free Ca2þ
increases, calmodulin binds to Ca2þ and undergoes a conformational change
that also leads it to bind and wrap around the M13 peptide, which creates a
compact configuration of the cameleon and therefore brings the donor and
acceptor GFP mutants to within a distance where energy transfer may occur
236 Godfrey Smith et al.
with much greater eYciency. Thus, the presence of Ca2þ creates FRET by the
cameleon, which may be readily detected. Moreover, because Ca2þ aYnity can be
tuned by incorporating mutations into the calmodulin protein, cameleons may
detect free Ca2þ concentrations in the range 10 nM–10 mM, and this has been
done to visualize local Ca2þ signals in the nucleus, sarco/endoplasmic reticulum,
and the cytosol, by transfecting chimeras of the cameleon that also have the
appropriate localization signals encoded in the complementary DNA (Miyawaki
et al., 1997). A typical example of a cameleon used for FRET imaging of Ca2þ has
an excitation spectrum peak at 442 nm for the donor (blue mutant of GFP), with
the associated emission peaking at 486 nm. This wavelength FRET-excites the
GFP on the acceptor side, which then emits longer wavelength fluorescence with a
peak at 530 nm (Fig. 4).
Several protocols for detecting and measuring FRET eYciency have been
developed, of which some are more applicable to imaging of Ca2þ signals than
others. These include measuring donor quenching, that is, measuring the decrease
in the emission from the donor fluorophore, which appears because some of the
energy emitted by the donor fluorophore is used to excite the acceptor chromo-
phore/fluorophore. This is done by taking the ratio between donor and acceptor
fluorescence during FRET as the numerator, and the same ratio in the absence of
FRET (i.e., by removing either of the donor or acceptor fluorophores) as the
denominator. Like all ratiometric quantifications, this has the advantage that the
measure becomes independent of local variations in fluorescence. The disadvan-
tage is that both the donor and acceptor fluorophores may be quenched by other
factors that would misrepresent the results. Another method for measuring FRET
eYciency is by measuring donor quenching and acceptor photobleaching, that is,
the intensity of the donor emission in the presence of an acceptor relative to the
intensity of the donor emission in the absence of an acceptor. In practice, the
latter is done by first photobleaching the acceptor by illuminating it with light at
the peak excitation spectrum of the acceptor fluorophore before measuring the
donor emission. This protocol relies on the fact that fluorescing itself causes the
fluorophores to lose their ability to fluoresce, a process called photobleaching.
However, because photobleaching may take many minutes to achieve (up to
�20 min), this may become less available in live specimens. Finally, FRET
combined with fluorescence lifetime imaging (FLIM) has introduced a robust
option for Ca2þ measurements, because it largely is unaVected by experimental
conditions such as fluorophore concentrations and excitation intensities. In this
combined FRET–FLIM approach, the change in donor lifetime is measured in
the presence and absence of an acceptor. The principle behind this is that the
period of time the donor will fluoresce (i.e., the lifetime of the donor) depends on
the presence or absence of an acceptor (Levitt et al., 2009). As described above,
the measurement of donor emission in the absence of an acceptor may be done by
first photobleaching the acceptor. However, once the photobleached control
images have been captured, this protocol allows for detailed imaging of Ca2þ
signals over a short time period.
Donor:Blue orcyan FP Cam
Excitation:
A
B
Emission:
Excitation:
486 nm 530 nm
Wavelength
Rel
ativ
e em
issi
on in
tens
ity
Emission:
FRET<100 nm
0 Ca2+ +4 Ca2+
4 Ca2+
+Ca2+
−Ca2+
442 nm
442 nm
486 nm
530 nm
M13
Acceptor:Green oryellow FP
Fig. 4 Cameleon-based Forster resonance energy transfer (FRET). (A) Schematic of the cameleon in
the absence and presence of Ca2þ; note the conformational change in the calmodulin (Cam) and the
Cam-binding domain of myosin light chain kinase (M13) upon binding to Ca2þ that allows for FRET
between the donor and acceptor green fluorescent protein (FP) mutants. (B) The relative emission
intensities at different wavelengths indicate whether or not Ca2þ is present, and hence, FRET occurs.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 237
Since the introduction of cameleons, derivatives have been developed that fuse
Cam with the Cam-binding peptide M13 in a similar fashion as cameleons, but are
based on single GFPs instead of two GFP mutants with diVerent spectral proper-ties that necessitate Ca2þ imaging by FRET. These derivatives, called pericams,
can therefore be imaged by standard epifluorescence or confocal microscopes, and
present with broad Ca2þ sensitivities and diVerent localization sequences that
allow for measurements of a range of Ca2þ concentrations within specific orga-
nelles and intracellular targets (Kettlewell et al., 2009; Nagai et al., 2001).
238 Godfrey Smith et al.
VII. Parallel Scanning Confocal Systems
The main limitation of the traditional laser scanning confocal microscope sys-
tems discussed so far is their low full frame temporal resolution. At typical rates of
just a few Hertz, an increase of 1–2 orders of magnitude in the speed of image
acquisition is necessary to decipher the fast Ca2þ dynamics of systems such as
neuronal networks or cardiac and muscle tissues.
Recent developments in parallel multispot confocal illumination devices have
gone some way to addressing these concerns, providing high contrast optical
sectioning with typical rates in the 10–100 Hz domain. The multispot system uses
an aperture mask at an illumination plane conjugate with the sample, where
multiple illumination points with nonoverlapping airy disk profiles are projected
to the sample simultaneously. This illumination pattern is then changed in a
sequential fashion such that every image point is uniformly illuminated in a
given time interval. Two approaches have been used in recent years to successfully
achieve this fast pattern change: the Nipkow spinning disk where a patterned disk is
spun at high speeds, and programmable matrix systems (digital mirror and liquid
crystal arrays) where individual pixels can be switched ‘‘on’’ and ‘‘oV’’ at very highspeeds.
The parallel illumination approach has the obvious advantage that all image points
can be illuminated much faster than in a conventional single scan system, but the
drawback is that a 2D imaging detector is required to record the image, which
then becomes the limiting factor for capturing image frames. Limitations of the
detectors (typically CCD cameras) are further exacerbated by the requirement for
fast detection, as these devices have a noise floor below which the fluorescence signal
cannot be resolved. The fundamental problem is that in order to increase frame rates,
the exposure time of a frame has to be reduced, thus limiting the number of detectable
photons in an integration interval. Simultaneously, the transfer rate of data from
the camera has to be increased, a process by which the read noise of CCD cameras
increases. Many of these concerns are resolved by modern high-end electron multi-
plication (EM) CCD cameras such as the iXon 897 (Andor), Evolve (Photometrics),
or ImagEM (Hamamatsu). These EM-CCD cameras can amplify small photoelec-
tron signals above the read noise of the camera; providing the sensitivity and speed
required making these theoretical parallel approaches a practical reality.
VIII. Spinning Disk Confocal Microscopy
These systems illuminate the specimen simultaneously with a large number of
non-overlapping points of light by using multiple (hundreds to thousands) pinholes
arranged in a geometrically precise spiral pattern on a spinning disk (Nipkow disk).
The disk is placed at an image plane conjugate with the sample, and the illumination
is filtered by this disk. Thus, the specimen is raster scanned rather than single-spot
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 239
scanned, such that the entire 2D frame may be illuminated semi-simultaneously.
Fluorescence emitted from each illumination point then returns through the same
aperture of the mask to give a set of confocal points in the image. When the disk is
spun at high speed (typically several thousand revolutions per minute), an appar-
ently continuous image is obtained when viewed through an eyepiece. The resultant
image is traversed with scan lines, but precise synchronization of the illumination
and detection systems virtually eliminates this artifact in well configured systems
(Wilson et al., 1996). Thus, this approach allows for very fast scan rates without
significantly compromising the SNR (Wang et al., 2005).
As the input illumination power is spread over a much larger area than with
conventional scanning confocal systems, light throughput can be a serious impedi-
ment to successful implementation of spinning disk systems. This was partially
resolved by enhancing the original design by means of a second disk spinning in
sequence with the Nipkow pinhole array disk. This second disk is sited between the
dichroic and the light source and contains a microlens array that maps a miniature
lens to each pinhole (Tanaami et al., 2002), thus improving the illumination
eYciency by focusing the lightbeam onto the pinhole (Fig. 5A). This also reduces
the backscattering of light at the surface of the Nipkow disk, which substantially
increases SNR. The emission detection pathway is not aVected by this modifica-
tion. Use of specialist cameras and fast versions of the spinning disk head can now
enable imaging rates of up to 2 kHz. However, the drawbacks are that the pinholes
on the spinning disk are inflexible, and the dwell time per pixel is usually very short
(�100 ns), which may severely reduce the SNR, although the frequent illumination
of the same pixel as the disk rotates may compensate for this eVect.
IX. Programmable Matrix Microscopy
These instruments are based on the principle of spatially filtering full field
illumination in a defined pattern at high speed, so as to give it a prescribed,
dynamic structure. By using appropriate filtering patterns, the device can simu-
late the optical behavior of confocal scanning microscopes. These systems are
directly comparable to the spinning disk approach in that the illumination device
consists of an array of small apertures that act as both the illumination and
detection pinholes. The principal diVerence, and advantage, is that the elements
of the array are individually addressable, allowing far greater flexibility in exper-
imental design compared to the spinning disk (Hanley et al., 1999). As a practical
example, selectively and sequentially illuminating individual cells within the field
is possible with the array system, whereas the Nipkow disk can only operate at
full frame.
Two technologies have been used in implementing practical programmable
matrix systems. The first makes use of a digital micromirror device (DMD), an
array of micrometer-sized mirrors whose angle can be independently controlled
to direct illumination to an ‘‘on’’ (confocal) or ‘‘oV’’ (non-confocal) pathway.
A
Nipkow pinhole array disk
Objective
Lightdetector 1(conjugate)
B
Lightdetector 2(nonconjugate)
Light source
Dichroicbeamsplitter
Dichroicbeamsplitter
Dichroicbeamsplitter
Dichroicbeamsplitter
Objective
ReflectiveLCOS array
Light detector
Emitted light
Excitationlight beam
Microlens array disk
Microlenspinhole
Fig. 5 (A) Nipkow spinning disk. The figure shows two light beams reaching the specimen and the
light detector, although, in reality, multiple beams reach the specimen and the light detector simulta-
neously. (B) The optical pathway of a programmable matrix microscope. The liquid crystal on silicon
(LCOS) array consists of numerous mm-sized liquid crystal ‘‘pixels’’ that can be switched independently
to reflect the lightbeam onto multiple spots simultaneously.
240 Godfrey Smith et al.
The alternative uses a reflective liquid crystal display (LCD), an array of microm-
eter-sized liquid crystal ‘‘pixels,’’ the base of which is coated with a reflective layer
(Fig. 5B). Each element is individually addressable, creating a controlled polariza-
tion pattern that is optically transformed into a confocal intensity pattern. So far,
the liquid crystal approach forms the basis for the commercial utilization of this
technique. This allows for using both the confocal image (conjugate with the array
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 241
mask) and nonconjugate image to provide further potential enhancements in
dynamic image quality (Heintzmann et al., 2003).
The real power of the programmable matrix approach lies in the ability to
redefine the array composition, without any additional hardware changes. This
enables complete flexibility for optimization of the illumination pattern for the
sample and experiments in hand—from low speed high resolution to high speed
lower resolution studies.
X. Advantages and Disadvantages of Confocal Microscopy
From an experimental point of view, confocal Ca2þ imaging oVers many benefits
over conventional fluorescence microscopy, especially with respect to spatial and
temporal resolution and the ability to optically section the specimen along the
Z-axis down to a resolution of 0.5–1 mm, enabled by the presence of a confocal
aperture with a pinhole in the light path that eVectively gives this imaging modality
its uniqueness, as well as its name. Importantly, confocal microscopy may be
performed in live cells that are electrically, mechanically, and by all other mea-
sures, viable. Moreover, confocal microscopy does not impair the ability to
manipulate the surrounding environment in terms of solutions in which the cells
or specimen are bathed, solution pens, temperature devices, application of phar-
macological drugs and inhibitors, patch pipettes, electrical and mechanical manip-
ulators, etc., as long as these do not interfere with the actual light pathway.
However, the significant drawbacks with confocal microscopy relate firstly to the
unavoidable scattering as well as chromatic aberration with subsequent focal plane
oVsetting that reduces confocal signal collection (Bliton et al., 1993) of the illumi-
nation light as it penetrates the specimen, especially since wavelengths of
�300–600 nm are commonly used for excitation, as those are very prone to
scattering; eVectively limiting the penetration depth to �40–50 mm (or less,
depending on the laser output and the spectral properties of the specimen), and
secondly to the fact that although only the emitted fluorescence produced at or
very near to the focal point will be recorded, excitation will still occur along the
whole Z-axis of the specimen (given molecules able to be excited are present along
the Z-axis). The latter point presents often considerable problems because the
photodamage generated along the entire Z-axis may kill live specimens. Though
this may be mitigated by lowering the laser power, the restricted light capturing
from the small volume accommodated by the pinhole tends to drive toward
microscopists increasing laser power. This point is further accentuated by excita-
tion at short wavelengths (300–500 nm) including UV light, as those are severely
damaging to live specimens due to the associated high energy. Finally, higher
purchase and maintenance costs relative to conventional epifluorescence and wide-
field microscopy may also represent a disadvantage to the researcher.
A diVerent excitation approach with many of the same advantages as confocal
laser scanning microscopes as described above, while also substantially limiting the
242 Godfrey Smith et al.
major drawbacks of confocal microscopy such as the restricted penetration depth
and photodamage, is oVered by multiphoton microscopy systems.
XI. Multiphoton Excitation Laser Scanning Microscopy
A conceptually diVerent microscope that also allows optical sectioning and high
spatial and temporal resolution live specimen imaging, and that also comes with
other added benefits, is the multiphoton microscope. From a biophysical perspec-
tive, multiphoton imaging rests on an excitation principle that relies on excitation
of the fluorescent molecule by photons that alone do not have enough energy, but
need to combine by simultaneously coming in to a very close proximity of the
fluorophore.
Although the theoretical concept of multiphoton excitation is relatively simple and
has been known and also utilized by physicists for many decades, biological applica-
tions ofmultiphoton excitation are of amore recent date (Denk et al., 1990).Whereas
in confocal single-photon laser illumination, the fluorophore is excited by the absorp-
tion of a single photon, as it provides suYcient energy for the fluorophore to reach an
excited state. Upon return to the ground state, a photon of longer wavelength than
the excitation photon is emitted, and it is this process that creates the fluorescence.
With multiphoton excitation, a fluorophore is excited by the near-simultaneous
(within 10�18 s) absorption of two or more photons that combined provide enough
energy to promote the fluorescent molecule from a ground state to an excited state;
two photons for two-photon (2P) excitation, three photons for three-photon (3P)
excitation, etc., with 2P excitation being by far the most common multiphoton
excitation modality. Thus, with 2P excitation, the fluorophore absorbs two photons
simultaneously, each twice the wavelength and half the energy required formolecular
excitation, and likewise, in 3P excitation, each of the three excitation photons have
three times the wavelength, but only one-third of the energy compared to single-
photon excitation (Helmchen and Denk, 2005).
Once excited, the emitted fluorescence is then proportional to the square of the
excitation intensity in 2P absorption (third power in 3P excitation), and this 2P
excitation (measured in Goeppert-Meyer Units) occurs only at the focal point, as
it is only here that the density of the excitation photons is high enough to ensure
a simultaneous photon arrival to the fluorophore that is suYcient to excite it.
Though this may not occur with all photons in the volume, the probability of it
occurring for the vast majority of the photons is very high. This nonlinear
excitation constitutes the most important physical diVerence from confocal sin-
gle-photon excitation, in that excitation will be confined to a small ellipsoid
volume around the focal point, whereas above and below this point, the density
of photons suYciently close to one another, or in other words the light intensity,
will not be high enough to generate any excitation (Fig. 6). This eVectively means
that out-of-focus fluorescence will not be generated, which again removes the
need for a confocal aperture with a pinhole in the emission light pathway.
Single-photon
BContinuous single-photon laser
Femtosecond pulsed 2P laser
Time
Time
100 fs
10 ns
Excitation
Excitation and emission intensitiesPhotonsFocal point
Emission2P
Single-photon 2P Single-photon 2P
A
Fig. 6 Two-photon (2P) excitation microscopy, including a comparison to single-photon confocal
microscopy. (A) Schematic of illumination lightbeams, in which excitation occurs along the whole Z-
axis with single-photon confocal microscopy, though with highest intensity at the focal point, whereas
excitation is confined to a narrow area around the focal point with 2P microscopy. Correspondingly,
emission occurs along the entire z-axis with confocal microscopy; though out-of-focus light is blocked
by the confocal aperture, whereas emission is confined to a narrow area around the focal point with 2P
excitation microscopy. (B) Schematic of the continuous laser used for single-photon excitation and a
pulsed titanium:sapphire (Ti:Sapphire) laser used for 2P excitation.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 243
Therefore, the optical sectioning and spatial and temporal resolution one may
achieve will be close to that achieved by confocal microscopy or a little poorer,
but not better (Cox and Sheppard, 2004). A clear benefit of this is that there need
not be a confocal aperture. Although crucial for confocal single-photon micros-
copy, confocal apertures themselves reduce light transmission, but also require
that the emission light path is returned through the scanhead including the
mirrors, which further causes a loss of light. Instead, the emitted fluorescence
after 2P excitation may be focused directly onto a PMT without having to be
descanned, as in confocal microscopy, such that 2P microscope systems are
generally more light sensitive. However, the introduction of a confocal aperture
with a pinhole into the 2P emission pathway may improve the spatial resolution
to a confocal standard.
244 Godfrey Smith et al.
Thus, although the capture of the emitted fluorescence may be set up with
diVerent light paths in 2P microsscopes compared to confocal single-photon
microscopes, it is conceptually similar in that emission light needs to be split by
the excitation light with the insertion of an appropriate dichroic mirror. However,
in the case of 2P microscopy, the emitted fluorescence will in most cases have a very
much shorter wavelength than the excitation wavelength, which is in contrast to
confocal single-photon and conventional epifluorescence microscopes.
Because of the abovementioned properties of 2P excitation and diVerences to
single-photon confocal excitation, 2P excitation oVers several important advan-
tages over single-photon excitation. First, because light scattering declines steeply
with increasing wavelengths, red and IR (670–1100 nm) light can penetrate and
hence excite fluorescent molecules much deeper into the specimen than single-
photon lasers providing excitation wavelengths in the range from �300 to
�600 nm. With suYcient power outputs and optimized optics including corrective
optics for pulse dispersion, penetration depths approaching 1000 mm (1 mm) may
be achieved, which is considerably deeper than the �40–50 mm achievable with
single-photon excitation. Thus, 2P excitation secures an up to �20-fold deeper
penetration depths by using excitation light of twice the wavelength. To achieve
comparable deep imaging with single-photon confocal microscopy, tissue or layer
removal with histologic techniques or penetration by the objective would have been
required. Such mechanical approaches would for obvious reasons compromise the
‘‘intactness’’ and viability of the tissue. For this reason, confocal microscopy is
mainly restricted to studies of single, isolated cells or tissue surfaces, whereas 2P
excitation laser scanning microscopy allows deep tissue imaging of intact organs.
Similarly, 2P excitation microscopes are comparably more often upright rather
than inverted, to allow for tissue preparation to be set up on the stage, whereupon
the objective lens is lowered down to within the optical range for imaging. The
opposite is true for confocal microscopes, which more often than not come
inverted. Furthermore, even with 2P excitation deep tissue imaging, one can be
assured that the vast majority of the fluorescence comes from the focal point and
not from residual scattering (scattering is greatly reduced, but not obliterated, by
long-wavelength excitation). This is because, even in strongly scattering tissue such
as the heart, the density of scattering exciting photons is too low to generate
significant fluorescence. Therefore, this together with the long excitation wave-
lengths also contributes to 2P excitation microscopes being much less sensitive to
light scattering than regular widefield epifluorescence or confocal microscopes.
However, it should be noted that penetration depths also heavily depend on the
specimen, as diVerent tissue properties may degrade the illumination as well as the
emitted fluorescent light. In particular, collagen and myelin are known to scatter
light and thereby restrict penetration (Helmchen and Denk, 2005).
Additionally, IR light is much less phototoxic than shorter wavelengths, such
that it correspondingly may cause only negligible photodamage. Also, because of
the nonlinear excitation, photodamage and bleaching are restricted to the focal
point. Importantly, the reduced photodamage and fluorophore bleaching allows
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 245
for longer imaging periods. 2P excitation spectra of most fluorophores are wider
than the equivalent single-photon excitation spectra, whereas the associated emis-
sion spectra after 2P and single-photon excitation are not diVerent, such that the
same excitation wavelength may potentially excite several fluorophores with dis-
tinct emission spectra (Bestvater et al., 2002; Xu et al., 1996). This property will be
explored in more detail later in the chapter. Moreover, whereas the positioning of
the confocal aperture relative to the light detector as well as the focal plane is
critical to ensure that the images of the source and detector apertures are cofo-
cused, this is not the case for 2P excitation microscopes, in which the position of
the light detector is not critical, because fluorescence emission will only be gener-
ated at and tightly around the focal volume.
However, several problems persist with 2P excitation. First, 2P excitation requires
very high light intensities; intensities that would instantly vaporize the specimen if
the light was delivered continuously. However, this is overcome by using lasers that
provide ultrabrief (10�13 s) pulses at very high frequencies (�80–100 MHz), which
thereby generate very high instantaneous energy, but suYciently low average energy
to avoid any substantial damage to the specimen. Thus, the distance between each
pulse is typically �10 ns, whereas the width of each pulse itself is typically 100 fs
(Fig. 6). The high repetition rates match fluorescence lifetimes closely (see above)
such that a good balance between excitation eYciency and onset of saturation is
achieved (Helmchen and Denk, 2005). The most widely used lasers that fulfill this
criterion and provide the necessary wavelengths, are the solid state titanium:sap-
phire (Ti:Sapphire) oscillating (pulsed) lasers. These lasers are tunable, such that the
latest versions are capable of delivering wavelengths within the range �670–
1100 nm, thus, from visible red to infrared (IR), though this range is continuously
expanding while also coming with higher power outputs, as manufacturers keep
developing their product lines. Though most 2P excitation applications may not be
power-limited, more available power does benefit applications requiring deep
tissue penetration. Typical power outputs in the latest Ti:Sapphire lasersmay exceed
4 W at 800 nm, though the optics cause a major loss in power from the outlet of the
laser to the focal point of the objective lens. Glass components of the optical
pathway also cause a dispersion of the �100 fs laser pulses, but this may be
compensated for by corrective optics in order to maximize 2P excitation (Diels
et al., 1985). Other limitations of 2P excitation microscopy are that reflected light
imaging is not possible; only fluorescence imaging, and that it is not suitable for
imaging highly pigmented specimens, as these absorb IR and near-IR light.
XII. Ca2þ Indicators for Use in Confocal andMultiphoton Microscopy
For many years, biology has benefited from the fluorescent dyes or constructs
that bind and therefore can measure the free concentration of Ca2þ [Ca2þ]. Theseindicators have been used to examine the levels and time course of changes
246 Godfrey Smith et al.
in [Ca2þ] within the cytosol and the various organelles within the cell (e.g.,
nucleus, mitochondria, sarco/endoplasmic reticulum). The two main categories
of molecules used for this purpose are small synthetic organic molecules based
on the fast Ca2þ buVer BAPTA (Tsien, 1980) or modified versions of natural
Ca2þ binding proteins (Miyawaki et al., 2003). Both categories ‘‘sense’’ Ca2þ by
chelating the ion which changes the structure/chemical properties of the ligands.
Several modes of fluorescence have been utilized to report Ca2þ (summarized in
Fig. 7).
Absorbance/quantum yield: In this case, Ca2þ binding causes a change in the
intensity of the fluorescence from the dye; typically, Ca2þ binding causes an
increase in fluorescence (Fig. 7A). This mode is the most common employed by
the fluorescent Ca2þ indicators used in confocal or 2P microscopy. In particular,
+Ca
+Ca +Ca
+Ca
−Ca
−Ca
−Ca
−Ca
Quantum yieldA B
DC
Spectral shift
Wavelength
Fluorescence life time Förster resonance energy transfer
Time (ns)
Wavelength
Wavelength
Ligh
t int
ensi
tyLi
ght i
nten
sity
Ligh
t int
ensi
tyLi
ght i
nten
sity
Fig. 7 Major categories of fluorescence properties of Ca2þ indicators. (A) Change in dye absor-
bance and quantum yield generates a Ca2þ-sensitive change in the fluorescence intensity (e.g., Fluo-3/
4 Rhod-2, Oregon Green, and Fura-Red (inverse relationship)). (B) Spectral shift in the excitation
spectrum as a result of Ca2þ binding to an indicator allows ratiometric measurements (e.g., Fura-2/3/
4/6/FF). (C) Changes in fluorescence life time as a result of Ca2þ binding to an indicator; the
fluorescence decays exponentially within ns of the end of excitation. The rate of decay is Ca2þ
dependant; for example, the decay of Ca2þ-bound Fluo-3 fluorescence is faster than the decay of
unbound Fluo-3. (D) Change in Forster resonance energy transfer (FRET) efficiency as a result of
Ca2þ binding to either the acceptor or donor proteins; Ca2þ binding changes the distance between the
two linked fluorescent proteins. The distance between the donor and acceptor proteins determines the
degree of FRET; an increase in FRET efficiency causes a decrease in donor fluorescence and an
increase in acceptor fluorescence. Black lines indicate excitation spectra and gray lines indicate
emission spectra. Dotted lines represent the Ca2þ-free form of the dye, whereas solid lines represent
the Ca2þ-bound form of the dye.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 247
Fluo-3, Fluo-4, and Oregon Green all show dramatic increase in fluorescence
upon binding to Ca2þ, while Fura-Red fluorescence decreases upon Ca2þ binding.
The optical arrangement required for these dyes is the simplest, that is, single
excitation and emission wavelengths.
Spectral shift: In this case, Ca2þ binding causes a shift in either the excitation
or emission wavelengths. Fura-2 and related dyes show a shift in the excitation
spectrum of the dye with minimal changes in the emission (Fig. 7B). In contrast,
Indo-1 and related dyes show large shifts in the emission spectrum of the dye
with minimal changes in the excitation spectrum. This spectral shift is used to
allow ratiometric measurements. In the case of Fura-based dyes, while exciting
at �340 nm, a rise of Ca2þ will cause an increase in fluorescence, whereas while
exciting at 380 nm causes a decrease. The ratio of fluorescence at these two
wavelengths is a unique function of the [Ca2þ] and is independent of the
concentration of the dye. This is particularly useful when measuring cells that
move and thereby alter the amount of dye within the light path, or comparing
cell compartments with diVerent dye concentrations (e.g., nucleus versus the
cytosol). However, despite all these advantages, the Fura- and Indo-based dyes
are rarely used in confocal microscopy because excitation wavelengths are
short (<400 nm) and therefore not convenient for commonly available lasers.
The shortest wavelength routinely available in commercial systems is 405 nm,
which can be used to excite Fura-based dyes close to the 380 nm excitation
maximal. In this mode, Fura acts as an inverse indicator, a property that has
some value in single-photon confocal and 2P imaging (Ogden et al., 1995;
Wokosin et al., 2004).
Fluorescence lifetime: In this case, the decay of fluorescence at the end of a pulse
of excitation light will take a finite time (ns) to decay. The time course of decay is
aVected by the binding of Ca2þ (Fig. 7C). In the case of Fluo-3, the decay of
fluorescence or lifetime is shorter for the Ca2þ bound form (Sanders et al., 1995).
Since the lifetime is independent of the concentration of the dye, the relative
population of bound and free dye can easily be calculated from this technique.
However, the nontrivial analysis of emission decay required to obtain lifetime data
has minimized the use of this technique.
FRET eYciency: In this case, a donor molecule absorbs photons and can transfer
the associated energy to a close by acceptormolecule via a nonradiative process that
operates at distances less than the wavelength of light (up to 10 nm). The closely
adjacent molecule accepts the energy transfer (thus excites) and then emits light at a
distinct wavelength (Fig. 7D). The most popular pairs of donor and acceptor
molecules are cyan fluorescent proteins (donor) and yellow fluorescent proteins
(acceptor). The eYciency of the energy transfer depends on the proximity of the
two molecules; within 10 nm, the eYciency is high but this drops dramatically as
the distance between the two molecules increases (proportional to 1/(distance)6).
This mechanism can be used to detect Ca2þ binding to either construct since the
change in tertiary structure will alter FRET eYciency and therefore altered
FRET signal.
248 Godfrey Smith et al.
XIII. Use of Dyes for Single-Photon Confocal Microscopy
In theory, all the fluorescent Ca2þ indicators created for cell biology can be used
in confocal or 2P excitation modes, although in practice, only a limited number of
dyes are routinely used for a series of practical considerations. By far, the majority
of confocal applications use single-wavelength excitation and emission dyes. These
dyes have several advantages: (i) their excitation wavelength is �500 nm or longer
and therefore can use the emission from readily available Argon or Krypton-
Argon lasers, and (ii) the majority of dyes have a good dynamic range (see
below). But they also suVer from a series of disadvantages: (i) calibration is diYcult
because the signal will be a function of both the concentration of the dye (unknown
and variable) and the concentration of Ca2þ and (ii) dye signal may vary with time
due to loss of the dye from the volume or due to photobleaching. All dyes
photobleach, but some dyes are more susceptible than others; in particular,
Fluo-3 and Fluo-4 are amongst those that most rapidly bleach (Thomas et al.,
2000). Dyes that show significant spectral shifts can be used in a single wavelength
mode, for example, Fura-based dyes can be imaged in confocal microscopy by
using a 405 nm laser. This produces an inverse reporter since fluorescence
decreases as Ca2þ increases (Wokosin et al., 2004). Similar measurements using
Indo-based dyes are less easy because the narrow excitation spectrum means that
405 nm laser light only poorly excites the dye and Indo-based dyes appear to be
inherently more prone to photobleaching.
XIV. Use of Dyes for 2P Excitation Microscopy
The utility of a dye for imaging Ca2þ using 2P excitation does not follow directly
from the behavior of the dye in single-photon excitation. There is no guarantee
that 2P excitation will successfully excite the dye since the eYciency of two-photon
excitation appears to be relative to the fluorophore structure (Kim et al., 2008).
A number of Ca-sensitive indicators have been studied over a limited range of 2P
wavelengths (Xu et al., 1996), some dyes (e.g., Oregon Green) have a better 2P
cross-section than the more common single wavelength dyes (Fluo-3); none
showed the expected increase in cross-section as the excitation wavelength
approached 900 nm. In contrast, the 2P cross-section of Fura-based dyes appeared
readily excitable by wavelengths approximately to double the appropriate single-
photon wavelength (Wokosin et al., 2004). In particular, �800 nm light provides
an inverse Ca2þ-sensitive signal that corresponds to the excitation of this dye at
�400 nm, as advocated previously (Ogden et al., 1995). However, few studies to
date have explored the more commonly used dyes (Fluo-3/4 and Rhod-2) at the
longer wavelengths achievable currently with tunable Ti:Sapphire lasers (up to
1100 nm). As shown in Fig. 8, the Fluo-based dyes do not show the peak of
fluorescence anticipated from approximately double single-photon wavelengths
(�1000 nm). The small peaks in the excitation spectrum observed at 400–470 nm
1000
100
10
1000
100
10
1000 1P × 22P
1P × 2
2P
900
Wavelength (nm)
1000 1200
Excitation wavelength (nm)
800
800
1000900800700
700
Fluo-3
Rhod-2B
A
Flu
ores
cenc
e (a
.u.)
600
400
200
0
1000 1100 1200900800700
Flu
ores
cenc
e (a
.u.)
600
400
200
0
Fig. 8 Excitation spectra of Ca2þ bound forms of (A) Fluo-3 and (B) Rhod-2. Emitted fluorescence
was collected at 500–650 nm (Fluo-3) and 550–650 nm (Rhod-2). Black line represents the single-photon
(1P) excitation spectra measured on a Perkin–Elmer spectrophotometer (2 nm slit width); the 1P
wavelength has been scaled by a factor of 2 (�2) to generate comparable wavelengths to two-photon
(2P) spectra. Gray line represents the 2P excitation spectrum of an equivalent concentration of dye (10
mM); excitation light was provided by a Coherent Chameleon XP Ti:Sapphire laser attached to a Zeiss
510 upright laser scanning microscope. Laser power was altered at each wavelength to ensure equivalent
excitation power across the excitation wavelengths. Insets show spectra plotted on log scale.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 249
250 Godfrey Smith et al.
during single-photon excitation were also paralleled by similar spectra at
800–950 nm during 2P excitation, but longer wavelengths failed to produce the
increased fluorescence anticipated from the single-photon spectrum. The behavior
of the Rhod-dyes (related structurally to Fluo-dyes) has yet to be examined in
detail mainly because accessing wavelengths >1100 nm is technically diYcult.
The practical application of 2P excitation to excite Ca2þ-sensitive dyes is limited;
the majority of applications used �800 nm to excite either Fluo-3 or Rhod-2.
In doing so, these studies are using the higher powers available from lasers at
these wavelengths, rather than attempting to access the higher quantum yields that
may be present at longer wavelengths. Further work is clearly required to deter-
mine the optimal 2P conditions to excite these readily available dyes at 1000–
1100 nm, or alternatively design new dyes with fluorophore structures that are
more easily excited by the 2P approach (Kim et al., 2008).
XV. Is It Worth Converting the Intracellular FluorescenceSignal to [Ca2þ]?
Ca2þ-sensitive dyes are frequently used in conjunction with confocal microscopy
to simply indicate the timing or frequency of transient Ca2þ events. Under these
circumstances, it is not considered necessary to convert the fluorescence signal into
estimates of intracellular [Ca2þ]. This is avoided partly for experimental ease, since
conversion requires knowledge of the concentration of the dye and its aYnity for
Ca2þ, and partly because absolute Ca2þ concentrations are not considered vital to
the interpretation of the experiments. In a number of cases this may be justified,
but whenever amplitude or time course of a Ca2þ transient is considered an impor-
tant variable, then calibration becomes essential for the following several reasons:
1. It is important to distinguish changes in background Ca2þ from one experi-
mental scenario to the next since this determines the subsequent intracellular Ca2þ
buVer power. Any event that generates a Ca2þ transient, for example, Ca2þ influx
via plasmalemmal Ca2þ channels or Ca2þ release from an internal store, will
increase total intracellular Ca2þ by a specific amount, but the extent to which
this increases the free cytosolic Ca2þ concentration will depend on the cellular
buVer power for Ca2þ. This is illustrated in Fig. 9, where total cellular Ca2þ is
increased by a standard amount; the subsequent free Ca2þ transient amplitude can
be �40% larger simply because of small (�10%) changes in resting [Ca2þ]. There-fore, if the basal Ca2þ concentration changes, the interpretation of changes in
transient amplitude has to be made with caution.
2. Intracellular [Ca2þ] levels that almost saturate the indicator cannot be used to
examine moderate changes in the amplitude of the Ca2þ transient. In the absence
of information concerning the maximal Ca2þ signal, it is diYcult to know how close
the dye is to saturation and therefore how sensitive the signal is to changes in peak
Ca2þ level.
Time (s)
Tim
e (s
)
0
3.0
1.5
300(mM)
(mM)
150
T1
T2
Buffercurve
0.5
0
0.5
1.0
1.0
F2
F1
Free[Ca2+]
[Ca2+]free
[Ca2+]total
Total [Ca2+]
Fig. 9 Illustration of the conversion of increments in total cellular Ca2þ to free Ca2þ signal using the
cellular buffer power. Increase of the total cellular Ca2þ by equivalent amounts (T1 & T2) causes a rise
of free [Ca2þ] of F1 and F2 due to differences in the background [Ca2þ]. Cellular Ca2þ buffer is
illustrated by the relationship between total cellular Ca2þ and free Ca2þ. Note that while the amplitude
of the transient increase in free [Ca2þ] depends on the increase in total Ca2þ and the cellular buffer
power, the time course of the decrease will depend on the extent of activation of cellular Ca2þ pumps
and exchangers. Generally, the rate of these processes depends on the free [Ca2þ], therefore the decay ofthe Ca2þ transients of different amplitudes may differ substantially.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 251
XVI. Calibration of Single Wavelength Dyes
Calibration of single wavelength dyes is based on two main assumptions: firstly,
that the dye is at equilibrium with intracellular Ca2þ, that is, the kinetics of the
change of intracellular Ca2þ are slow compared to the rate constants for associa-
tion and dissociation. For the commonly used Fluo-type dyes at an intracellular
concentration of<100 mM, the half time for association and dissociation are of the
order of�1 ms and therefore for most circumstances the equilibration assumption
holds. The second assumption is that all the dye molecules sense the same amount
of light, that is, there is no filtering intrinsic to the biological preparation or by the
dye itself. Typically, the absorption coeYcient of Ca2þ indicators is of the order of
50,000 M�1 cm�1, therefore significant absorbance (>0.05) would occur in a cell
10 mm thick containing >100 mM dye. This consideration is important for single-
photon excitation and is one of the constraints that limit single-photon imaging to
thin (<50 mm) specimens.
252 Godfrey Smith et al.
With these two criteria satisfied, the Ca2þ concentration can be calculated from
the fluorescence from single wavelength dyes using the following equation:
Ca2þ� � ¼ Kd F � Fminð Þ= Fmax � Fð Þ
where F is the fluorescence signal from the cell/tissue; Kd the dissociation constant
of Ca2þ for the indicator (units M); Fmin the minimum fluorescence achieved when
the dye is essentially Ca2þ free, which practically can be approximated by exposing
the dye to a [Ca2þ], that is, 0.01Kd of the dye; and Fmax is the fluorescence achieved
when the dye is completely Ca2þ bound, which practically can be achieved with
[Ca2þ] of 100Kd of the dye. Measurements of these constants with some degree of
precision inside a cell are, however, diYcult. The dissociation constant can be
measured outside the cell in solutions approximating the intracellular mileau, but it
has been a frequent observation that the value of the Kd is altered by the intracel-
lular environment in a way that is diYcult to mimic by solution chemistry, for
example, mimicking intracellular viscosity and the range of negatively charged
intracellular proteins (Poenie, 1990). Therefore, the best practice is to measure the
dissociation constant within the cell type of interest. The easiest way to achieve this
is by using a glass microelectrode to gain access to the intracellular space. The use
of a series of solutions with a high concentration of Ca2þ buVer (EGTA or
BAPTA) with specific [Ca2þ] can be used to make a series of single cell measure-
ments to allow estimation of Kd. However, it is important to note that this
technique cannot be applied to multicellular preparations where multiple cells in
a tissue are diVerentially loaded with the dye.
XVII. Estimation of Fmax Values
This should be measured on a cell-to-cell basis even within multicellular prepara-
tions and involves exposing the inside of the cell to�50 mMor higher Ca2þ, depend-ing on the aYnity of the dye for Ca2þ. These levels are generally toxic to cells, but if
tolerated for a short time (1–2 s), this may be suYcient to estimate Fmax. These
intracellular [Ca2þ] levels can be achieved rapidly within single cells by perfusion
with a Ca2þ ionophore and raised extracellular Ca2þ (Loughrey et al., 2003).
A second ingenious method used in single voltage clamp experiments is to use an
amphotericin-containing patch pipette that facilitates monovalent cation exchange
across the membrane within the patch and therefore allows low resistance access to
the cell. At the endof the experiment, themembrane is ruptured under the patchusing
a rapid pressure step and the resultant influx of Ca2þ from the patch pipette generates
a rapid rise of intracellular [Ca2þ] that can be used to assess Fmax (Diaz et al., 2001).
A simpler but less reliablemethod is to simply use themicroelectrode to penetrate the
cell and allow extracellular Ca2þ influx in order to record Fmax, but generally Ca2þ
influx occurs in parallel with a rapid loss of intracellular dye so the signals would have
to be interpreted with caution.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 253
Depending on the tissue, there are alternative approaches to estimate Fmax; in
some nerve cells, rapid frequent stimulation of cells can generate intracellular Ca2þ
levels that approach saturation of the dye (Maravall et al., 2000), thus allowing
Fmax values to be estimated.
XVIII. Estimation of Fmin or the Dynamic Range of the Dye
Estimation of Fmin or the dynamic range of the dye is more diYcult. The ratio of
Fmax/Fmin measured outside the cell cannot be assumed to apply inside; estimates
suggesting that values of �70–80% of those seen in free solutions are common
(Poenie, 1990). Again, an averaged value can be obtained using patch pipettes as a
means of establishing buVered [Ca2þ] inside cells. Alternatively, the Fmin in each
experiment can be estimated if the intracellular Ca2þ can be lowered to 1–10 nM
(for Fluo-3) by decreasing extracellular Ca2þ, but this assumes that intracellular
Ca2þ can be readily manipulated by changes in the extracellular environment
which is not always the case for every cell type. The eVect of under- or overestima-
tion of the dynamic range of the indicator is shown in Fig. 10C. Based on the
reported dynamic range of Fluo-based dyes (�100�), large under/overestimates of
the dynamic range (by up to 30%) cause only small errors in Ca2þ estimation and
only in the lower range of [Ca2þ] values relative to the Kd of the dye.
XIX. Consequence of Errors in Estimation of Intrinsic andDye Fluorescence
Prior to conversion of the indicator-based fluorescence to [Ca2þ], the back-
ground or intrinsic fluorescence of the cell/tissue has to be subtracted from the
signal. All cells have an intrinsic fluorescence mainly due to the metabolites beta
nicotinamide adenine dinucleotide (NADH) and flavin adenine dinucleotide
(FAD); their excitation wavelengths are 350–500 nm and emission wavelengths
�450–600 nm. The relative fluorescence of these two metabolites depends on the
metabolic state of the cell/tissue and degree of photobleaching. Thus, intrinsic
cellular fluorescence is significant and variable. The most advisable approach is to
use a dye with a significant basal fluorescence that is many times (>10�) that of the
intrinsic value. This cannot always be achieved; the Fluo-based and Rhodamine-
based dyes are by far the most popular dye groups used in confocal and 2P excitation
microscopy. Their main attraction is a large dynamic range as a result of a low
fluorescence signal from the Ca2þ free form. In this situation, Fmin values are fre-
quently comparable to that of the intrinsic fluorescence of the cell and therefore it is
important to quantify either by parallel measurements on nonloaded tissue or from a
single cell prior to the introduction of the dye. Error in estimation of background
fluorescence (which can be up to 100%) has dramatic eVects on the calculation of
Error in background estimationA B
Error in absolute fluorescence
Kd= 0.4mM Kd= 0.4mM
%E
rror
in [C
a2+] m
easu
rem
ent
%E
rror
in [C
a2+] m
easu
rem
ent
Kd= 0.8mM Kd= 0.8mM
100
−100%
−30%
−10%
+10%
+30%+100%
−50%
+50%
80
Rel
ativ
e flu
ores
cenc
e
60
40
20
0.1
[Ca2+]/Kd
1 10
40
0.1
[Ca2+]/Kd
1 100
100
80
Rel
ativ
e flu
ores
cenc
e
60
40
20
0
20
0
−20
40
20
0
−20
−40
C Error in dynamic range
Kd= 0.4mM
%E
rror
in [C
a2+] m
easu
rem
ent
Kd= 0.8mM
−30%
−10%
+10%
+30%
0.1 1 10
[Ca2+]/Kd
100 4
80
Rel
ativ
e flu
ores
cenc
e
60
40
20
0
2
−2
−4
0
Fig. 10 [Ca2þ] calibration errors. (A) Error due to changes in background fluorescence plotted
against varying [Ca2þ] values normalized by the indicator Kd (0.4 mM, Fluo-3 and 0.8 mM, Fluo5F).
Dynamic range of the dye was set at 100 (maximum attributable to Fluo-3). A typical cellular Ca2þ
concentration range is highlighted by gray boxes (100 nM to 1 mM) for each of the twoKd values. The left
axis (thick line) highlights the relative fluorescence versus [Ca2þ]. The right axis (thin lines) represents the%error in [Ca2þ] due tovariations in the intracellular background.Two levels of backgroundfluorescencewere considered: (i) background fluorescence is equal in magnitude to Fmin; errors of �50% and�100%
background were considered, and (ii) background fluorescence is equal to 5� Fmin; errors of �10% and
�20% of background fluorescence. These two combinations of background and errors superimpose
exactly (i.e., �50% superimposes on �10% and �100% superimposes on �20%) to produce the 4 error
lines shown. (B) Error in absolute fluorescence levels; errors of�10% and�30% are shown. (C) Error in
dynamic range of Fluo- and Rhod-based dyes; errors of�10% and �30% are shown.
254 Godfrey Smith et al.
intracellular [Ca2þ], particularly at either end of the sensitive range of the indicator, asshown in Fig. 10A. If the dye has a Ca2þ aYnity midway between the extremes of
intracellular Ca2þ, then the error can be small and approximately constant. But if the
dye has a lowerCa2þ aYnity, whichmay be desirable to resolve changes in peakCa2þ,then errors associated with the minimum cellular Ca2þ levels can be large.
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 255
These errors can be compounded by errors in estimates of fluorescence or the
variability of signals from one cell to the next. Fig. 10B shows the errors in Ca2þ
based on simple errors in fluorescence changes. The graph illustrates the risk
inherent in using dyes with a relatively high aYnity relative to the physiological
signal. Small errors in the range of fluorescence signals translate to large errors of
intracellular Ca2þ such that the ability to discriminate changes in maximum
physiological response is severely impaired. This can be significantly improved
by using lower aYnity dyes, but at the cost of poor resolution of minimum or
background intracellular [Ca2þ].
XX. Multimodal and Multiple Fluorophore Confocal andMultiphoton Microscopy
Although Ca2þ is an important signaling molecule in a variety of cell types, it by
no means operates alone. Rather, Ca2þ both temporally and spatially interacts
with many other properties and processes in the cell that only in concert orches-
trate cellular function. Thus, some of these processes are dictated by Ca2þ, butsome or not. A good example of this interplay is excitation–contraction coupling
in muscle cells, in which the action potential depolarizes the plasma membrane of
the cell, which causes a small influx of Ca2þ through the membrane. This inward
Ca2þ current stimulates the ryanodine receptor to release bulk Ca2þ from the
sarcoplasmic reticulum, which upon binding to the myofilaments induces the
actin–myosin interaction and the subsequent cellular contraction (Bers, 2002).
The cellular contraction may be imaged by simple black-and-white contrast
edge-detection microscopy, but this is not the case for the intracellular Ca2þ and
membrane potential characteristics that both require more sophisticated methods
such as fluorescence microscopy. Thus, simultaneous imaging with the use of
multiple fluorophores present at the same time in the specimen or combinations
of diVerent imaging modalities in some sense is required for capturing complex
information.
Thus, loading or injecting the specimen with multiple fluorophores allows for
simultaneous recording of diVerent signals, or if simultaneous recordings are not
technically possible, diVerent signals may be recorded sequentially without having
tomanipulate, move, or in any other way perturb the specimen between recordings.
In the latter case, only the optical pathways of the microscope would be altered
between recordings, whereas the specimen would not, since it would already be
loaded with diVerent fluorophores. The use of multiple fluorophores require either
the ability to direct separate emission wavelength bands onto diVerent light detec-tors, or to spectrally separate diVerent fluorophores by diVerent excitation wave-
lengths. Depending on the hardware, both confocal and multiphoton microscopes
can fulfill these requirements and therefore allow for measurements with multiple
fluorophores. Such experiments can be done by simultaneously loading the
256 Godfrey Smith et al.
specimen with a Ca2þ-sensitive fluorophore and a fluorophore that is sensitive for
another characteristic of the cell, which may also be Ca2þ in a diVerent compart-
ment of the cell with diVerent dynamics or a diVerent concentration range, or a
second fluorophore that may be sensitive to the plasma membrane voltage in an
excitable cell (also called a potentiometric dye). Measurements of Ca2þ and mem-
brane voltage (resting membrane potentials and action potentials) may then be
conducted either simultaneously by exciting both fluorophores at the same time and
capture spectrally diVerent fluorescence emission signals, or sequentially by exciting
each fluorophore separately, that is, one after the other, under otherwise similar
experimental conditions. The latter approach would assume that the experimental
conditions remain the same. Several factors may necessitate this, such as an inabili-
ty to diVerentiate between diVerent emission signals, or an inability to excite more
than one fluorophore at any given time, for example, if the excitation spectra do not
overlap and only one excitation wavelength may be delivered at one time.
The advantage of using multiple fluorophores either simultaneously or sequen-
tially is to increase the information content of the imaging, especially how diVerentprocesses relate to each other spatially and temporally. However, several issues
may limit the applicability of such measurements. Introducing a fluorophore to the
specimen may also change the dynamics of the cellular parameter of interest,
especially in live specimens that rely on stable and constant intra- and extracellular
environments. For instance, most Ca2þ indicators are also Ca2þ chelators that
buVer free Ca2þ, and most fluorophores or the medium they are delivered in may
change biochemical and biophysical properties of the intracellular environment.
This may be accentuated by simultaneous loading with several dyes. DiVerent dyesmay also quench, sequester, or in other ways inhibit each other. Finally, excitation
in itself may cause changes or damage to the specimen, and although this to some
degree is unavoidable, the degree of change or damage may be diVerent or evenaccentuated during sequential recordings.
The single-photon excitation and emission spectra of multiple Ca2þ- and
voltage-sensitive fluorescent dyes are well known. Clearly, some dyes have over-
lapping excitation or emission peaks, or present with broad excitation or emission
spectra such that even if the peaks are separated from one another, the tails of the
spectra still overlap considerably. Overlapping excitation spectra means that
diVerent dyes may be excited simultaneously, but overlapping emission spectra
may result in severely reduced signal specificity, and therefore, certain combina-
tions of fluorescent dyes may be less applicable, such as the potentiometric Di-4-
ANEPPS and Di-8-ANEPPS dyes, and the Ca2þ-sensitive Fluo-3 dye, all exten-
sively used by numerous laboratories for single-fluorophore purposes. All of these
dyes have single-photon excitation peaks at �480–500 nm and emission peaks
at 520–610 nm, respectively, with especially Di-4-ANEPPS and Di-8-ANEPPS
having very broad emission spectra that peak at �610 nm, but that considerably
overlap with the Fluo-3 emission spectrum, even though the latter has its peak
at �525 nm and therefore numerically diVers from Di-4-ANEPPS and
Di-8-ANEPPS and is more narrow (Fig. 11A). This problem may to some degree
800700
Wavelength (nm)
Rhod-2 and RH-237
600500
600
B
Em
issi
on in
tens
ity (
a.u.
)
500
400
300
200
100
0
800
A
800
700
700
Wavelength (nm)
600
600
500
500
Fluo-3 and Di-4-ANEPPS
400
300
200
100Em
issi
on in
tens
ity (
a.u.
)
0
C
800700Wavelength (nm)
600500
Fluo-3 and RH-237
Em
issi
on in
tens
ity (
a.u.
)
700
600
500
400
300
200
100
0
Fig. 11 Single-photon emission spectra of voltage-sensitive Di-4-ANEPPS and RH-237, and Ca2þ-sensitive Fluo-3 and Rhod-2 dyes. The following emission spectra were obtained by spectrophotometry
after simultaneously loading cardiac muscle cells in a high Ca2þ solution (60 mM) with Ca2þ- andvoltage-sensitive dyes and exciting at 488 nm. The presence of intact excitable cells in a Ca2þ-richenvironment provides substrate for both Ca2þ- and voltage-sensitive dyes. (A) Fluo-3 (Ca2þ; first peak)and Di-4-ANEPPS (voltage; second peak). (B) Rhod-2 (Ca2þ; first peak) and RH-237 (voltage; second
peak). (C) Fluo-3 (Ca2þ; first peak) and RH-237 (voltage; second peak).
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 257
be avoided as the Ca2þ and voltage signals are spatially separated between intra-
cellular and membrane compartments of the cell, though in muscle cells this may
turn out to be diYcult because of the dense network of plasma membrane trans-
verse tubules that penetrate into the interior of the cell. The same overlap problem
exists with the voltage-sensitive RH-237 and the Ca2þ-sensitive Rhod-2 dyes, with
emission peaks occurring at �580 and �660 nm, respectively, but with especially
the RH-237 emission spectrum being very broad (Fig. 11B). In contrast, Fluo-3
and RH-237 are more distinctly separated from one another. Both Fluo-3 and
RH-237 dyes may be excited by the same single-photon excitation wavelength at
258 Godfrey Smith et al.
�500 nm (though RH-237 would not be optimally excited by this wavelength), but
have emission spectra that may be spectrally diVerentiated, as Fluo-3 has a narrowemission spectrum that peaks at �525 nm, whereas RH-237 peaks at �660 nm
with a broad spectrum (Fig. 11C). Thus, this combination appears attractive as it
diVerentiates the Ca2þ and membrane potential signals, for which it has also been
used successfully (Fast and Ideker, 2000).
However, several problems arise when transferring from single-photon to 2P
excitation, although several of the voltage-sensitive dyes present with consistent
and reproducible 2P excitation spectra that very much resemble the double single-
photon excitation spectra and may thus be confidently used for meaningful multi-
photon imaging. In contrast, the 2P behavior of many of the Ca2þ-sensitive dyes issomewhat diYcult to interpret (see previous discussion and Fig. 8), although the
ratiometric Fura dyes may be 2P excited in order to provide a meaningful Ca2þ
signal that also captures transient changes over a millisecond scale with high
fidelity (Wokosin et al., 2004). This may be because the Fura dyes have a single-
photon excitation spectrum in the UV range (340–380 nm), and therefore the 2P
excitation spectrum, which approximately is double the single-photon spectrum,
occurs at �800 nm wavelengths, in which the 2P laser power outputs are not
limited. Moreover, this study indicated that several of the Fura dyes, in particular
Fura-4F, may work well when excited with a single IR wavelength, despite their
use as ratiometric single-photon dyes, as judged by the dynamic ranges and SNR
obtained during 2P excitation microscopy in single cardiac muscle cells during
diVerent Ca2þ conditions. In contrast, the Fluo- and Rhod-based Ca2þ-sensitivedyes, all with single-photon excitation peaks at �500–550 nm, present with 2P
excitation spectra that are not immediately predicted by the doubled single-photon
spectra (Fig. 8). In these cases, the 2P excitation spectra are at least partly broken
up and appear blueshifted compared to the doubled single-photon spectra.
Although doubling the single-photon excitation spectrum is often a good predictor
for the 2P excitation spectrum, deviations from this do occur, although these
deviations may neither be systematic nor well understood (Xu et al., 1996; Zipfel
et al., 2003). Alongside this, a reoccurring problem is that the available Ti:Sapphire
pulsed 2P lasers are power-limited at the long wavelengths of 1000–1100 nm that
would correspond to the doubled single-photon excitation spectra of Fluo-3 and
Rhod-2. Because not all fluorophores are easily transferable from single-photon
excitation, for which they were developed, to 2P excitation, this therefore has made
it problematic to use multiple fluorophores simultaneously during 2P excitation
microscopy, and the issue has not yet been fully resolved.
A diVerent approach to capture more complex information has been to combine
several multimodal microscopy techniques inways that also encompass confocal and
multiphoton systems, but also this comes with both advantages and disadvantages.
For instance, diVerentmodes of contrast used on the same specimenmay increase the
information extracted from the images and reduce artifacts. Multimodal microscopy
may also allow for a wider repertoire of fluorophores. However, if confocal and
multiphoton imaging are combined, it requires descanning and insertion of a
9. Confocal and Multiphoton Imaging of Intracellular Ca2þ 259
confocal aperture with a pinhole into the light pathway, which may reduce the
fluorescence capture after 2P excitation and thus lead to a loss of signal, though a
confocal aperturemay also be set up to increase the spatial resolution ofmultiphoton
images, by restricting the PSF tails. Because of these limitations, the reality is often
that it is diYcult, though not impossible, to achieve optimal performance from each
individual mode when several modes are combined. Nonetheless, the advantages of
simultaneous or near-simultaneous light capture by diVerent modes of microscopy
may, under the right circumstances, far outweigh the disadvantages.
Examples include combinations of confocal or multiphoton with epifluorescence
or diVerential interference contrast microscopes to capture light emission restricted
to the focal plane as well as capturing a widefield view, either simultaneously or
sequentially without having to reorient or replace the specimen. Other options also
include setting up a microscope system that combines confocal and 2P excitation
imaging modes, or 2P excitation and second-harmonic generation (SHG) imaging.
Although these applications tend to serve narrow and specific purposes, they may
allow for imaging of local versus global Ca2þ signaling, or Ca2þ signaling in
combination with for example, metabolic parameters by using 2P excitation to
excite metabolites such as NADH and FAD, or collagen that in particular con-
tributes to the SHG signal (Masters, 2006). A final example of multimodal micros-
copy techniques that may successfully be combined includes the combination of
FRET and FLIM imaging to quantify FRET between two fluorophores, as in the
case of the Ca2þ-sensitive cameleon described above. These examples are not
exhaustive, but serve to illustrate the potential of combining diVerent fluorophoresor microscopy modalities in order to gain information of a more detailed nature.
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CHAPTER 10
METHODS IN CELL BIOLCopyright 2010, Elsevier Inc.
The Use of Aequorins to Record andVisualize Ca2þ Dynamics: From SubcellularMicrodomains to Whole Organisms
Sarah E. Webb,* Kelly L. Rogers,† Eric Karplus,‡ andAndrew L. Miller**Biochemistry and Cell Biology Section and State Key Laboratory of Molecular NeuroscienceDivision of Life ScienceHKUST, Clear Water BayKowloon, Hong Kong, PR China
†The Walter and Eliza Hall Institute of Medical ResearchParkville, Australia
‡Science Wares Inc.FalmouthMassachusetts, USA
A
OGY,All rig
bstract
VOL. 99 0091hts reserved. 263 DOI: 10.1016/S0091
-679X-679X
I. In
troduction II. E xpression of Apoaequorin, GFP-Apoaequorin, and Other Apoaequorin-BasedSpectral Variants in Cells, Tissues, and Whole Organisms
A. Expression of Apoaequorin B. Expression of GFP-Apoaequorin C. Summary of Section IIIII. In
troducing Coelenterazines into Cells, Tissues and Embryos IV. T echniques for Detecting Aequorin Luminescence V. C onclusionsR
eferencesAbstract
In this chapter, we describe the practical aspects of measuring [Ca2þ]transients that are generated in a particular cytoplasmic domain, or within a
specific organelle or its periorganellar environment, using bioluminescent,
/10 $35.00(10)99010-9
264 Sarah E. Webb et al.
genetically encoded and targeted Ca2þ reporters, especially those based on
apoaequorin. We also list examples of the organisms, tissues, and cells that
have been transfected with apoaequorin or an apoaequorin-BRET complex, as
well as of the organelles and subcellular domains that have been specifically
targeted with these bioluminescent Ca2þ reporters. In addition, we summarize
the various techniques used to load the apoaequorin cofactor, coelenterazine,
and its analogs into cells, tissues, and intact organisms, and we describe recent
advances in the detection and imaging technologies that are currently being
used to measure and visualize the luminescence generated by the aequorin-
Ca2þ reaction within these various cytoplasmic domains and subcellular
compartments.
I. Introduction
One of the most significant recent developments in the Ca2þ signaling field has
been the general acceptance of the wide-spread heterogeneity of Ca2þ activity
within individual cells; not only at rest, but also most importantly, during stimula-
tion (Berridge, 2009; Rizzuto and Pozzan, 2006; Rutter et al., 2006; Whitaker,
2008). This has led to the concept of dynamic subcellular ‘‘Ca2þ microdomains.’’
As suggested by Rizzuto and Pozzan (2006), this term (especially with regard to its
spatial dimensions) has several diVerent meanings depending on one’s area of
interest. In this chapter, however, like Rizzuto and Pozzan, we use the term in a
general way to describe Ca2þ dynamics that do not involve the entire cell cyto-
plasm, but that remain localized to a specific cytoplasmic domain, or occur within
a particular organelle or its periorganellar environment. Thus, one of the current
challenges researchers are facing in the field of Ca2þ imaging is that of resolving
changes in [Ca2þ] within, and between, various subcellular microdomains.
An eVective strategy to address this challenge that is common to both fluorescence-
and luminescence-based imaging techniques is to exclusively visualize Ca2þ
dynamics in specific microdomains using genetically encoded and targeted Ca2þ
reporters (GET-CRs). These come in two general forms, fluorescent GET-CRs
and bioluminescent GET-CRs, respectively. At the other end of the size spectrum
is the exciting prospect of imaging Ca2þ signals derived from GET-CRs within
freely moving, large, organisms, for example, adult mice (Rogers et al., 2007).
This presents a diVerent set of technical challenges to researchers in the Ca2þ
imaging field.
Fluorescent GET-CRs include the camgaroos (Baird et al., 1999; Griesbeck
et al., 2001), G-CaMPs (Nakai et al., 2001; Ohkura et al., 2005), pericams (Nagai
et al., 2001), case-sensors (Souslova et al., 2007), grafted EF-hands (Zou et al.,
2007), and cameleon-types (Miyawaki et al., 1997; Ishii et al., 2006; Tsuruwaka
et al., 2007; and reviewed by Zorov et al., 2004; McCombs and Palmer, 2008).
Bioluminescent GET-CRs include single protein entities such as aequorin
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 265
(Cheung et al., 2006; Torrecilla et al., 2000), obelin (Stepanyuk et al., 2005),
mitrocomin (Inouye and Sahara, 2009), clytin (Inouye, 2008), and photina
(Bovolenta et al., 2007). In addition, there is a growing number of aequorin-
derived bioluminescence resonance energy transfer (BRET)-based complexes
such as the GFP-aequorins (Ashworth and Brennan, 2005; Baubet et al., 2000;
Martin et al., 2007 ; Rogers et al., 2005, 2007), as well as other wavelength-shifted
variants (Gorokhovatsky et al., 2004). To date, apoaequorin alone, or apoae-
quorin in tandem with another BRET protein, has been genetically expressed in a
diverse range of diVerent species; either in the whole organism or in specific
tissues within an intact organism (See Table I and Fig. 1). For example, apoae-
quorin has been ubiquitously expressed in whole zebrafish embryos (Cheung
et al., 2006) or specifically targeted to the Malpighian tubules in Drosophila
(Rosay et al., 1997), while the BRET complexes GFP-apoaequorin and YFP-
apoaequorin have been specifically targeted to neuronal cell subsets of Drosophi-
la (Martin et al., 2007), and the endodermis and pericycle of Arabidopsis roots
(Kiegle et al., 2000), respectively. Furthermore, apoaequorin or apoaequorin-
BRET complexes have been expressed either ubiquitously in the cytosol of cells in
culture or, using specific targeting sequences, in distinct organelles of cells in
culture (see Table II, and a recent review by Gerasimenko and Tepikin, 2005).
Specific organelles targeted include: the ER (Montero et al., 1997), mitochondria
(Rizzuto et al., 1992), the Golgi apparatus (Pinton et al., 1998), the nucleus (Brini
et al., 1993, 1994), gap junctions (George et al., 1998), subplasma membrane
domains (Marsault et al., 1997; Nakahashi et al., 1997), secretory vesicles
(Mitchell et al., 2001), and the outer mantle of secretory granules (Pouli et al.,
1998), to name but a few examples.
In this chapter, we focus on describing the practical uses of bioluminescent GET-
CRs, especially those based on apoaequorin. In addition, we provide the reader (in
table form) with a review of the literature to date listing representative examples of
whole organisms, tissues, and cells that have been transfected with apoaequorin or
an apoaequorin-BRET complex, as well as a list of organelles and subcellular
domains that have been specifically targeted (see Tables I and II). We also summa-
rize diVerent strategies used for loading various derivatives of the apoaequorin
cofactor, coelenterazine (Shimomura et al., 1989) into cells, tissues, and intact
organisms (summarized in Table III). Furthermore, we describe recent advances in
detection and imaging technologies used to measure and visualize light generated by
the aequorin-Ca2þ luminescent reaction within cells, tissues, and intact organisms
(summarized in Table IV and illustrated in Figs. 2 and 3). Our hope is that this
chapter will provide a starting point for researchers wishing to use GET-CRs to
measure or visualize Ca2þ dynamics from cells, tissues, or intact organisms. Fur-
thermore, the references provided in Tables I–IV should lead them to more detailed
information regarding a biological system and/or experimental setup that will com-
plement their own research interest. For loading holoaequorin into cells and embry-
os, we refer readers to the practical methodologies described inMiller et al. (1994), as
Table IExamples where apoaequorin or apoaequorin-BRET complexes have been targeted to a diverse range of diVerent species
Kingdom Class Species Apoaequorin or apoaequorin-BRET targeted to: References
Animalia Mammals Mus musculus (mouse) Whole organism Yamano et al. (2007)
Mitochondrial matrixa Rogers et al. (2007)
Fish Danio rerio (zebrafish) Whole organismb Cheung et al. (2006)
Trunk musculature Cheung (2009)
Amphibians Xenopus laevis (African clawed frog) Plasma membrane of oocytes Daguzan et al. (1995)
Insects Drosophila melanogaster (Fruit fly) Malpighian tubules—diVerent
cellular components
Rosay et al. (1997)
Mushroom bodies and antennal lobesa Martin et al. (2007)
Plantae Dicots Nicotiana plumbaginifolia (Tobacco) Whole organism Knight et al. (1991a)
Arabidopsis thaliana Whole organism Knight et al. (1995, 1996),
Sedbrook et al. (1996)
Guard cells Dodd et al. (2006)
Endodermis and pericycle of rootsc Kiegle et al. (2000)
Solanum tuberosum (Potato) Whole organism Fisahn et al. (2004)
Monocots Triticum aestivum (Winter wheat) Whole organism Nagel-Volkmann et al. (2009)
Moss Physcomitrella patens Whole organism Russell et al. (1996)
Fungi Funguses Phyllosticta ampelicida Whole organism Shaw et al. (2001)
Neurospora cassa Whole organism Nelson et al. (2004)
Aspergillus awamori Whole organism Nelson et al. (2004)
Aspergillus niger Whole organism Nelson et al. (2004),
Bencina et al. (2005)
Yeast Saccharomyces cerevisiae Whole organism Batiza et al. (1996)
Schizosaccharomyces pombe Whole organism Deng et al. (2006)
Protista Amoebozoa Dictyostelium discoideum (Slime mold) Whole organism Cubitt et al. (1995)
Diatoms Phaeodactylum tricornutum Whole organism Falciatore et al. (2000)
Monera Bacteria Escherichia coli Whole organism Knight et al. (1991b)
Bacillus subtilis Whole organism Herbaud et al. (1998)
Streptococcus pneumoniae Whole organism Chapuy-Regaud et al. (2001)
Blue-green algae Anabaena strain sp. PCC7120 Whole organism Torrecilla et al. (2000)
aGFP-aequorin constructs were used.bTransient transfection of apoaequorin mRNA was used.cYFP-aequorin constructs were used.
0 s 2 s 4 s 6 s
8 s 10 s 12 s 14 s
16 sPho
tons
/pix
el
18 s 20 s 22 s
0.20
0.16
0.12
0.08
0.04
0
A
1–2 3–4 5–6 7–8 >8
0 min
Photons/pixel
3 min 6 min 9 minB
0 s 15 s 12
1
30 sOK107
Pho
tons
/pix
el
C
0 16 32 64 128 225Photons
100 ppb 300 ppb 500 ppb 750 ppbAir
D
Fig. 1 Examples of the spatial patterns of Ca2þ signals generated by whole organisms, or by specific
tissues or subcellular organelles within intact organisms, where aequorin or GFP-aequorin were
genetically expressed. (A) Newborn mice stably expressing GFP-aequorin targeted to the mitochondrial
matrix were injected intraperitoneally with native coelenterazine and bioluminescence activity was
recorded with the animals un-restrained and freely moving. These are consecutive video images on to
which have been superimposed the corresponding bioluminescence images. Each panel represents 2 s of
accumulated light. Scale bar is 5 mm. Reproduced with permission, fromRogers et al. (2007). (B) An 18-
somite stage (i.e., �18 h postfertilization (hpf)) zebrafish embryo that was injected with apoaequorin
mRNA at the one-cell stage to transiently express apoaequorin throughout the whole embryo, and then
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 267
268 Sarah E. Webb et al.
not much has changed with respect to these particular techniques since Volume 40 of
the Methods in Cell Biology series was published.
II. Expression of Apoaequorin, GFP-Apoaequorin, and OtherApoaequorin-Based Spectral Variants in Cells, Tissues, andWhole Organisms
Microinjected holoaequorin has been used since the late 1960s for monitoring
changes in [Ca2þ]i in diVerent cells and tissues. The earliest reports describe the use
of holoaequorin to detect Ca2þ transients in muscle and nerve cells (Baker et al.,
1971; Ridgway and Ashley, 1967) as well as during activation in medaka eggs
(Ridgway et al., 1977). This approach is only practical, however, for introducing
aequorin into giant cells and large embryos, which are easy to microinject. The
more recent development, from the mid 1980s to early 1990s, of genetic engineering
techniques to introduce and express apoaequorin (the protein moiety of aequorin)
cDNA in cells, tissues, and whole organisms (Inouye et al., 1989; Knight et al.,
1991a,b; Nakajima-Shimada et al., 1991; Prasher et al., 1985; Saran et al., 1994), as
well as to target apoaequorin to specific organelles within cells (Brini et al., 1993;
Rizzuto et al., 1992), has paved the way for aequorin to be used as the Ca2þ
reporter of choice in many more biological systems today, from cells in culture to
complex multicellular organisms.
GFP-aequorin was developed approximately 10 years ago in order to improve
the stability and light emission properties of aequorin for single-cell imaging
(Baubet et al., 2000). Based on the naturally occurring phenomenon of BRET,
GFP-aequorin emits a red-shifted light emission (l¼509 nm) relative to that of
aequorin alone (l¼470 nm) in the presence of elevated free Ca2þ ion concentra-
tions. GFP-aequorin has a number of advantages over aequorin for monitoring
changes in cellular Ca2þ concentrations, including increased stability and total
light output. Furthermore, the expression level and distribution of the GFP reflects
the expression level and distribution of apoaequorin; thus, the expression of
apoaequorin can be directly visualized in living cells or tissues. Although the
incubated with f-coelenterazine starting at the 64-cell stage to reconstitute aequorin. Each panel
represents 120 s of accumulated light and consecutive panels are stepped at 60-s intervals. Scale bar is
200 mm. (C) D. melanogaster (P[GAL4] OK107 line) stably expressing GFP-aequorin in the mushroom
bodies. Exposed fly brains were incubated for >1 h at room temperature with native coelenterazine,
prior to imaging. The first panel shows the whole brain and the localization of GFP in the mushroom
bodies. The following panels show consecutive bioluminescent images, each panel representing 15 s of
accumulated light, following treatment with 70 mM KCl to induce Kþ-depolarization. Scale bar is
100 mm. Reproduced with permission, from Martin et al. (2007). (D) Nine-day old seedlings of
Arabidopsis thaliana (ecotype RLD1) that constitutively express apoaequorin were incubated in the
dark overnight in coelenterazine solution. These images show the total Ca2þ-dependent biolumines-
cence recorded from seedlings exposed to air or to diVerent concentrations of ozone for 1 h. Scale bar is
5 mm. #John Wiley and Sons Ltd. Reproduced with permission, from Evans et al. (2005).
Table IIExamples where apoaequorin or apoaequorin-BRET complexes have been targeted to the cytosol of cells in culture or todistinct intracellular organelles either in cells in culture or intact organisms
Species (common
name
and/or scientific
name) Cell type
Cytosolic expression of
apoaequorin or
apoaequorin-BRET Organelle(s) targeted Comments References
Kingdom: Animalia
Human (Homo
sapiens)
ECV304 (umbilical
vein endothelial cells)
✓ Mitochondria Transient expression Lawrie et al. (1996)
Diploid fibroblasts � (used indo 1-AM
and fura 2-AM
Mitochondria 1� cell cultures(Transient
expression)
Padua et al. (1998)
HEK-293 cells (embry-
onic kidney cell line)
✓ – Some transient
and some stable
expression
Sheu et al. (1993)
✓ – Stable expression Button and
Brownstein (1993)
✓ Nucleus HSV gene
transfer vector
Chamero et al. (2002)
✓ ER lumen,
mitochondrial
matrix
Transient expression Brini et al. (2005)
HEK 293T cells ✓ Nucleus,
mitochondria, ER
Expression of GA
and RA via HSV
gene transfer
vector
Manjarres et al. (2008)
HeLa (immortalized
epithelial cell line)
✓ Nucleus Stable expression Brini et al. (1993)
� (used Fura-2) ER lumen Transient and stable
expression
Montero et al. (1995)
� (used fura 2) ER 1� cell cultures(HSV gene
transfer vector)
Alonso et al. (1998)
✓ Mitochondria,
Golgi, ER
Transient expression Pinton et al. (2000)
� (used indo-1) Golgi, ER Stable expression Missiaen et al. (2004)
Normal adult and
Hailey-Hailey disease
keratinocytes
� (used proton-induced
X-ray emission)
Golgi 1� cell cultures(Transient
expression)
Behne et al. (2003)
Jurkat cells (immorta-
lized T-lymphocytes)
� (used indo PE3 (AM)) ER Transient expression Narayanan et al. (2003)
(continues)
Table II (continued )
Species (common
name
and/or scientific
name) Cell type
Cytosolic expression of
apoaequorin or
apoaequorin-BRET Organelle(s) targeted Comments References
African green
monkey
(Simia
aethiops)
Kidney COS cell
cultures
✓ – Transient expression Button and Brownstein
(1993)
COS7 cells ✓ ER Transient expression Kendall et al. (1994)
✓ Mitochondriaþouter
mitochondrial
membrane
Transient expression Brandenburger et al.
(1999)
Cow (Bos taurus) Bovine adrenal
medulla
chromaYn cells
� (used fura 2) ER 1� cell cultures (HSV
gene transfer
vector)
Alonso et al. (1998)
� (used fura 2) Mitochondria 1� cell cultures (HSV
gene transfer
vector)
Montero et al. (2000)
✓ (also Fura 4F) ER, nucleus
mitochondria,
1� cell cultures(HSV gene trans-
fer vector)
Villalobos et al. (2002)
✓ Nucleus 1� cell cultures (HSV
gene transfer
vector)
Chamero et al. (2002)
� Secretory granules 1� cell cultures (AdV
gene transfer
vector)
SantoDomingo et al.
(2008)
Bovine adrenal zone
glomerulosa cells
✓ Mitochondrial matrix 1� cell cultures(Transient
transfection)
Brandenburger et al.
(1996)
✓ Mitochondriaþouter mi-
tochondrial membrane
1� cell cultures(Transient
transfection)
Brandenburger et al.
(1999)
Mouse
(Mus
musculus)
Muller glial cells of adult
retinal explants
✓ – Explant culture. Ex-
pression of GA via
AdV gene transfer
vector
Agulhon et al. (2007)
A-11 (nonmetastatic)
and 3LL (metastatic)
Lewis lung cancer cell
lines
✓ – Stable expression Yoshida et al. (1998)
Soma and neurites from
adult superior cervical
ganglion neurons
� (used fura 2) Mitochondria 1� cell cultures (HSV
gene transfer
vector)
Nunez et al. (2007)
Pancreatic b-cells fromintact islets
� (used fura 2) Mitochondria Explant culture. Ex-
pression of GA via
HSV gene transfer
vector
Quesada et al. (2008)
Myoblasts isolated from
the extensor digi-
torum longus muscle
in mdx and normal
C57B1/10 mice
✓ Sub-sarcolemma 1� cell cultures(Transient
expression)
Basset et al. (2004)
Myotubes isolated from
hind leg muscles of
mdx and C57BL101
mice
✓ SR, mitochondriaþ -
plasma membrane
1� cell cultures(Transient
expression)
Robert et al. (2001a)
C2C12 skeletal muscle
cell line
� (used fura 2 AM) Mitochondria Stable expression Challet et al. (2001)
NIH 3T3 fibroblasts � (used Fura-2) ER HSV gene transfer
vector
Alonso et al. (1998)
✓ Outer surface of intracel-
lular membranes
Transient expression Biagioli et al. (2005)
MIN6 (pancreatic b-cellline)
� (used fura 2 AM) Mitochondria Stable expression Nakazaki et al. (1998)
� (used fura 2 AM) ER, secretary vesicle Transient expression
and AdV gene
transfer vector
Mitchell et al. (2001)
Intact pancreatic islets of
Langerhans from
Balb/c mice
✓ Nuclear Explant culture
(HSV gene trans-
fer vector)
Villalobos et al. (2005)
Neuro2A (neuroblasto-
ma cells)
✓ – Transient expression
of GA
Baubet et al. (2000)
Rat
(Rattus
norvegicus)
Cerebellar granule cells � (used fura 2) ER 1� cell cultures(HSV gene trans-
fer vector)
Alonso et al. (1998)
Skeletal muscle
myotubes
✓ (also used fura-2) Mitochondria, nucleus,
SR
1�cell cultures
(Transient
expression)
Brini et al. (1997)
� SR and ER 1� cell cultures(Transient
expression)
Robert et al. (1998)
✓ ER lumen and lumen of
the terminal cisternae of
the SR
1� cell cultures(Transient
expression)
Brini et al. (2005)
(continues)
Table II (continued )
Species (common
name
and/or scientific
name) Cell type
Cytosolic expression of
apoaequorin or
apoaequorin-BRET Organelle(s) targeted Comments References
L6 myogenic cell line � (used indo PE3 (AM)) ER Transient expression Narayanan et al. (2003)
Ventricular myocytes
from neaonatal
Wistar rats
✓ Mitochondria 1� cell cultures(Transient
expression)
Robert et al. (2001b)
A7r5 cells (aortic
smooth muscle cell
line)
✓ Plasma membrane Transient expression Marsault et al. (1997)
Aortic
smooth muscle cells
� (used fura 2 AM) Mitochondria 1� cell cultures(Transient
expression)
Szado et al. (2003)
Anterior pituitary cells � (used fura 2) ER 1� cell cultures(HSV gene trans-
fer vector)
Alonso et al. (1998)
GH3 cells (pituitary cell
line)
� (used fura 2) ER HSV gene transfer
vector
Alonso et al. (1998)
✓ Nucleus HSV gene transfer
vector
Chamero et al. (2002)
✓ Nucleus, ER Expression of GA
via HSV gene
transfer vector
Chamero et al. (2008)
PC12 cells (Adrenal me-
dulla pheachromo-
cytoma cell line)
� (used fura 2) ER HSV gene transfer
vector
Alonso et al. (1998)
✓ Nucleus HSV gene transfer
vector
Chamero et al. (2002)
✓ Secretory granule
membrane
Transient expression Moreno et al. (2005)
✓ Mitochondria Transient expression Dıaz-Prieto et al. (2008)
✓ Nucleus, mitochondria,
ER
Expression of GA
and RA via HSV
gene transfer
vector
Manjarres et al. (2008)
INS-1 cells
(derived from insulin-
secreting pancreatic
b-cell tumor)
✓ Mitochondria Stable expression Kennedy et al. (1996)
� ER Stable expression Maechler et al. (1999)
H4-IIE cells (hepatoma
cell line)
✓ ER Stable expression Chan et al. (2004)
Tail artery (from male
Wistar rats)
� SR Explant culture
(AdV gene trans-
fer vector)
Rembold et al. (1997)
Hamster
(Cricetulus
griseus)
CHO-K1 cells ✓ – Stable expression Button and Brownstein
(1993), Sanchez-Bueno
et al. (1996)
CHO.T cells ✓ Mitochondria Transient expression Rutter et al. (1996)
CHO cells ✓ Peroxisomes
Mitochondria, ER
Transient expression Lasorsa et al. (2008)
Fruit fly
(Drosophila
melanogaster)
Schneider 2 (S2) cells ✓ – Stable expression Torfs et al. (2002)
Kingdom: Plantae
Soybean (Glycine
max)
Cells in suspension of
[L]., cell line 6.6.12
✓ – Stable expression Mithofer et al. (1999)
Tobacco (Nicoti-
ana tabacum)
Leaf discs ✓ – Stable expression Cessna et al. (2000)
Parsley (Petrose-
linum crispum)
Cells in suspension ✓ – Stable expression Blume et al. (2000)
Kingdom: Fungi
Aspergillus
nidulans
Whole organism ✓ Mitochondria Stable expression Greene et al. (2002)
Saccharomyces
cerevisiae
Whole organism ✓ – Stable expression Nakajima-Shimada et al.
(1991)
� ER lumen Stable expression Strayle et al. (1999)
� Mitochondria Stable expression Jung et al. (2004)
Kingdom: Protista
Trypanosoma
brucei brucei
Procyclic cells ✓ Nucleus Stable expression Xiong and Ruben (1996)
✓ Mitochondria Stable expression Xiong et al. (1997)
AdV, Adenovirus; HSV, Herpes Simplex Virus; CHO.T cells, CHO cells that over-express human insulin receptors; HEK 293T, 293 cells transformed with large
T-antigen from SV40.
Table IIIExamples of the some of the reported coelenterazine loading protocols
Species Coelenterazine loading protocol reported References
Kingdom: Animalia
Mus musculis (Mouse) Native coelenterazine was introduced into adult mice (at 4 mg/kg) by tail-vein
injection and into new-born mice (at 2–4 mg/g) by intraperitoneal injection.
Light emission was recorded immediately
Rogers et al. (2007)
Minced tissues or cells of tissues were incubated with 0.2 ml RPMI 1640
containing 10 mM coelenterazine at 37 �C for 5 h
Yamano et al. (2007)
Danio rerio (Zebrafish) Embryos that had been dechorionated at the 64-cell stage were incubated with
50 mM f-coelenterazine, prepared in 30% Danieau’s solution
Cheung et al. (2006)
Xenopus laevis (African clawed frog) Oocytes were incubated in 2.5 mM coelenterazine in a medium containing
5 mM b-mercaptoethanol
Daguzan et al. (1995)
Drosophila melanogaster (Fruit fly) Malpighian tubules from 4 to 14-day old adults were incubated in Schneider’s
medium containing 2.5 mM coelenterazine for 4–6 h in the dark
Rosay et al. (1997)
Exposed fly brains were incubated in fly ringers solution containing 5 mMnative coelenterazine for >1 h at r.t.
Martin et al. (2007)
Kingdom: Plantae
Nicotiana plumbaginifolia (Tobacco) Seedlings were floated on water containing 2.5 mM coelenterazine o/n at r.t. in
the dark
Knight et al. (1991a, 1996)
Arabidopsis thaliana Seedlings were submerged in 10 mM coelenterazine (which had been dissolved
in ethanol) in dH2O for 7.5 h at r.t. in the dark
Sedbrook et al. (1996)
Seedlings were floated on water containing 2.5 mM coelenterazine o/n at r.t. in
the dark
Knight et al. (1996)
Solanum tuberosum (Potato) Plants were incubated in 5 mM hcp coelenterazine for 8 h Fisahn et al. (2004)
Physcomitrella patens (Moss) Ground up moss tissue was incubated in 0.5 MNaCl, 5 mMmercaptoethanol,
5 mM EDTA, 0.1% gelatin, 10 mM Tris–HCl pH 7.4 containing 2.5 mMcoelenterazine for 6 h or o/n
Russell et al. (1996)
Kingdom: Fungi
Neurospora crassa / Aspergillus niger /
Aspergillus awamori
Vogel’s medium containing 2.5 mM coelenterazine (prepared in methanol) was
inoculated with 1�105 spores/ml. Inoculated medium was incubated for
24 h at 30 �C in the dark
Nelson et al. (2004)
Phyllosticta ampelicida 2.5 mM coelenterazine was pipetted over colonies growing in 1/2� potato
dextrose agar and incubated for 4 h
Shaw et al. (2001)
Schizosaccharomyces pombe Cells were incubated in EMM medium containing 20 mM coelenterazine for
4 h at 30 �CDeng et al. (2006)
Saccharomyces cerevisiae 0.1 volume of 590 mM coelenterazine (prepared in methanol) was added to
25–30 ml yeast culture and incubated for 20 min at r.t.
Batiza et al. (1996)
Kingdom: Protista
Dictyostelium discoidium (Slime mold) Cells were incubated with coelenterazine solution to a final concentration of
50 mM for 24 h at r.t. in the dark
Cubitt et al. (1995)
Kingdom: Monera
Escherichia coli Cells, diluted in 100 mMKCl, 1 mMMgCl2, Tris–HCl, pH 7.5, were incubated
with coelenterazine (final concentration of 2.5 mM) o/n at r.t. in the dark
Knight et al. (1991b)
Bacillus subtilis Bacteria were incubated in TS medium containing 20 mg/ml kanamycin and
2.5 mM coelenterazine h for 1 h in the dark at r.t.
Herbaud et al. (1998)
Anabaena sp. Cells were incubated with coelenterazine (to a final concentration of 2.5 mM)
for 4 h in the dark
Torrecilla et al. (2000)
Tissue Culture Cells
H4-IIE cells Cells were incubated with 5 mM coelenterazine in Krebs–Ringer modified
buVer containing 1 mM EGTA for 1 h at r.t.
Chan et al. (2004)
Neuro2A cells Cells were incubated with 5 mM coelenterazine, 10 mM b-mercaptoethanol
and 4 mM EDTA in PBS for 2–4 h at 4 �CBaubet et al. (2000)
HeLa cells Cells were incubated with 2.5 mM coelenterazine for 6–8 h at 37 �C Brini et al. (1993)
Jurkat cells Cells were incubated with 5 mM coelenterazine-n for 2 h at 37 �C Narayanan et al. (2003)
COS7 cells Cells in suspension were incubated with 2.5 mM coelenterazine in DMEM o/n
at 4 �CKendall et al. (1994)
r.t. is room temperature; o/n is overnight.
Table IVExamples of the detectors used to image aequorin-generated luminescence
Company Type of detector reported Examples of specimens visualized References
Berthold Technologies (U.K.) Ltd,
Hertfordshire, UK
LB980 intensified tube camera Nicotiana plumbaginifolia (tobacco)
seedlings
Wood et al. (2001)
Biospace Lab., Paris, France Cooled GaAs ICCD (3rd generation) Mus musculus (mouse)—intact
animals
Rogers et al. (2007)
Dittie Thermografie, Bonn, Germany Giotto 1.12 microchannel plate-linked
image intensifier tube (1st generation)
Triticum aestivum (winter wheat)
seedlings
Nagel-Volkmann et al. (2009)
EG&G, Berthold Technologies (U.K.)
Ltd, Hertfordshire, UK
Luminograph LB980 low-light camera
system
Phyllosticta ampelicida Shaw et al. (2001)
Hamamatsu Photonics GmbH Deutsch-
land, Herrsching, Germany
C2400-40H ICCD Petroselinum crispum (parsley)
suspension cultures
Blume et al. (2000)
Hamamatsu Photonics Co., Hamamatsu,
Japan
Ultrasensitive VIM camera system (a CCD
camera equipped with an intensifier,
Model C-1400-47)
Arabidopsis thaliana intact plants Furuichi et al. (2001)
VIM photon counting camera Bovine adrenal chromaYn cells Villalobos et al. (2002)
Mouse superior cervical ganglion
neurons
Nunez et al. (2007)
Mouse intact pancreatic islets Villalobos et al. (2005),
Quesada et al. (2008)
C2400-20M ICCD CHO.T cells Rutter et al. (1996)
Photek Ltd., East Sussex, UK RA-IPD H4-IIE cells Chan et al. (2004)
Danio rerio (zebrafish) embryos Cheung et al. (2006)
IPD 3 Drosophila melanogaster (fruit fly)
mushroom bodies
Martin et al. (2007)
Adult mouse retinal explants Agulhon et al. (2007)
Photek 216 ICCD Arabidopsis thaliana seedlings Evans et al. (2005)
EDC-02 ICCD Arabidopsis thaliana seedlings/
leaves
Knight and Knight (2000),
Grant et al. (2000)
Photometrics, Tucson, AZ CH220 CCD imager Arabidopsis thaliana seedlings Sedbrook et al. (1996)
Photonic Science, Robertsbridge, UK Cooled extended ISIS video camera Neuro2A (mouse neuroblastoma
cells)
Baubet et al. (2000)
Princeton Instruments, Trenton, NJ TE/CCD512BKS CCD Nicotiana plumbaginifolia (tobacco)
seedlings
Sai and Johnson (2002)
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 277
improved light emission properties of GFP-aequorin are still not completely
understood, it is thought that they likely relate to an improved stability of the
apoaequorin protein or to an increased quantum yield of the Ca2þ-activatedphotoprotein complex, or perhaps both.
The cloned GFP-apoaequorin gene has been engineered to target diVerentsubcellular compartments using similar strategies as those developed and described
already for aequorin (Rizzuto et al., 1992). This approach has also been extended
to include other spectral variants of these aequorin-based reporters, by replace-
ment of the GFP gene with the sequences encoding Venus (YFP), mRFP, and
more recently, mOrange (Bakayan et al., 2009; Curie et al., 2007; Manjarres et al.,
2008). The development of these bifunctional reporters together with new imaging
technologies (see Section IV) has considerably extended the number of applica-
tions possible with aequorin and in particular, has facilitated important advances
in multicompartment measurements of Ca2þ concentrations and in noninvasive
whole animal Ca2þ-imaging studies of the mammalian system.
A. Expression of Apoaequorin
1. Protocol 1: Preparation of Transgenic Zebrafish that Express Apoaequorin in a Tissue-SpecificManner (e.g., in the skeletal musculature)
a. MaterialspiP-HE (apoaequorin plasmid; Inouye et al., 1989)
ap-SK plasmid (a-actin promoter; Higashijima et al., 1997)
pIRES2-EGFP plasmid (Clontech Laboratories, Inc., Mountain View, CA,
USA)
pCMVTNT vector (Promega Corp., Madison, WI, USA)
f-coelenterazine (C-6779; Molecular Probes, Invitrogen Corp., Eugene, OR,
USA)
Methyl cellulose (M0387; Sigma–Aldrich Corp., MO, USA)
b. Methodsi. Preparation of the pa-KS-aeq-IRES-EGFP plasmid. To prepare the pa-KS-
aeq-IRES-EGFP plasmid, use PCR to amplify the apoaequorin gene from the
piP-HE plasmid, with the following oligonucleotide primers: 50-accagaattcatgacaag-caaacaatactcagtcaagcttacatcagac-30 and 50-accagtcgacttaggggacagctccaccgtagag-30,such that EcoR1 and Sal1, are added to the 50 and 30 ends of the apoaequorin
gene, respectively. The apoaequorin gene can then be cloned into the pIRES2-EGFP
plasmid using these restriction sites. Excise the aeq-IRES-EGFP fragment with
EcoR1 and Not1, and then clone it into the ap-SK plasmid to obtain an aeq-
IRES-EGFP fragment with an a-actin promoter (i.e., a-aeq-IRES-EGFP). In paral-
lel, amplify the SV40 late polyadenylation signal (pA) from the pCMVTNT vector
using the following oligonucleotide primers: 50-accagcggccgccagacatgataagatacattg-30 and 50-accagagctctctagaaccggttaccacatttgtagaggtttt-30, adding Not1 to the 50 end,
278 Sarah E. Webb et al.
and Age1,Xba1, and Sac1 to the 30 end of the SV40 late polyadenylation signal. The
SV40 late polyadenylation signal can then be cloned into the pBluescriptII-KSþ
plasmid with Not1 and Sac1, after which the a-aeq-IRES-EGFP fragment can also
be cloned into this plasmid using Xho1 and Not1. The enhanced green fluorescent
protein (EGFP) marker gene is regulated by the IRES-sequence for the subsequent
identification of transgenic fish. Use of the IRES-sequence enables the translation of
both apoaequorin and the EGFP marker from a single mRNA; thus, the expression
level and distribution of EGFP reflects the expression level and distribution of
apoaequorin (Fahrenkrug et al., 1999; Wang et al., 2000).
ii. Generation of transgenic zebrafish that express apoaequorin in the skeletal
muscles. Linearize the pa-KS-aeq-IRES-EGFP plasmid with Xba1 and then
microinject �1 nl (i.e., �100–200 pg) into the center of the blastodisc of the
zebrafish embryos at the one-cell stage. The microinjection pipettes and pressure
injection system used are described in detail in Webb et al. (1997). The injected
embryos should then be maintained at�28.5 �C and screened for the expression of
EGFP after 24 hpf. Embryos (F0) that express EGFP should be raised to adult-
hood for further transgenic germ line screening. This involves: (1) The F0 fish being
crossed with the wild-type fish to get the F1 generation. (2) If some of the F1
embryos express EGFP, then this indicates that one of their parents was transgenic
(i.e., it was heterozygous). The F1 embryos that express EGFP can then be raised
to adulthood and intercrossed with one another to produce the F2 generation.
(3) In the F2 generation, 50% of the oVspring should be heterozygous, 25% should
be homozygous, and 25% should be wild-type. (4) The homozygous F2 transgenic
fish may then be identified by crossing the EGFP-expressing fish with wild-type
fish; if all of the F3 oVspring express EGFP, then their transgenic parent was a
homozygote; if 50% of the F3 embryos express EGFP, then their transgenic parent
was heterozygote. The homozygous F2 fish can then be intercrossed with one
another to obtain stable transgenic lines.
iii. In vivo reconstitution of aequorin. Dechorionate the a-actin-apoaequorintransgenic embryos when they are at the eight-cell stage (we dechorionate embryos
manually with watchmaker’s forceps) and incubate them in a custom-designed
injection/imaging chamber (described in Webb et al., 1997) with 20 mg/ml f-coe-
lenterazine in 30% Danieau’s solution to reconstitute the active aequorin. Prepare
the f-coelenterazine as a stock solution of 2 mg/ml in methanol and dilute it in 30%
Danieau’s solution just prior to use. In this transgenic F2 fish line, the EGFP and
thus the apoaequorin, are expressed in the musculature at low levels at �12 hpf
(i.e., the�6-somite stage) and the level of expression increases in an approximately
linear manner up to�24 hpf. In addition, at�24 hpf EGFP and thus apoaequorin
were expressed throughout the entire musculature, that is, in both the slow and fast
muscles. Thus, this line of muscle-specific apoaequorin-expressing transgenic
zebrafish can be used to visualize and characterize the Ca2þ signals generated in
the trunk musculature during its formation and function. For imaging, these later-
stage embryos may be immobilized with 3% methyl cellulose. We have collected
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 279
both temporal and spatial information regarding the Ca2þ signals generated by the
musculature from these embryos up to 96 hpf. For imaging, we use two custom-
built Photon ImagingMicroscope Systems (PIMS; Science Wares, Falmouth, MA,
USA), one based on an IPD-425 (Photek Ltd., Sussex, UK) and the other based on
a back-illuminated EMCCD (DU-897 iXonEMþ camera) that was purchased from
Andor Technology (Belfast, Northern Ireland, UK) and then optimized by Science
Wares Inc. for detecting single photon events at the emission rates typical for
aequorin-based imaging (see Section IV for further details).
B. Expression of GFP-Apoaequorin
2. Protocol 2: Transient and Stable Transfection of Mammalian Cells with GFP-ApoaequorinUsing Plasmid DNA
Many of the expression vectors designed for gene delivery are commercially
available. They contain a number of units, including an immediate-early enhancer/
promoter sequence such as human cytomegalovirus (HCMV), a multiple cloning
region for insertion of the reporter gene, an antibiotic resistance gene (e.g., Ampi-
cillin) for selection of the vector in Escherichia coli, and an additional antibiotic
resistance gene for selection in mammalian cells (e.g., Neomycin G148). For
mammalian expression, a good starting point is to clone the reporter gene into a
vector containing an HCMV promoter (Williams et al., 2005). The constitutive
immediate-early HCMV promoter drives high levels of GFP-aequorin expression
in many mammalian cell lines following transient transfection.
Transfection reagents (e.g., cationic liposomes) enable recombinant DNA deliv-
ery into the nucleus of many immortalized cell types such as HEK293, HeLa,
COS7, CHO and NIH/3T3 cells, with high eYciency. Following transient trans-
fection, stable clones can then be isolated using a combination of drug selection
(e.g., Neomycin (G418) resistance) and cell sorting using flow cytometry. On the
other hand, primary cells (e.g., cortical neurons) are usually transfected with very
low eYciency (i.e., less than 1–5%) using this method and better results can be
obtained by using recombinant viral vectors for gene delivery (Rogers et al., 2005).
a. MaterialsTransfection reagent (e.g., FugeneÒ6 reagent, Roche Applied Science; Lipofec-
tamine, Invitrogen; PolyFect, Qiagen).
Ultrapure plasmid DNA (1 mg/ml)—Plasmid DNA can be purified using a
plasmid purification kit (e.g., QiagenÒpurification kits).
Optimem media (Invitrogen)
35-mm Petri dishes or 8-chamber slides (e.g., those available fromMatTek corp.
or Ibidi Gmbh). Any culture dish will do providing the bottom of the dish is
optimized for high-resolution microscopy on an inverted setup (e.g., dishes
prepared with a glass coverslip mounted underneath a hole cut in the bottom).
280 Sarah E. Webb et al.
This allows the use of objective lenses with high numerical apertures for maximum
light collection.
Native or h-coelenterazine (supplied by Molecular Probes, Invitrogen, US or
Interchim, France; 1-5 mM stock solution prepared in 100% ethanol).
b. Methodsi. Preparation of cells that are transiently transfected with GFP-
apoaequorin. Healthy cell monolayers can be transfected when they are approxi-
mately 50–75% confluent and imaged within 24–48 h following transfection. Wash
cells 1� and resuspend with serum-free medium (with no antibiotic/antimycotic)
and then place cells back into the 37 �C/5% CO2 incubator. For transfecting a 35-
mm dish, prepare a microcentrifuge tube containing 150 ml of serum-free media.
Add 4.5 ml of transfection reagent (this amount may be increased according to the
cell type or reagent used). Vortex the tube and then add 1.5 ml of plasmid DNA
(1 mg/ml). Mix by flicking the tube and leave the tube at room temperature for
approximately 30–45 min to allow the formation of a complex. Following incuba-
tion, remove the cells from the incubator and add 100 ml of the mix drop-wise and
gently swirl the dish before placing back into the incubator. Although optional,
healthy cells can be more easily maintained if the medium is changed after 6 h with
fresh medium containing fetal calf serum (FCS). After 24–48 h, replace the normal
growth medium with a serum-/phenol red-free medium (used for imaging), and
incubate the cells for 1 h with 5mM coelenterazine (native or h-according to the
type of study). Single cells expressing high levels of GFP-aequorin can be selected
by their GFP fluorescence, and used for bioluminescence Ca2þ imaging studies.
ii. Preparation of stable cell lines expressing GFP-apoaequorin. Follow the
protocol for preparing cells for transient transfection (see previous section). At
48 h after transient transfection, start the selection process by adding a selective
medium containing the appropriate antibiotic (e.g., Geneticin, or Puromycin). The
antibiotic concentration used (starting at an upper concentration of 1 mg/ml)
needs to be optimized for diVerent cell types. The medium should be changed
with fresh selective medium every 2–3 days over a period of several weeks. During
this time, the concentration of antibiotic may be gradually decreased. Isolation of
GFP-positive clones into 96-well plates can then be facilitated using flow cytome-
try. In our experience, approximately 10% of the clones survive and proliferate
after a first round of FACS sorting. Since stable transfection is a random integra-
tion event and a large amount of variability is expected, the clones should be
selected based on the level of GFP intensity and homogeneity as well as cellular
morphology, and where instrumentation is available (e.g., Nikon’s Biostation or
the Incucyte from Essen Instruments), clones can also be selected based on growth
curves.
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 281
3. Protocol 3: Preparation and Transfection of Organotypic Brain Slices with RecombinantAdenovirus-5 Vector Containing the GFP-Apoaequorin Gene
Recombinant viral vectors, such as human adenovirus serotype 5 (Ad5), adeno-
associated virus (AAV), Sindbis viruses, or retroviruses (e.g., Lentiviruses), are
highly eYcient expression vectors for gene delivery in mammalian cells or tissues
(Tenenbaum et al., 2004). Both Ad5 and the Sindbis virus have been used to
mediate high levels of expression of GFP-apoaequorin in primary neuronal cul-
tures, brain slices, and retinal explant cultures (Rogers et al., 2005). Ad5 was found
to preferentially infect glial cells in cortical- or hippocampal-derived tissue as well
as Muller cells (a type of glial cell) in retinal explant cultures (Rogers et al., 2005).
a. MaterialsVibratome (e.g., Model VT-1000, Leica)
Gas tank (95% O2 / 5 % CO2)
Glass petri dish (3–5 cm diameter)
Membrane filter inserts (e.g., 12 mm TranswellÒ Permeable Supports, Corning)
12-well tissue culture plates
Artificial cerebrospinal fluid (ACSF), pH 7.4 (124 mMNaCl, 3 mMKCl, 2 mM
CaCl2, 1.3 mMMgCl2, 25 mMNaHCO3, 1.25 mMNaHPO4, and 10 mM glucose)
Superglue, Blades, Low temperature melting Agar, Culture medium (50 %
MEM, 25 % HBSS, 25 % Horse Serum, 6.5 mg/ml glucose, 2 mM L-glutamine,
100 U/ml penicillin, 100 mg/ml streptomycin, pH 7.2).
Ad5-GA Viral particles (1�5 x 108 particles).
Specimen chamber for live imaging
b. MethodsOrganotypic slices can be prepared similarly to previously reported methods
(Stoppini et al., 1991). Briefly, prepare 400-600 mm slices as described in section 3b.
Once the slices are cut, gently transfer them using a Pasteur pipette onto a culture
membrane insert and into a 12-well culture plate containing prewarmed culture
medium. Slices can be kept in culture for 4–5 days and viral particles added directly
to the medium approximately 48 h prior to imaging. After viral transfection and
verification of GFP expression, the membrane culture insert with attached slice can
be moved to a larger Petri dish containing growth media (e.g., 35 mm) and the
membrane excised with a scalpel blade. Care should be taken to avoid folding of
the membrane insert, which will cause injury to the tissue. The membrane with
attached slice should then be carefully inserted into an imaging chamber and a
tissue anchor placed on top to secure the tissue (e.g., Series RC-20, Harvard
Apparatus). Once the slice is mounted onto an inverted microscope, a simple
gravity flow system for delivering buVer together with a small peristaltic pump
connected to the output can be used for perfusion (Mohammed et al., 2007).
282 Sarah E. Webb et al.
i. Transgenic mice expressing GFP-apoaequorin reporters. GFP-aequorin has
also been expressed in a number of mammalian cell lines, as well as in organisms
such as Plasmodium bergei (Billker et al., 2004), Drosophila melanogaster (Martin
et al., 2007), and in mice (Rogers et al., 2007). Transgenic mice can provide a
source of cells, tissues, or organs for studies ex vivo or for studies in vivo (Rogers
et al., 2007). In the case of targeted reporters, these are especially useful because
they provide information regarding the localization of any probe-derived signals,
which would otherwise be diYcult because of the low to moderate resolution
aVorded by bioluminescence imaging. In addition, an inducible or conditional
expression system could be introduced into the vector (e.g., Cre/Lox or Tet
inducible elements), to ensure the absence of a phenotype in the event that expres-
sing the reporter ubiquitously from the early stages of development is found to be
detrimental. A conditional expression system also enables expression to be acti-
vated in a specific cell population or at diVerent stages of development. Indeed, the
UAS-Gal4 system enables the specific expression of GFP-aequorin in neuronal
subsets of the fly brain, allowing specific neuroanatomical mapping of Ca2þ
signaling pathways (Martin et al., 2007).
Transgenic mice can be generated via one of two methods: (1) using a ‘classical’
transgenesis approach, where the transgene is randomly integrated into the
genome (Constantini and Lacy, 1981), or (2) by homologous recombination,
which enables directed integration of the transgene (e.g., knock-in of the HPRT
locus; Rogers et al., 2007). Similarly to pronuclear injection, lentiviral vectors can
be used to deliver the transgene into the fertilized mouse egg (Ikawa et al., 2003).
However, these methods can result in random and multiple integrations of the
transgene. In contrast, homologous recombination targets transgene insertion in a
single-copy to a known site in the genome ensuring a more predictable expression
pattern and phenotype based on the known integration site (Bronson et al., 1996).
Transgenic mice conditionally expressing mitochondrially targeted GFP-apoae-
quorin have already been generated using this method (Rogers et al., 2007).
Targeted insertion of the transgene was made 5’ to the X-linked hypoxanthine
phosphoribosyltransferase (HPRT) locus (X-chromosome).
4. Protocol 4: Preparation of Acute Brain Slices from Transgenic Mice ExpressingMitochondrially Targeted GFP-Aequorin
a. MaterialsVibratome (e.g., Model VT-1000, Leica)
Oxygen (95% O2/5% CO2)
Glass petri dish (3–5 cm diameter)
ACSF, pH 7.4 (124 mM NaCl, 3 mM KCl, 1.3 mM MgCl2, 25 mM NaHCO3,
1.25 mM NaHPO4 and 10 mM glucose)
Superglue, Blades, Low temperature melting Agar
Specimen chamber for live imaging (e.g., RC-20 chamber from Harvard
apparatus)
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 283
b. MethodsACSF should be prepared fresh on the day of experiments. Two 50 ml falcon tubes
filledwithACSF (Ca2þ free) canbeplacedat�20 �Cfor approximately 1–2hprior to
the preparationof brain slices. This partly frozenmedium is used tofill the bath on the
Vibratome where the brain will be sliced. Once the Vibratome is ready, rapidly
remove the brains from neonates, dissect the cerebellum away, and place the brain
ventrally against the agar block. Horizontal or coronal slices (400 mm) can be cut and
transferred immediately to a small Petri dish containing oxygenated ACSF and
coelenterazine (10 mM) and incubated at room temp, in the dark for 45–60 mins.
Once the slice has been inserted into the imaging chamber, a slice anchor (Harvard
Apparatus) can be used to secure the tissue. Slices shouldbe continuously perfused (at
a flow rate of 1 ml/min) with oxygenated ACSF containing 2 mM CaCl2. Biolumi-
nescence signals can be monitored as previously described (Rogers et al., 2007).
C. Summary of Section II
The commercial development of products, kits, or services enabling genetic
engineering of cells or animals means that these technologies are no longer out
of reach to biologists who have experience in imaging but who have little molecular
biology experience. BRET-based imaging depends on the degree of spectral over-
lap, relative orientation, and the distance between donor and acceptor dipoles.
Fluorescent proteins derived from coelenterates or their variants are therefore
likely to be the most suitable acceptors owing to their structural similarity with
GFP (Tsien, 1998). The recent development of RFP-aequorin (Manjarres et al.,
2008) has provided the means to simultaneously monitor Ca2þ signals from two
diVerent microdomains within a single cell.
III. Introducing Coelenterazines into Cells, Tissues and Embryos
Coelenterazine is the small (�400 Da) prosthetic group that binds with apoae-
quorin to form the active aequorin complex. As coelenterazine is subject to oxida-
tion, it is normally supplied in a sealed vial free of O2. Prior to reconstitution, the
coelenterazine should be stored at �20 �C. In addition, coelenterazine is poorly
soluble in water and so stock solutions are normally prepared in methanol. In this
form, coelenterazine is stable for �3 months at �20 �C. Coelenterazine was
originally isolated from Aequorea victoria; however, in the 1970s a procedure for
chemically synthesizing coelenterazine was developed (Inoue et al., 1975; Kishi
et al., 1972). This procedure has since been used for preparing coelenterazine and
its analogs (Jones et al., 1999). Many of the coelenterazine analogs possess proper-
ties diVerent from those of native coelenterazine. These include half-life, aequorin
regeneration rate, luminescence capacity, emission maximum and membrane
permeability, the latter being due to the lipophilic nature of coelenterazine.
For example, f-coelenterazine has the same half-life as native coelenterazine
284 Sarah E. Webb et al.
(Shimomura, 1991) but when it is reconstituted with apoaequorin to form an
aequorin complex (f-aequorin) the level of luminescence produced on reaction
with Ca2þ is almost 20-fold higher than that produced when native coelenterazine
is used. In addition, f-coelenterazine has the highest permeability through cell
membranes (Shimomura, 1997).
As coelenterazine is lipophilic, apoaequorin-expressing cells, tissues, and whole
organisms can simply be incubated in coelenterazine solution. However, this
method is successful only in tissue culture cells and in simple organisms that
have a large surface area-to-volume ratio where eYcient diVusion occurs. In
more complex, multicellular organisms such as developing vertebrate embryos,
reconstituting aequorin is more of a challenge. In the case of our a-actin-apoae-quorin transgenic zebrafish, we started our f-coelenterazine incubation as early as
the eight-cell stage (i.e., 1.25 hpf ) when the embryonic cells had a large surface
area-to-volume ratio, and embryos were incubated continually in this 20 mg/ml
coelenterazine both up until, and during data collection, which took place from 16
to 48 hpf. In the case of the apoaequorin-expressing transgenic mice, Rogers et al.
(2007) introduced native coelenterazine into adult mice (at 4 mg/kg) by injection
into the tail vein and into new-born mice (at 2–4 mg/g) by intraperitoneal injection.These, and other protocols used for introducing coelenterazine into various intact
organisms and tissue culture cells are summarized in Table III.
IV. Techniques for Detecting Aequorin Luminescence
Currently, several diVerent types of equipment are commercially available that
can be used to detect or visualize aequorin-generated luminescence. These range in
capability, price, design, and commercial availability. At the lower cost end, there
is the simple test tube/culture dish luminometer, which provides only temporal
Ca2þ signaling information, costs just a few hundred US dollars, and is supplied by
several diVerent companies. At the higher cost end are several custom-designed
imaging systems, which provide both temporal and spatial luminescent informa-
tion, as well as bright-field and fluorescence images (if required), to enable the
correlation of Ca2þ signaling events with morphological features and other cellular
changes. These systems are obviously a lot more costly and are built to order by a
small number of specialist companies. Some examples of the types of detectors that
have been used to image aequorin-generated luminescence are shown in Table IV.
It may be diYcult to justify the purchase of expensive single photon imaging
equipment at early stages in a project. Often a photon counting photomultiplier
tube (PMT) can be used in place of an imaging photon detector (IPD) to determine
the timing and amplitude of bioluminescence signals in living systems. By adding a
near-IR light source and an appropriate blocking filter in front of the PMT, a
relatively inexpensive near-IR sensitive camera can be used to continuously moni-
tor morphological development while the PMT reports total bioluminescence
activity. An example of this type of system is shown in Fig. 2A. Accurate correlation
Dark box
IRcamera
Zoomlens
Feed-through
Shutter
Light-proofchamber
FilterLens
IRlighting
Beam-splitter Lens
IR-sensitivecamera
Sample stage(large field of view)
Doorswitch
HV/amplifier/discriminator
PMT
PMT1
ICCD
Sample
ManualPMTshutterIR
blockingfilter
IR ring light
Temperature-controlledstage
Monitor
ComputerPMT system
controller
PerfusionPerfusion
B
A
C
Mirror
CellsGlass
window
Dichroicmirror
(585 DCXR)
D630-60
PMT2
D535-50
Perfusionchamber
1252
IRimage
Count plot
Fig. 2 Schematic representations of three recently developed luminescence detection systems. (A)
A system that combines a PMT with an IR light source and an IR-sensitive camera. This enables both
single photon detection in the visible light spectrum and monitoring the sample continuously in real-
time using IR light. The system was designed and built by Science Wares, Inc., (Falmouth, MA). (B)
A two-channel luminometer, which was designed to simultaneously collect temporal Ca2þ signaling
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 285
286 Sarah E. Webb et al.
of bioluminescence signals with morphological events facilitates planning, optimi-
zation, and standardization of protocols for subsequent imaging experiments.
Over the past few years, several novel aequorin-based detection system designs
have been reported. One is the two-channel luminometer, which was made for
dual-wavelength aequorin measurements (see Fig. 2B). This system was designed
by Manjarres et al., 2008 and built by Cairn Research (Faversham, UK). Here, the
luminescence emission from two spectrally distinct aequorins (GFP-aequorin and
RFP-aequorin) that are coexpressed in diVerent subcellular locations within the
same cells is divided by a dichroic mirror and the resulting beams of light are
filtered at 535 and 630 nm and then collected by two separate PMTs (Manjarres
et al., 2008).
Another new Ca2þ detection device is an imaging system capable of acquiring
real-time bioluminescence data from living (and unrestrained) small animals such
as mice (see Fig. 2C). This system was designed by Roncali et al. (2008). It is based
around a Photon ImagerTM intensified CCD camera (Biospace Lab., France)
operating in a photon counting mode. The ICCD camera is set on top of a light-
tight chamber and records optical signals at a video rate of 25 Hz. Motion can also
be monitored by using two cameras, one that records the signal of interest and the
other that is used to video-track the animal. The latter can be achieved by
illuminating the field of view with infrared light. The signals from both cameras
are recorded simultaneously and electronically synchronized. A detailed descrip-
tion of this equipment is given by Roncali et al. (2008). A similar approach has
been used with microscope-based imaging systems to continuously acquire biolu-
minescence image data emitted at short wavelengths while using longer wavelength
illumination to simultaneously record transmitted light images that show mor-
phology (Speksnijder et al., 1990).
One of the most recent developments in bioluminescence detection involves
using an EMCCD detector for single photon imaging (Martin et al., 2007;
Rogers et al., 2008). The best bioluminescence imaging detectors are capable of
single photon detection, and this requires that the detector somehow amplify
the detected signal above background noise. All electronic imaging detectors
ultimately convert incident photons from the sample into detected electrons,
information from cells expressing GFP- and RFP-aequorin in diVerent organelles. The system was
designed by I. M. Manjarres, P. Chamero, M. T. Alonso, and J. Garcıa-Sancho (Universidad de
Valladolid and Consejo Superior de Investigaciones Cientıficas, Valladolid, Spain), and B. Domingo,
F. Molina and J. Llopis (Universidad de Castilla-LaMancha, Albacete, Spain), and was built by Cairne
Research (Faversham, UK). (C) A photon counting-based system, with a video monitoring function
(via an IR-sensitive camera), for whole-body optical imaging of un-restrained, freely moving small
animals, such as mice. The system was designed by E. Roncali, K. L. Rogers and B. Tavitian (Labor-
atiore d’Imagerie Moleculaire Experimentale, INSERM U803, Orsay, France) and M. Savinaud,
O. Levrey and S. Maitrejean (Biospace Lab, Paris). Panels (B) and (C) are modified from Fig. 1 in
Manjarres et al. (2008) and Roncali et al. (2008), respectively.
Sample Photo
Photons
Photo
ns
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 287
which get amplified by an electronic circuit. The output of the circuit is monitored
by a computer for recording purposes, and the collected data is then postprocessed
to generate meaningful images (see Fig. 3).
Bioluminescence can be diYcult to image because there is often very little signal,
and because it is diYcult to predict exactly when and where the signal will be
produced. Because the signals are often quite small, it is important to use the most
eYcient optical system possible to collect the light. An overview of the main types
of optical systems used for collecting light from bioluminescent samples is given by
Karplus (2006) and see Fig. 4. In microscopy, it is important to select an objective
with the highest ratio of numerical aperture to magnification (NA/mag). In macro-
imaging, it is important to select a lens with smallest possible working distance and
the lowest possible f-stop, typically known as a ‘‘fast’’ lens. It can be helpful to have
an electronic source on hand to test the eYciency of an optical system in biolumi-
nescence imaging mode and ensure that the various components of the system are
performing as expected (Creton and JaVe, 2001).Another important consideration for bioluminescence imaging is that the sam-
ple being imaged must be kept in a light-tight enclosure to eliminate any direct or
reflected ambient light from reaching the imaging detector, as it could easily
overwhelm the bioluminescence signal. Because the sample is normally kept in
complete darkness during bioluminescence imaging, it can be beneficial to periodi-
cally obtain bright-field and fluorescence images to determine the stage of devel-
opment or morphological condition of the sample.
Such bright-field and fluorescence images are typically orders of magnitude
brighter than bioluminescent images, so they can be obtained in a very short
period of time compared with that required to accumulate a meaningful biolumi-
nescence image. Until the recent development of deep cooled, back-thinned, elec-
tron multiplying charge-coupled devices (EMCCDs), detectors capable of single
photon imaging for bioluminescence were not well suited to acquiring bright-field
and fluorescence images because they used microchannel plates to amplify and
transmit images inside the detector. Microchannel plates blur and distort the
OpticsImaging
detector andelectronics
ns Photons Electrons Computerrecording
Photon dataprocessing
Photon images
Fig. 3 In bioluminescent imaging, some of the photons emitted from a sample are collected by an
optical system and directed onto a detector that converts the incident photons into electrons. These
electrons are processed by electronic circuits that provide data to a computer indicating the time and
position of the detected photons. The computer program then postprocesses the photon data to generate
images.
Imaging mode
Imagedata
Photonimagingsoftware
Shutter driver
Multicore CPU
Dark box
Dark box
BioluminescenceA
B
C
Bright-field Fluorescence
Fluorescencecondenser
Fluorescencecondenser
Epifluorescentfilter set
Epifluorescentfilter set
Fiber optic light guide
Light frommercuryarc lamp
Fiber opticlight guide
Bright-fieldcondenser
Electronic shutter
Electronic shutter
Bioluminescent/fluorescent sample
MotorizedXY stage
MotorizedObjective turret
Motorizedfilter turret
Tubelens
Reducinglens
Electronicshutter
Light fromhalogen lamp
FL slider
Motorizedmirror
Tubelens
Reducinglens
Electronicshutter
*Waterchiller
*Only required for RA-IPD
RA
-IP
D s
yste
mE
MC
CD
sys
tem
alo
neC
omm
on fo
r E
MC
CD
,R
A-I
PD
and
PM
T
EMCCD EMCCD
RA-IPDRA-IPD RA-IPD
CCDCCDCCD
EMCCD
Waterchiller
To microscopeinterface (in CPU)
To microscopeinterface (in CPU)
To EMCCD interface (in CPU)
To RA-IPD interface (in CPU) andRA-IPD controller
To framegrabber(in CPU)
EMCCD interface
Microscopeinterface
Imagedata
Photonimagingsoftware
Shutter driver
Multicore CPU
RA-IPDcontroller
Microscope interface
RA-IPD interface
Frame grabber
Motorizedfocus
Motorizedfocus
High NAobjective
Halogenlamp
Mercuryarc lamp
Fig. 4 Schematic representation of luminescence imaging systems based around a modified EMCCD
and an RA-IPD, which can be used to acquire bright-field and fluorescence imaging information as wel
as collect bioluminescence data. (A) Components that are common to both the EMCCD- andRA-IPD
based systems. (B and C) Components that are specific for the (B) EMCCD- and (C) RA-IPD-based
288 Sarah E. Webb et al.
l
-
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 289
images they transmit, have a limited dynamic range, and can also be permanently
damaged if they are exposed to too much signal. While a nonintensified detector
could be used to acquire bright-field and fluorescence images, the problem of blur
and distortion in the bioluminescence image would still remain. In addition, it was
a challenge to adjust the size and registration of images acquired from two diVerentdetectors.
An EMCCD sensor can be fabricated on a substrate alongside one or more
amplifiers with programmable gain settings, so a computer can rapidly change
from a low-gain to a high-gain setting. Thus, a computer-controlled EMCCD is
capable of acquiring bright-field and fluorescence images at low-gain settings, as
well as bioluminescence images at high-gain settings (see Fig. 4). This makes it
possible to eliminate blurring and distortion in all three types of images to be
acquired, improving the spatial resolution in the bioluminescence images by 2–3
times over what can be achieved with a microchannel plate based detector. Because
all three types of images can be obtained with the same sensor, the scale and
registration are identical as well. Furthermore, back-thinned EMCCDs have a
significantly higher quantum eYciency in the visible light spectrum, compared with
photocathode materials used in intensifier-based detectors, so they are able to
respond more eYciently to weak signals.
While EMCCDs are capable of detecting single photon events when the electron
multiplying gain is high enough to overcome the read noise of the output amplifier,
the electron multiplying gain mechanism is subject to substantial statistical varia-
tion. For example, when an output signal of 1000 electrons occurs, it is easily
detected as a meaningful event, but it is not possible to be certain how many input
electrons generated this signal—it may have been just 1, or 2, or 5 input electrons.
As a result, EMCCDs operating in photon counting mode have limited ability to
track large intensity changes. The maximum signal intensity that can be recorded
reliably is essentially determined by the frame rate at which the sensor is read out.
The range can be extended at the expense of the field of view by selecting a small
region of interest, and/or at the expense of spatial resolution by binning together
adjacent pixels on the sensor.
Two additional limitations of EMCCDs arise from the circuitry used to read out
the image data. First, in order to record the signal detected by the CCD sensor,
systems, respectively. The EMCCD can be modified to acquire bioluminescence information as well as
bright-field and fluorescence images. On the other hand, a resistive-anode Imaging Photon Detector
(RA-IPD) can be used in conjunction with a CCD camera, the latter to acquire bright-field and
fluorescent images, when a higher dynamic range and temporal resolution are needed for the biolumi-
nescence signal. For both the EMCCD and RA-IPD-based imaging systems, a high level of automation
for the microscope makes it possible to rapidly switch between the various imaging modes, and also
makes it possible to have the computer run automated acquisition sequences over extended periods,
typically overnight. The motorized focus allows the computer to acquire image stacks in any imaging
mode for three dimensional reconstructions. Both systems were designed and built by Science Wares
Inc., Falmouth, MA, USA.
290 Sarah E. Webb et al.
there must be a period of dead time while the accumulated image data is shifted out
of the active area of the sensor. Second, the signals used to shift the image data
from the active area into the readout amplifier can also impose noise on the output
that looks identical to single photon events in the photon counting mode. Signifi-
cant advances have been made to minimize the noise (called clock-induced charge)
that is generated. However, these two eVects are still significant such that increas-
ing the readout rate beyond a certain level actually degrades the net single photon
imaging performance of the EMCCD.
In situations where significant bioluminescence intensity changes are taking
place over periods less than a second, an intensifier-based detector is likely to be
a better choice than an EMCCD (see Fig. 4). The spatial resolution of commer-
cially available intensifier-based detectors is usually in the order of tens of microns
at the detector input window, which is adequate for many applications, but not as
good as what can be achieved with commercially available EMCCD detectors,
which typically have a pixel size of 8 or 16 mm. There are two main types of
intensifier-based detectors, those with a phosphor image output that is optically
coupled to a visible light CCD, and those with an electrically encoded anode that
produces position sensitive pulses for each detected event. The temporal resolution
of detectors with an optically coupled phosphor output is typically in the range of
tens of milliseconds due to the persistence time of the phosphor and the frame rate
of the CCD.
The best temporal resolution for single photon imaging can be in the order of
tens of nanoseconds, and is achieved only by intensifier-based detectors with an
electrically encoded anode. The dynamic range of such detectors is constrained by
the dead time of the pulse processing electronics, which is typically in the order of a
few microseconds per detected event. Improvements continue to be made in the
spatial resolution of microchannel plates and throughput speed of encoded anode
detectors (Lapington, 2004; Siegmund et al., 2005), but the cost and complexity of
operating such detectors has prevented them from being used widely for biolumi-
nescence imaging so far.
Another factor that should be considered when selecting a detector for biolumi-
nescence imaging is the expected signal-to-noise ratio of the recorded image data
(Karplus, 2006). Frequently, accepting lower spatial resolution can result in a
better signal-to-noise ratio. In situations where a fast or brief signal needs to be
identified with high temporal accuracy, a photocathode detector is often capable of
a better signal-to-noise ratio than an EMCCD. Even though an EMCCD detector
can have 2�–20� higher quantum eYciency than a detector with a photocathode,
at the high readout rates needed for good temporal resolution, incident photons
can still be lost during the dead time needed to transfer image data into the readout
frame of the EMCCD, and the photons that are detected can be obscured by clock-
induced readout noise. Figure 5 shows a comparison of the bioluminescence
images acquired by an EMCCD and an RA-IPD photon imaging system at two
diVerent stages of zebrafish development.
Cleavage period
i iAPview
Sideview
APview
Sideview
4-cell stage1 hpf
ii EM-CCD
1 2 3 4 >5 1–2 3–4 5–6 7–8 >9 1 2 3 4 >5 1–2 3–4 5–6 7–8 >9
Pos
ition
ing
Pro
paga
tion
Dee
peni
ngA
ppos
ition
iii IPD 425 ii EM-CCD iii IPD 425
Sphere stage4 h
Blastula periodBA
Fig. 5 Comparison of the bioluminescence images acquired by the EMCCD and RA-IPD photon
imaging systems during (A) the Cleavage Period and (B) Blastula Period of zebrafish development. (Ai
andBi) Schematics of an embryo froma side (animal pole—AP) viewand top viewat the (Ai) Four-cell stage
(i.e., �1 hpf) and (Bi) sphere stage (i.e., �4 hpf) to show the morphology of the embryo and the typical
patterns of Ca2þ signals (in red) observed at these two stages of development. (Aii, Aiii and Bii, Biii)
Representative AP views of f-aequorin loaded embryos to show the changes in intracellular free Ca2þ that
occur (Aii andAiii) at diVerent timesduring the secondcell division cycle (i.e., two- to four-cell stage) and (Bii
andBiii) at sphere stage. The imageswere acquired using anAndorEMCCD-based imaging system (Aii and
Bii) and a Photek IPD 425-based imaging system (Aiii and Biii). In both cases, luminescence was accumu-
lated for 30 s. Color scales indicate luminescent flux in photons/sec. Scale bars are 200 mm.
10. The Use of Aequorins to Record and Visualize Ca2þ Dynamics 291
292 Sarah E. Webb et al.
V. Conclusions
Living cells, tissues and whole organisms are essentially defined by their complex
spatial structures. Underlying such broad morphological characteristics are much
finer molecular assemblies, such as microdomains within the cytoskeleton, plasma
membrane, nucleus and cytoplasm. It is becoming clear that such microdomains are
the loci of many key signaling events, including those that involve Ca2þ and as such,
they are becoming amajor area of interest for investigators in theCa2þ signaling field.
In recent years, researchers have made spectacular advances on the live-cell imaging
front, due in part to the development of techniques that combine breaking Ernst
Abbe’s diVraction barrier (Abbe, 1873) being compatible with examining living
systems. Such techniques include stimulated emission depletion (STED) microscopy
(Hell, 2007), photoactivated localization microscopy (PALM) (Betzig et al., 2006),
and stochastic optical reconstruction microscopy (STORM) (Rust et al., 2006). It is
imperative, therefore, that Ca2þ imaging also joins the super-resolution revolution,
and indeed significant progress has been made on this front with the development of
fluorescence-based techniques such as single channel Ca2þ nanoscale resolution
(SCCaNR) microscopy (Wiltgen et al., 2009). The continued development of both
fluorescent and luminescent GET-CRs will undoubtedly alsomake a contribution to
this advancement, especially if, in the case of the latter, the intensity of the Ca2þ-mediated luminescent emission can be increased. We find ourselves, therefore, at a
most exciting and opportune time to extend our understanding of Ca2þ signaling in
living cells from the microscopic to the nanoscopic level.
Acknowledgments
We thank Philippe Brulet, Marc Knight, and Jean-Rene Martin, who kindly gave us permission to
use their previously published work. Special thanks also to Osamu Shimomura for his generous support
of aequorin-based imaging over the years. We acknowledge financial support from Hong Kong RGC
GRF grants: HKUST-6241/04M,-6416/06M,-661707 and-662109.
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INDEX
1,2-Bis(o-aminophenoxy)ethane-N,N,N’,
N’-tetraacetic acid (BAPTA), 17–18
ionic strength dependent, 5, 7
pH dependent, 4
structural formulas for, 5
temperature, apparent aYnity, 6, 7
2P excitation microscopy
Fluo-based dyes, 248, 249
Rhod-dyes, 249, 250
A
Acetoxymethyl (AM) ester, 120
Aequoria victoria, 154
Aequorins
apoaequorin expression, 268, 277–279
bioluminescent GETCRs, 265
BRET protein complex, 265
coelenterazine, 283–284
expression of GFP-Apoaequorin, 279–283
luminescence
bioluminescence imaging, 287
ca 2þdetection device, 286
detectors types, 267
EMCCD detector, 286, 288
equipment, 284
factors, 290
intensifier-based detector, 290
photon counting photomultiplier, 284
schematic representations, 285
single photon imaging, 290
two-channel luminometer, 286
mitochondrial target, 160
strategies, 265
AM. See Acetoxymethyl (AM) ester
Apoaequorin
BRET complexes
cytosol cell culture, 269–273
diverse range species, 266
materials, 277
methods
in vivo reconstitution, 278
p�-KS-aeq-IRES-EGFP plasmid
preparation, 277–278
transgenic zebrafish generation, 278–279
Azid-1, 33–35
B
Bioluminescence
EMCCD detector, 286
fluorescent GET-CRs, 264
GETCRs, 265
optical systems, 287
single photon detection, 286
vs. EMCCD and an RA-IPD photon imaging
system, 291
Bioluminescence resonance energy transfer
(BRET), 255
aequorins, 265
cytosol cell culture, 269–273
diverse range species, 266
Blue fluorescent protein (BFP), 155
BRET. See Bioluminescence resonance
energy transfer
C
Ca2þ binding compounds, 104, 105
Ca2þ buVers
association constant (KCa), 7, 10
BAPTA family, 5–7
basic steps, solution preparation, 16–18
dissociation constant (Kd), 3
EGTA, 4–6
fluorescent indicators, 3–4
ionic strength corrections, 11–12
measurement, free [Ca2þ], 12–13Michaelis-Menten form, 9
potential complications, 18–19
proton activity coeYcient, 12
software programs, 19–23
spreadsheet for calibration calculations, 13–16
stability constants, 3
temperature corrections, 10–11
Ca2þ flux measurements, 99, 100, 108
301
302 Index
Ca2þ in mitochondria, measurement
cytosolic and mitochondrial Ca2þ, 146Rhod-2 indicator, 143–145
Rhod-2, cytosol, 145–146
Ca2þ, manipulation
buVering changes, 130–132
divalent Cation Ionophores, 129–130
extracellular buVers
BAPTA, 126–127
EGTA, 126–127
lowering Extracellular [Ca2þ], 127–129Ca2þ-selective electrodes
dissociation constant, measurements of, 70
dynamic range of, 69
ETH 129, 86
key advantage, 68
microelectrodes (MEs)
bath calibration, 81
Ca2þ-selective ligand, preparation and use
of, 78–80
calibration procedure, 81–82
double-barreled, 80–81
electrolyte filling of, 78
ETH 1001, ionophore, 82
extracellular [Ca2þ], measurement, 83–85
glass tubing preparation, 76
microelectrode pulling and silanization, 77
solution perfusion, 81
troubleshooting, 85–86
minielectrode
application of, 75–76
electrode potential of, 73
inhomogeneities, eVect of, 75
lifetime of, 72
resistance of, 72
response times of, 73–74
selective ligand, preparation and use, 71–72
storage of, 75
Nicolski-Eisenman equation, 68
Ca2þ-selective liquid membranes, coeVcients, 96,
97
Ca2þselective microelectrodes (CaSMs)
construction
microelectrode, 94
micropipette fabrication, 93
silanization, 93
properties
response time, 97–98
response to Ion Activity, 94–96
selectivity, 96
spatial resolution, 96–97
self referencing
calculation of flux, 103
correction for Ca2þ buVering, 104
diVerential concentration determination,
101–102
diVerential concentration measurement,
98–101
measurement of voltage gradients, 104–105
positional artifacts, 105–107
Caged Ca2þ chelators. See Photolabile Ca2þ
chelators
Calcium release-activated channels (CRAC),
185, 194–195
Cameleon, genetically encoded calcium sensors
CFP/citrine couple, 159
ECFP/EYFP-based cameleons, 161
F46L mutation, 160
FIP-CBsm, 156
FRET, 156
mechanism, 161–163
myosin light chain kinase, 155
YC concentrations, 159
YC2.12 fluorescence, 160
YCX.60, 161
CaSM. See Ca2þselective microelectrodes
CaSMs, construction
microelectrode construction, 94
micropipette fabrication, 93
silanization, 93
CaSMs, properties
Response Time, 97–98
Response to Ion Activity, 94–96
Selectivity, 96
Spatial Resolution, 96–97
CaSMs, self referencing
calculation of flux, 103
correction for Ca2þ buVering, 104
diVerential concentration determination,
101–102
diVerential concentration measurement, 98–101
measurement of voltage gradients, 104–105
positional artifacts, 105–107
Cell-attached patch recordings, 192–193
Coelenterazine, 283–284
Confocal and multiphoton imaging
absorbance/quantum yield, 246–247
advantages and disadvantages, 241–242
calibration of single wavelength dyes, 251–252
conversion of increments, 251
fluorescence lifetime, 247
Fmax values estimation, 252–253
Fmin estimation, 253
Forster resonance energy transfer microscopy
acceptor photobleaching, 236
cameleon, 235, 237
donor quenching, 236
eYciency, 246, 247
Index 303
FRET-FLIM approach, 236
GFP, 235–236
pericams, 237
intrinsic and dye fluorescence, 253–255
laser scanning confocal microscopy
2D frame scan, 231
Jablonski diagram, 232, 233
signal-to-noise, 231
Stokes shift, 232
limitations in speed, 230
multimodal and multiple fluorophore
Di-8-ANEPPS, 256, 257
excitation-contraction coupling, 255
Fura dyes, 258
membrane voltage, 256
overlapping excitation spectra, 256
RH-237 emission spectrum, 257
multiphoton excitation laser scanning
microscopy
2P excitation microscopy, 242, 243
biophysical perspectives, 242
high repetition rates, 245
IR light, 244
NA objectives, 228
parallel scanning confocal systems, 238
programmable matrix microscopy
digital micromirror device, 239
filtering patterns, 239
liquid crystal display, 240
PSF, 229
signal detection system, 227, 228
spatial resolution, 229
spectral shift, 247
spinning disk confocal microscopy, 238–239
total internal reflection fluorescence
microscopy
evanescent wave, 234
refractive index, 233, 234
use of dyes
2P excitation microscopy, 248–250
single-photon confocal microscopy, 248
CRAC. See Calcium release-activated channels
Cytoplasmic [Ca2þ]i, regulation of, 57
D
Diazo compounds, Ca2þ chelators
chemical properties, 39–41
photolysis, eVects of, 41–42
Dibromo-BAPTA (Br2-BAPTA), 4, 11
pH dependent, apparent aYnities, 4
structural formulas for, 5
temperature and ionic strength, apparent Ca2þ
aYnity, 5
Digital micromirror device (DMD), 239
DM-nitrophen, Ca2þ chelators
[Ca2þ]i changes, 37–39absorbance of, 36
Ca2þ-and Mg2þ-aYnities of, 36
caged Mg2þ chelator, 36
kinetic behavior of, 38
quantum eYciency, 35
structure of and reaction scheme, 35
Dual-wavelength ratiometric dyes, 114, 115,
118–120
E
Electrode calibration curves, 15, 16
Electron multiplying charge-coupled devices
(EMCCD)
bioluminescence detection, 286
computer-controlled EMCCD, 289
limitations, 289
single photon events, 289
vs. RA-IPD photon imaging system, 290, 291
EMCCD. See Electron multiplying charge-
coupled devices
Enhanced yellow fluorescent protein (EYFP)
circular permutation, 163–164
Ethylene glycol bis(�-aminoethylether)-N,N,N’,
N’-tetraacetic acid (EGTA), 17–18
ionic strength, apparent aYnity, 6
pH dependent, 4–5
structural formulas for, 5
temperature, apparent aYnity, 6
Excel spreadsheet, Ca2þ calibration buVers, 14, 15
Excitation spectra
Fluo-3, 117
Fura-2, 117
F
Fluorescence lifetime imaging (FLIM), 236
Fluorescent Ca2þ Indicators
dual-wavelength ratiometric dyes, 114, 115,
118–120
single-wavelength nonratiometric dyes, 114,
115, 118–120
Fluorescent Ca2þ Indicators, properties, 115
Forster resonance energy transfer (FRET)
microscopy
acceptor photobleaching, 236
advantage and disadvantage, 236
cameleon, 235, 237
donor quenching, 236
eYciency, 246, 247
304 Index
Forster resonance energy transfer (FRET)
microscopy (cont.)
FRET-FLIM approach, 236
GFP, 235–236
pericams, 237
Fura-2, 33
G
Genetically encoded calcium sensors
Aequoria victoria, 154
Cameleon family
CFP/citrine couple, 159
ECFP/EYFP-based cameleons, 161
F46L mutation, 160
origin, 155, 157
sensor mechanism, 161–163
YC concentrations, 159
YC2.12 fluorescence, 160
YCX.60, 161
yellow fluorescent protein, 159
Camgaroos, 163–164
cases 12 and 16, 167
GCaMPs, 165–167
green fluorescent protein, 154
pericam, 164–165
subcellular locations
endoplasmic reticulum, 168
golgi, 169
mitochondria, 168
peroxisome, 169
plasma membrane, 169
tissue-specific expression
comparative studies, 174–176
GCaMP, 172–173
inverse pericam, 171–172
TN-L15, TN-XL, 173, 174
TN-XXL, 174
YC2.1, 169–170
YC3.3er (citrine-based sensor), 171
uses, 176–177
GFP-apoaequorin
recombinant viral vectors
materials, 281
methods, 281–282
transgenic mice expressing mitochondrially
materials, 282
methods, 283
transient and stable transfection
materials, 279–280
methods, 280
Green fluorescent protein, 154
H
Human cytomregalovirus (HCMV), 279
I
Icrac. See Calcium release-activated currents
Imaging photon detector (IPD), 284
Indicator fluorescence signal, conversion
nonratiometric fluorescent indicator,
calibration, 133–134
ratiometric fluorescent indicator, calibration,
134–138
Indicators, loading in the cells, 121–122
aqueous Solubility of AM Esters, 121
dye compartmentalization
assessing extent of compartmentalization,
122–124
minimizing compartmentalization,
121–122
dye leakage, 124–125
procedure, 124–125
Inositol 1,4,5-trisphosphate receptors (IP3R)
ATP ligands, 198
behavior analysis, 203
Ca 2þ signals, 190
Ca2þ ligands, 198
cell analysis, 195–196
current–voltage relationship, 204
cytopasm-out configuration, 202
DT40 cell expression, 193
electrical recording, 193
endoplasmic reticulum, 190
inner nuclear membrane expression, 198
intrinsic pore open, 190
IP3, 198
nuclear path-clamp recording, 194
single-channel recording, 191, 193
whole-cell recordings, 192
Intracellular Ca2þ
confocal and multiphoton imaging
2P excitation microscopy, 248–250
advantages and disadvantages, 241–242
Fmax values estimation, 252–253
Fmin estimation, 253
FRET, 235–237
indicators, 245–247
intrinsic and dye fluorescence, 253–255
limitations in speed, 230
LSCM, 230–233
multimodal and multiple fluorophore,
255–259
Index 305
multiphoton excitation laser scanning
microscopy, 242–245
parallel scanning confocal systems, 238
programmable matrix microscopy, 239–241
single wavelength dyes, 251–252
single-photon confocal microscopy, 248
spinning disk confocal microscopy, 238–239
TIRF, 233–234
patch clamp methods, 185
Intracellular calcium signals
fluorescent Ca2þ Indicators
Ion channel modulation, Ca2þ chelators, 49–52
calcium channels, 51–52
potassium and nonspecific cation channels,
49–51
J
Jablonski diagram, 232, 233
L
Laser scanning confocal microscopy (LSCM)
2D frame scan, 231
Jablonski diagram, 232, 233
signal-to-noise, 231
Stokes shift, 232
Liquid crystal display (LCD), 240
M
MaxChelator, 22
Microelectrodes (MEs), Ca2þ-selectivebath calibration, 81
Ca2þ-selective ligand, preparation and use of,
78–80
calibration procedure, 81–82
double-barreled, 80–81
electrolyte filling of, 78
ETH 1001, ionophore, 82
extracellular [Ca2þ], measurement, 83–85
glass tubing preparation, 76
microelectrode pulling and silanization, 77
solution perfusion, 81
troubleshooting, 85–86
Minielectrode, Ca2þ-selectiveapplication of, 75–76
electrode potential of, 73
inhomogeneities, eVect of, 75
lifetime of, 72
resistance of, 72
response times of, 73–74
selective ligand, preparation and use, 71–72
storage of, 75
Multiphoton excitation laser scanning
microscopy
2P excitation microscopy, 242, 243
biophysical perspectives, 242
high repetition rates, 245
IR light, 244
Muscle contraction, Ca2þ chelators, 52–53
Myosin light chain kinase (MLCK), 155
N
Nipkow spinning disk, 239, 240
Nitr compounds, Ca2þ chelators
[Ca2þ]i changes, cells, 33–34azid-1, 33
BAPTA, 29, 30
fura-2, 33
nitr-5, 30–31
nitr-7, 30–31
nitr-8, 30–31
Nitr-5, 30–31
Nitr-7, 30–31
Nitr-8, 30–31
Nuclear patch-clamp recording
conventional techniques, 193
ER membrane, 195
IP3R, 195–196
methods
asymmetric recording solutions, 201
cytoplasm-out configuration, 202
DT40 Cell culture, 196–197
equipments, 199
nuclei isolation, 197–198
optimal filtering frequency, 201
pipette tip approaches, 201
recording configuration, 201
single channel record analysis, 202–208
solutions, 198–199
P
Patch clamp methods
calcium-selective channels, 184
intracellular calcium, 185
materials, 195
methods
calcium release-activated currents, 194–195
cell-attached patch recordings, 192–193
fire-polishing pipettes, 191
giga-ohm seals, 190–192
306 Index
Patch clamp methods (cont.)
perforated patch recordings, 193–194
rig assemble, 187–188
Sylgard application, 188, 189
principles, 186–187
recombinant channels, 185, 186
Perforated patch recordings, 193–194
Photolabile Ca2þ chelators, 33
biological applications
cytoplasmic [Ca2þ]i, regulation of, 57
filopodial activity, control of, 58
ion channel modulation, 49–52
muscle contraction, 52–53
rate-limiting steps, 58
synaptic function, 53–57
calibration, 45–48
diazo compounds
chemical properties, 39–41
photolysis, eVects of, 41–42
DM-nitrophen
[Ca2þ]i changes, 37–39absorbance of, 36
Ca2þ-and Mg2þ-aYnities of, 36
caged Mg2þ chelator, 36
kinetic behavior of, 38
quantum eYciency, 35
structure of and reaction scheme, 35
introduction into cells, 42–43
light sources, 43–45
nitr compounds
[Ca2þ]i changes, cells, 33–34azid-1, 33
BAPTA, 29, 30
fura-2, 33
nitr-5, 30–31
nitr-7, 30–31
nitr-8, 30–31
properties of, 32
purity and toxicity, 48–49
Photomultiplier tube (PMT), 227, 231
Photon counting photomultiplier (PMT), 284
PMT. See Photon counting photomultiplier
Point spread function (PSF), 229
Programmable matrix microscopy
digital micromirror device, 239
filtering patterns, 239
liquid crystal display, 240
Proton activity coeYcient, 12
R
Ratiometric fluorescent indicators
Fura-2, 117
Indo-1, 117
principle, 114, 115, 118–120
Recombinant viral vectors
materials, 281
methods, 281–282
S
Scatchard plot analysis, 13, 15, 16
Shuttle buVers, 104, 105
Single channel recording
current-amplitude histograms, 203
dwell-time histogram, 207
electrophysiological records, 202
IP3R cation-selectivity, 206
kinetic analyses, 206
maximum interval likelihood method, 207
Mg 2þ, 198mutations, 205
open probability, 208
Sigworth–Sine transformation, 207
stability plot, 206
Single-photon confocal microscopy, 248
Single-wavelength nonratiometric dyes, 114, 115,
118–120
SOCE. See Store-operated calcium entry
Software programs, Ca2þ buVers, 19–20
accuracy, 21
ideal software criteria, 20
javascript web versions, 22–23
MaxChelator, 22
use and adaptability, ease of, 21
Store-operated calcium entry (SOCE), 185
Synaptic function, Ca2þ chelators, 53–57
T
Tissue-specific expression
comparative studies, 174–176
GCaMP, 172–173
inverse pericam, 171–172
TN-L15, TN-XL, 173, 174
TN-XXL, 174
YC2.1, 169–170
YC3.3er (citrine-based sensor), 171
Total internal reflection fluorescence (TIRF)
microscopy
evanescent wave, 234
refractive index, 233, 234
Y
Yellow fluorescent protein, 159
VOLUMES INSERIES
Founding Series EditorDAVIDM. PRESCOTT
Volume1 (1964)Methods in Cell Physiology
Edited by David M. Prescott
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Volumes in Series 311
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