single-molecule and super-resolution imaging in living cells a dissertation submitted to the

SINGLE-MOLECULE AND SUPER-RESOLUTION IMAGING IN LIVING CELLS

A DISSERTATION

SUBMITTED TO THE DEPARTMENT OF CHEMISTRY

AND THE COMMITTEE ON GRADUATE STUDIES

OF STANFORD UNIVERSITY

IN PARTIAL FULFILLMENT OF THE REQUIREMENTS

FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

Hsiao-lu Lee

December 2011

This dissertation is online at: http://purl.stanford.edu/sg023jd4045

© 2011 by Hsiao-lu Lee. All Rights Reserved.

Re-distributed by Stanford University under license with the author.

ii

http://purl.stanford.edu/sg023jd4045

I certify that I have read this dissertation and that, in my opinion, it is fully adequatein scope and quality as a dissertation for the degree of Doctor of Philosophy.

William Moerner, Primary Adviser


Michael Fayer


Robert Pecora

Approved for the Stanford University Committee on Graduate Studies.

Patricia J. Gumport, Vice Provost Graduate Education

This signature page was generated electronically upon submission of this dissertation in electronic format. An original signed hard copy of the signature page is on file inUniversity Archives.

iii

iv

ABSTRACT

Since the first successful detection single molecules over two decades ago,

single-molecule spectroscopy has developed into a burgeoning field with a wealth of

experiments at room temperature and inside living cells. Probing asynchronous and

heterogeneous populations in situ, one molecule at a time, is not only desirable, but

critical for many biological questions. Further, super-resolution imaging based on

sequential imaging of sparse subsets of single molecules, has seen explosive growth

within the last five years. This dissertation describes both the application of live-cell

single-molecule imaging as an answer to important biological questions, and

development and validation of fluorescent probes for targeted super-resolution

imaging.

Chapter 1 is a general introduction to fluorescence, single-molecule

spectroscopy, live-cell imaging, and super-resolution imaging. In this chapter, single-

molecule experiments in living cells are discussed, and the probes and targeting

schemes used for such experiments are summarized and compared. Chapter 2

describes experimental details of single-molecule imaging in live cells. Chapter 3

presents the application of live-cell single-molecule imaging to studying the

interaction of oligoarginine molecular transporters with cell plasma membranes.

Chapter 4 describes the design, development, and validation of a target-specific

photoactivatable fluorogen (a chromophore that is dark until converted to a fluorescent

form using light) for super-resolution imaging in live mammalian and bacterial

systems. Chapter 5.presents the real-time single-molecule imaging and sub-diffraction

localization of native sodium ion channels in live neuronal models (differentiated

PC12 cells). This experiment utilized novel fluorescent saxitoxins created by de novo

synthesis.

Data, figures, tables, and excerpts are used with permission in this dissertation from

the following publications:

v

(1) Ondrus, A.*; Lee, H.-L.D.*; Iwanaga, S.; Parson, W.; Andresen, B.; Moerner,

W.E.; DuBois, J. Fluorescent saxitoxins for live cell imaging of single voltage-

gated sodium ion channels beyond the optical diffraction limit. In prep.

(2) Lee, H.-L.D.; Lord, S. J.; Iwanaga, S.; Zhan, K.; Xie, H.; Williams, J. C.; Wang,

H.; Bowman, G. R.; Goley, E. D.; Shapiro, L.; Twieg, R. J.; Rao, J.; Moerner, W.

E. Superresolution Imaging of Targeted Proteins in Fixed and Living Cells Using

Photoactivatable Organic Fluorophores. J. Am. Chem. Soc. 2010, 132, 15099-

15101.

(3) Lord, S. J.; Lee, H.-L.D.; Moerner, W. E. Single-Molecule Spectroscopy and

Imaging of Biomolecules in Living Cells. Anal. Chem. 2010, 82, 2192-2203.

(4) Lord, S. J.; Lee, H.-L.D.; Samuel, R.; Weber, R.; Liu, N.; Conley, N. R.;

Thompson, M. A.; Twieg, R. J.; Moerner, W. E. Azido Push–Pull Fluorogens

Photoactivate to Produce Bright Fluorescent Labels. J. Phys. Chem. B 2010, 114,

14157-14167.

(5) Lee, H.-L .; Dubikovskaya, E. A.; Hwang, H.; Semyonov, A. N.; Wang, H.; Jones,

L. R.; Twieg, R. J.; Moerner, W. E.; Wender, P. A. Single-Molecule Motions of

Oligoarginine Transporter Conjugates on the Plasma Membrane of Chinese

Hamster Ovary Cells. J. Am. Chem. Soc. 2008, 130, 9364-9370.

(6) Lord, S. J.; Conley, N. R.; Lee, H. -L. D.; Samuel, R.; Liu, N.; Twieg, R. J.;

Moerner, W. E. A Photoactivatable Push−Pull Fluorophore for Single-Molecule

Imaging in Live Cells. J. Am. Chem. Soc. 2008, 130, 9204-9205.

vi

ACKNOWLEDGMENTS

I have been very fortunate to have W.E. has a research advisor. Through his

insights, support, and well-timed critique, he has taught me how good science is done

and communicated. Further, the joy and enthusiasm he enjoys for science is extremely

contagious and motivational, which is particularly important for difficult times. I could

not have asked for a better advisor.

The members of the Moerner group have contributed immensely to my

personal and professional development at Stanford. The group has been a source of

friendships as well as good advice and collaboration. Dr. Hanshin Hwang introduced

me to cell culture, a feat considering prior to graduate school all my experiments were

performed in a UHV chamber. Dr. So Yoen Kim whose thoroughness and good cheer

made her not only a great office mate, but also the go-to person for advice during my

first 2 years of graduate school. I have had the pleasure of working with Dr. Sam Lord

and Dr. Nick Conley, to whom I have turned many times for chemistry advice and

experimental discussion. Dr. Julie Biteen was generous and helpful to me in a number

of ways from helping me trouble-shoot bacterial experiments to fitting single point

spread functions. Dr. Steven Lee was helpful in many discussions on sub-diffraction

re-constructions. Whitney Duim helped me through learning many biochemical

methods, and has been a great source of support, and friendship. Matthew Lew has

been kind and generous with his expertise on MATLab and optics. Lana Lau has also

been a great source of support and awesome sounding board for crazy experimental

ideas. I also thank Marissa Lee for helpful fluorophores photophysics discussions.

My time at Stanford was made enjoyable in large part by the many friends that

have become a part of my life. I am grateful for time spent with roommates and

friends: Nicholas J. Ward, Xiao Li, Yale Huang, Tina Xu, Marissa Caldwell, Michael

Stern, Meredith Foster, Lydia Ho, Alisha Weight, and Harry Jao. I am also grateful to

my Wellesley College Professors for continuing to be supportive of me even though I

have graduated. My time at Stanford was also enriched by the warm communities of

dancers in the bay area.

vii

I would like to thank my family for their unyielding love and encouragement.

My aunt Leena , uncle TC, cousins Arthur and Vicky have made the bay area much

more of a home for me. My sister Raechel has always been there for me in celebration

and in distress. She would spend weekends and nights with me in lab. I would also

like to thank my sister Ivy for holding down the fort in Kaohsiung while Raechel and I

pursue our studies abroad. And most of all, I would like to thank my parents whose

love and sacrifice gave me the opportunity to pursue science; my mother Ching-Mei

has been an unfaltering tower of support in my life, and my father, Jueen, whose

curiosity and intellect has inspired my studies in science. This dissertation is dedicated

to them.

viii

TABLE OF CONTENT

Abstract .............................................................................................................................. iv

Acknowledgments .............................................................................................................. vi

Table of Content .............................................................................................................. viii

List of Tables ...................................................................................................................... xi

List of Figures .................................................................................................................. xii

1. Introduction and Background .................................................................................... 1

Fluorescence spectroscopy of molecules .................................................. 3 1.1

Single-molecule experiments in biological cells ....................................... 6 1.2 Sample requirements for single-molecule detection ............................................ 6 1.2.1

Additional requirements for imaging in live cells ............................................... 8 1.2.2

Super-resolution in live cells ............................................................................. 19 1.2.3

Single-molecule imaging in cells: a timeline..................................................... 25 1.2.4

Outlook ..................................................................................................... 28 1.3

Scope of the dissertation .......................................................................... 28 1.4

References ................................................................................................. 29 1.5

2. Experimental ............................................................................................................. 44

Microscopy Instrumentation ................................................................... 46 2.1 Microscope Configurations ............................................................................... 46 2.1.1

Detectors ............................................................................................................ 49 2.1.2

Optics ................................................................................................................ 51 2.1.3

Excitation Light Source ..................................................................................... 52 2.1.4

Cell Culture .............................................................................................. 53 2.2 Bacterial Cell Culture ........................................................................................ 53 2.2.1

Mammalian Cell Culture ................................................................................... 64 2.2.2

Immunofluorescence ................................................................................ 78 2.3 Immunofluorescence protocol ........................................................................... 81 2.3.1

Antibody conjugation ........................................................................................ 82 2.3.2

Total protein isolation from cells/cell lysis ............................................. 84 2.4 Total protein isolation from bacterial cells ........................................................ 84 2.4.1

Total protein isolation from mammalian cells ................................................... 85 2.4.2

Coverslip coating ...................................................................................... 86 2.5 Fibronectin surface coating ............................................................................... 86 2.5.1

Collagen surface coating ................................................................................... 86 2.5.2

Polyelectrolyte multilayer surface coating ........................................................ 87 2.5.3

References ................................................................................................. 87 2.6

3. Single-Molecule Motions of Oligoarginine Transporter Conjugates on the

Plasma Membrane of Chinese Hamster Ovary Cells ...................................................... 95

ix

Introduction to cell penetrating peptides ............................................... 98 3.1

Experimental .......................................................................................... 107 3.2 Fluorescent Conjugates ................................................................................... 107 3.2.1

Epi-illumination for single-molecule imaging ................................................. 108 3.2.2

Cellular sample preparation ............................................................................. 109 3.2.3

Imaging Arginine8-D-V tethered to polyelectrolyte multi-layers ................... 110 3.2.4

Passive diffusion inquiry ................................................................................. 110 3.2.5

Adaptive translocation inquiry ........................................................................ 111 3.2.6

Receptor-mediated endocytosis inquiry .......................................................... 111 3.2.7

Macropinocytosis inquiry ................................................................................ 112 3.2.8

High octaarginine concentration inquiry ......................................................... 113 3.2.9

Effects of addition of oxygen scavenger on imaging ...................................... 114 3.2.10

Single-molecule fluorescence imaging of positional trajectories ....... 115 3.3

Analysis of single-molecule motion ....................................................... 116 3.4

Results ..................................................................................................... 117 3.5 Fluorescently-labeled Arg8 conjugates are readily internalized ...................... 118 3.5.1

Single Arg8-D-V molecules disappear before photobleaching ....................... 120 3.5.2

Length of single-molecule trajectories yields residence time .......................... 122 3.5.3

Diffusion coefficients of single-molecule trajectories ..................................... 124 3.5.4

Conclusions ............................................................................................. 131 3.6

3.7 Acknowledgements ................................................................................ 132

References ............................................................................................... 133 3.8

4. Super-resolution Imaging of Targeted Proteins in Fixed and Living Cells

Using Photoactivatable Organic Fluorophores ............................................................. 139

4.1. Introduction ............................................................................................ 141 Requirements for pointillist super-resolution imaging .................................... 141 4.1.1

Small-molecule photoactivatable fluorophores: the azido-DCDHFs .............. 145 4.1.2

Cellular labeling considerations ...................................................................... 152 4.1.3

Our cellular labeling strategy: HaloTag/HaloEnzyme Enzymatic Targeting . 154 4.1.4

Experimental .......................................................................................... 157 4.2 Plasmids for HaloEnz Fusion Construction ..................................................... 157 4.2.1

Cell Labeling Protocol ..................................................................................... 159 4.2.2

Cell Imaging Protocol ..................................................................................... 161 4.2.3

Photoactivation Quantum Yield Characterization ........................................... 163 4.2.4

Results ..................................................................................................... 164 4.3 Photophysical characterization of Azido DCDHF HaloTag ............................ 164 4.3.1

Specificity and efficacy in in mammalian culture ........................................... 166 4.3.2

Specificity and efficacy in live bacterial samples ............................................ 168 4.3.3

Acknowledgements ................................................................................ 171 4.4

References ............................................................................................... 171 4.5

5. Fluorescent Saxitoxins for Super-Resolution Imaging of Voltage-Gated

Sodium Ion Channels on Live Neuronal Cells .............................................................. 179

Introduction ............................................................................................ 181 5.1 Cell membrane potential and action potential ................................................. 181 5.1.1

Sodium ion channels ....................................................................................... 184 5.1.2

x

Saxitoxin .......................................................................................................... 185 5.1.3

Experimental .......................................................................................... 187 5.2 Synthetic design of fluorescent STX ............................................................... 187 5.2.1

Cell culture ...................................................................................................... 188 5.2.2

Electrophysiology experiments ....................................................................... 190 5.2.3

Confocal microscopy on CHO and PC12 cells ................................................ 191 5.2.4

Single-molecule microscopy ........................................................................... 196 5.2.5

Wide-field image and data analysis ................................................................. 197 5.2.6

Results and discussion ........................................................................... 202 5.3 Photo-physical properties of STX-Cy5 and STX-DCDHF ............................. 202 5.3.1

Fluorescent saxitoxins bind specifically to NaV .............................................. 204 5.3.2

NaV block by STX, STX-Cy5 and STX-DCDHF in differentiated PC12 ....... 206 5.3.3

Reversible, specific NaV labeling by STX-Cy5 and STX-DCDHF ................. 209 5.3.4

NaV expression and distribution in NGF-differentiated PC12 ......................... 211 5.3.5

Single-molecule and super-resolution imaging ............................................... 212 5.3.6

Single-molecule tracking of labeled NaVs in NGF-differentiated PC12 ......... 213 5.3.7

Single-molecule imaging of NaV distributions on filopodia ............................ 216 5.3.8

Super-resolution imaging of labeled NaVs in filopodia and neuritic spines .... 218 5.3.9

Conclusion .............................................................................................. 220 5.4

Acknowledgement .................................................................................. 221 5.5

References ............................................................................................... 221 5.6

xi

LIST OF TABLES

Table 1-1 Fluorescent emitters for single-molecule imaging in living cells. .............. 10 Table 1-2 Various tag-probe labeling methods. .......................................................... 16 Table 1-3 Selected single-molecule experiments with relevance to cell imaging. ..... 25

Table 2-1 Important timeline in tissue culture technique development ....................... 64

Table 2-2 Commonly used continuous cell lines for microscopy. ............................... 65

Table 3-1 Amino acid sequences of a sampling of arginine-rich cell penetrating

peptides ....................................................................................................................... 100

Table 4-1 Photophysical properties of various photo-switchable molecules ............ 149

Table 4-2 Photophysical parameters of Azido DCDHF-V and derivatives ............... 164

xii

LIST OF FIGURES

FIG. Description Page

1-1 A Jablonski diagram 4

1-2 Potential energy diagram representing molecular vibronic transitions 6

1-3 Excitation (A) and emission spectra (B) of the principal endogenous

cellular fluorophores that contribute to autofluorescence.

9

1-4 Over expression of actin-GFP in a NIH3T3 cell 12

1-5 Super-localization of a single fluorescent molecule 21

1-6 Schematic showing the key idea of super-resolution imaging of a

structure by single-molecule super-localization and active control.

23

1-7 The importance of the Nyquist-Shannon criterion in interpreting

super-resolution structures

25

2-1 General microscope configurations for single-molecule imaging in

biological samples

47

2-2 Illustration of the gain stages and ADC conversion of photons into

counts by the EMCCD camera

50

2-3 General scheme for bacterial culture cultivation. 55

2-4 A typical bacterial culture growth curve 56

2-5 Schematic of inoculating an agarose plate for bacterial culture. See

the following steps for streaking sequence

57

2-6 Schematic for preparing bateria containing agarose gel pads on over

slips for imaging

60

2-7 Hemacytometer 71

2-8 Schematic for immunofluorescence experiments 78

3-1 Mechanisms of peptide uptake across the cellular membrane 102

3-2 Representative associations of a polycationic guanidinium

transporter with anionic cell membrane constituents

104

3-3 A. Structures of DCDHF-V labeled octaarginine (Arg8-D-V) and

tetraarginine (Arg4-D-V) B. Structure of DCDHF-V labeled lipid

analog (D-V-12)

108

3-4 Schematic of the imaging arrangement. 109

3-5 Effect of the enzymatic oxygen scavenger system for single

conjugate cellular imaging showing high contrast from the DCDHF

label at normal oxygen

115

3-6 Representative images of Arg8-D-V on the CHO plasma membrane

with decreasing concentration of conjugate (A B, and C).

118

3-7 Comparison of photo-bleaching time and residence time. 122

3-8 Residence time distributions and fitted residence times of Arg8-D-V,

Arg8-D-V on high [K+] treated CHO cells, Arg4-D-V, D-V-12,

Transferrin-Alexa594, and Arg8-D-V on cytochalasin D treated

CHO cells.

123

3-9 Distributions of observed single-molecule diffusion coefficients (D) 125

xiii

for A: Arg8-D-V, B: D-V-12, C: transferrin-Alexa594 conjugate,

and D: Arg8-D-V in high [K+] on the CHO plasma membrane

3-10 Normalized probability distributions for Arg8-D-V (black) and

Arg8-D-V on high [K+] buffer treated CHO cells (gray)

129

3-11 Normalized probability distributions for Arg8-D-V (red) and Arg8-

D-V on cytochalasin D treated CHO cells (purple)

130

3-12 Distributions of single-molecule diffusion coefficients and residence

times. On the left, distribution of single-molecule diffusion

coefficients (D) of Arg8-D-V in presence of 10 µM of excess

unlabeled Arg8 peptides

131

4-1 Schematic showing the key ideas of super-resolution imaging of a

structure by photo-control

144

4-2 Schematic structure of the DCDHF fluorophores. The amine donor

and DCDHF acceptor are connected by a π-conjugated linker.

146

4-3 Photo-conversion of dark azido fluorogens produce fluorescent

emitter

147

4-4 Chemical structure and spectra of an azido DCDHF-V photo-

activatable flurogen.

148

4-5 Demonstration of the photoactivation of azido DCDHF-V in live

cells

151

4-6 Single molecule trajectories of DCDHF-V-P-amine fluorophore

diffusing in the membrane of a CHO. The trajectory of a single copy

of the DCDHF-V-P-amine fluorophore diffusing in the membrane of

a CHO cell after photoactivation. Dotted red lines indicate when the

fluorophore was dark (i.e. blinking).

154

4-7 Schematic of the specific conjugation mediated by the HaloEnz

protein and its HaloTag ligand

155

4-8 Photochemical activation of azido DCDHF 155

4-9 A) Flow chart of experimental procedures in using HaloTag-

DCDHF-fluorogen. (B) HaloTag-DCDHF fluorogen forms covalent

bond with HaloEnzyme.

156

4-10 Effects of diffuse white light on Azido DCDHF HaloTag. 165

4-11 Evidence that the HaloTag-targeted fluorogen correctly labels

specific proteins and enables SR imaging in mammalian cells

167

4-12 Negative bacterial controls 169

4-13 Diffraction-limited imaging of DCDHF HaloTag inside live C.

crescentus cells expressing fusion proteins to FtsZ and PopZ

169

4-14 SR imaging of protein fusions inside live C. crescentus cells using

azido-DCDHF HaloTag

170

5-1 Action potential as a plot of membrane potential vs. time 182

5-2 Computational model of STX bound in the sodium channel pore,

side view.

186

5-3 Natural and synthetic saxitoxins for NaV studies. 188

5-4 Natural and synthetic saxitoxins for NaV imaging 188

xiv

5-5 Sequence comparison of the pore region in rNaV1.4, Na=1.7 (PC12),

and NaV1.2 (PC12)

195

5-6 Schematic for maxplot stacking of a set of images 197

5-7 Fluorescence intensities over time of STX-Cy5 on NaVs in the

membrane of NGF differentiated PC12 cells and surface

immobilized Cy5-NHS

199

5-8 Statistical localization precision for STX-Cy5 on PC12. 200

5-9 Absorbance/emission spectra of STX-Cy5 and STX-DCDHF. 203

5-10 Fluorescence enhancement of membrane-bound STX-DCDHF 204

5-11 Confocal images of specific rNaV1.4 labeling by STX-fluorophores

in CHO cells

205

5-12 Single-molecule wide-field images of specific rNaV1.4 labeling by

STX-fluorophores in CHO cells

206

5-13 Sodium ion channel current blocking by natural and synthetic

saxitoxins

207

5-14 On/off times and rate constants for Na+ current block by STX 208

5-15 Extracellular labeling of NGF differentiated PC12 cells by STX-

DCDHF

209

5-16 Live cell confocal imaging of NaVs. NaVs were labeled with 10 nM

STX-Cy5 or 15 nM STX-DCDHF

209

5-17 Quantification of probe wash-off and PC12 ± NGF fluorescence.

Media contained 10 nM STX-Cy5 or 15 nM STX-DCDHF

210

5-18 Quantification of probe wash off and PC12 ± NGF fluorescence.

Media contained 10 nM STX-Cy5 or 15 nM STX-DCDHF

212

5-19 Single-particle tracking of NaVs in the soma and neurites of an NGF

differentiated PC12 cell

214

5-20 Single-molecule imaging of NaVs in filopodia of an NGF


216

5-21 Maxplot images of NaVs labeled with STX-Cy5 and stretch rates in

filopodia of an NGF differentiated PC12 cell

217

5-22 Super-resolution imaging of NaVs in neuritic spines of an NGF


219

1

1.INTRODUCTION AND BACKGROUND

2

Abstract

In this dissertation, I focus on single-molecule spectroscopy and imaging

experiments that measure the signal from each individual fluorescent label in living

cells. While the related area of fluorescence-correlation spectroscopy1 has also been

applied to living cells,2 it will not be discussed in this dissertation. In this chapter, I

include a brief overview of fluorescence, key requirements for single-molecule

imaging, additional requirement for imaging in live cells, and applications of single-

molecule imaging, including pointillist super-resolution methods. This chapter does

not attempt to be an exhaustive review of all the relevant topics; rather, emphasis is

placed on capturing relevant theoretical background and recent progress. Lastly, this

chapter refers the reader to relevant (recent) reviews and background reading in each

section.

3

Fluorescence spectroscopy of molecules 1.1

Modern optical fluorescence microscopy relies on using targeted emitters to

label analytes of interest while providing high contrast over background. Good

fluorescent emitters in carefully prepared samples provide excellent signal-to-

background ratio and are available with a wide range of properties (e.g.

environmental-sensitivity), enabling molecule-differentiating read-out. All projects

included in this dissertation essentially rely on the use of fluorescent emitters.

Therefore, a short theoretical overview of the fluorescence process based on the

references cited will be given in this section.

Spectroscopic experiments study how light interacts with matter as a function

of wavelength/radiative energy. When light impinges upon a molecule, the oscillating

electromagnetic field from the light can couple the ground-state molecule with higher

energy states; if the appropriate wavelength of light is used, this may bring the

molecule to a higher excited state. Most molecules of interest in this thesis have

singlet ground states and we are interested in visible light, so the usual allowed

electronic transition is from the lowest singlet to the first excited singlet S0S1. The

excited molecule can then relax to the ground state radiatively, via either fluorescence

(S1S0, with no change of spin) or phosphorescence from the lowest triplet state

(T1S0), or non-radiatively by internal conversion or intersystem crossing. Figure 1-

1 depicts the energy levels and transitions that are typically relevant in fluorescence

microscopy.

4

Figure 1-1 A Jablonski diagram depicting molecular energy levels and transitions

with the states defined in the text. Solid arrows represent absorptive or emissive vertical (Franck–Condon) transitions;

squiggles represent non-radiative transitions. VR is vibrational relaxation; ISC is

intersystem crossing from singlet to triplet state. The approximate timescales are as

follows: absorption and fluorescence transitions are practically instantaneous (< fs);

VR occurs over ps; average times in the excited electronic state before fluorescence

emission are ns; ISC is a spin-forbidden transition so phosphorescence is typically

slow (s–s or longer); for typical wide-field single-molecule imaging, excitation times

are on the order of one photon absorbed per 10 s. Figure courtesy of Sam Lord.

The low irradiance condition of most laboratory fluorescence spectroscopy

enables us to treat light classically as an electromagnetic wave and treat the molecule

quantum-mechanically (i.e. in a semi-classical approach). In this manner, one can

calculate the time-dependent probability of the radiative transition using time-

dependent perturbation theory. This calculation shows that the probability that the

molecule would reach excited electronic state scales linearly with the absolute square

of the transition dipole moment, the irradiance of the on-resonance light, and the

duration of time that the target molecule is exposed to the light.

The one-dimensional transition dipole moment can be expressed as

0

0S

0

1S i

ixe , where i

ixe represents the sum of the x coordinates of electrons in the

molecule multiplied by the charge e, and the two 0 are the time-independent

wavefunctions of the S0 ground and the S1 excited states. This transition dipole

moment can be related to the classical oscillator strength, the Einstein coefficients, and

transition selection rules.

5

Figure 1-2 depicts the potential-energy surfaces of the ground and excited

states of a molecule, and the vibronic (i.e. electronic and vibrational) transitions

between the two surfaces. The Born-Oppenheimer approximation assumes that the

electronic and nuclear wavefunctions can be treated independently because electrons

move and rearrange on a time scale much faster than the nuclei. This is depicted by the

vertical arrows in Figure 1-2, which indicate that, while there is a change in the energy

of the system, there is no change in the nuclear configuration during an electronic

transition and therefore no change in the molecular structure. In other words,

electronic transitions are regarded as essentially instantaneous compared to the

timescale of nuclear motions. This result forms the basis for the Frank-Condon

analysis. The Franck–Condon principle states that, upon near-instantaneous excitation,

the excited molecule retains its exact ground-state structure, to the transition is vertical

with no change in nuclear coordinates. The resulting excited state of the molecule is

termed the Frank-Condon state. The excited molecule can then relax from the Franck–

Condon state to other lower energy excited-state structures via vibrational relaxation

and nuclear rearrangement.

The transition probability between the Franck–Condon state and the ground

state is determined by the overlap integral between vibrational wavefunctions in the

two electronic states. There are 3 major practical results stemming directly from this

principle: 1) Molecules are more likely to absorb higher-energy light (bluer) and emit

photons with lower energy (redder); 2) Molecular absorption and emission spectra are

often mirror-images of each other; 3) For rigid aromatic molecules such as

naphthalene or anthracene, where the vibronic bands in the spectra can be resolved at

room temperature in solution, the Franck–Condon principle also explains the relative

intensities of the different vibronic transitions.

6

Figure 1-2 Potential energy diagram representing molecular vibronic transitions.

Vertical transitions obey the Franck–Condon principle. The relative probability of an

electronic transition between two states depends on the overlap of the vibrational

wavefunctions in the ground and excited state (the orange and green wavefunctions),

i.e. the Franck–Condon factor.

For more information on molecular photophysics, please reference the following:

Fayer, M. D. Elements of Quantum Mechanics; Oxford University Press: New York,

2001.

Lakowicz, J. R. Principles of Fluorescence Spectroscopy; Springer: New York, 2006.

Valeur, B. Molecular Fluorescence; Wiley: New York, 2002.

Single-molecule experiments in biological cells 1.2

Owing to more than 20 years of active research and development (extensively

reviewed in several publications),3-6

it is now possible to routinely image single

molecules in microscopy. In the following, I highlight some key requirements for

achieving single-molecule imaging, and discuss additional considerations for imaging

single molecules in live cells.

Sample requirements for single-molecule detection 1.2.1

Achieving the required signal-to-noise ratio (SNR) for single-molecule

fluorescence measurements requires maximizing signal while minimizing background.

Therefore, fluorescent single-molecule imaging traditionally requires the following: 1)

7

a transparent, non-fluorescent host matrix; 2) fluorescent molecules that are resolved

by separating them in space (by more than the diffraction limit of ~200 nm), time, or

spectral properties; 3) good fluorescent probes that exhibit the combination of a large

absorption cross section, high photo-stability, and weak bottlenecks into dark states

such as triplet states. In addition, one must rigorously exclude undesirable fluorescent

impurities, minimize Raman scattering, and reject any Rayleigh scattered radiation at

the pumping wavelength.3

These aforementioned parameters directly contribute to the SNR which must

be greater than one for a relevant acquisition/averaging time. The SNR for detecting

one molecule in fluorescence spectroscopy is:7

dλbλλ

λλ

em

em

RDIRDhc

Iσ

TDhc

Iσ

BTDTR

DTR

λ

λ

noise rms

signal

F

F

Equation 1-1

where Rem is the photon emission rate of the single fluorophore, ΦF is the fluorescence

quantum yield of the fluorophore, σλ is the fluorophore absorption cross-section at the

excitation wavelength λ, T is the detector counting interval (or binning time), Iλ is the

excitation irradiance at the sample, h is Planck’s constant, c is the speed of light, B is

the overall measured background count rate, Rb the irradiance-dependent background

count rate from the sample, Rd is the dark-count rate from the detector, and D is the

instrument-dependent collection factor which includes losses due to the microscope

optics and filters as well as the detector quantum efficiency.7

The Poisson statistics representing the photon stream from a single emitter

results in the SNR improving with the square root of the binning time as long as dark

counts are not dominant. The two processes which limit SNR are background and

shot-noise. In the preferred shot-noise limited regime, the experimental SNR increases

as the square root of the irradiance as long as saturation of the emitting molecule is not

occurring. Fluorescence microscopy in living cells is generally in the background-

limited regime.

8

Please see Ref.3, Ref.4, and Ref.7 for more in-depth discussion.

Additional requirements for imaging in live cells 1.2.2

Single-molecule imaging in live cells offers insight into physiological

processes with molecular detail, but presents a number of methodological challenges.

Imaging single emitters in living cells introduces additional challenges including those

mentioned in 1.2.1.: 1) cellular autofluorescence, which can significantly lower the

SNR; 2) the need for a bio-orthogonal labeling strategy; 3) specificity of the

fluorescent probe to the analyte of interest. In the following, we will examine these

three considerations in depth.

Cellular autofluorescence 1.2.2.1

In order to maintain low background counts, one must take precaution to avoid

cellular auto-fluorescence. The term ‘‘autofluorescence’’ is used to distinguish the

intrinsic fluorescence of cells/tissues from the extrinsic fluorescence that results from

the exogenous fluorescent probes that are used to label cell and tissue structures.8

When excited with radiation of a suitable wavelength, some cell and tissue molecular

components behave as endogenous fluorophores: they fluoresce and contribute

background. These endogenous fluorophores can be found distributed throughout the

cell, such as emission from proteins that contain aromatic amino acids (e.g.

tryptophan, tyrosine, phenylalanine), as well as critical cellular cofactors such as

nicotinamide adenine dinucleotide (NADPH), flavins, and lipopigments.8 Flavins are

active players in cellular metabolic processes and emit in the visible spectrum.9 NADP

is a major electron acceptor and its reduced form, NADPH, is a coenzyme used as a

source of reducing power in many cellular reactions. NADPH has an excitation

maximum at 340 nm and emission maximum at approximately 450 nm.8 Plant cell

samples also contain many additional fluorophores which can produce background,

such as chlorophylls and flavonoids.10

Figure 1-3 from Wagnieres et al. presents the

excitation and emission spectra of these aforementioned endogenous cellular

fluorophores.11

Cellular autofluorescence can be avoided by selecting an imaging

9

wavelength longer than about 500 nm, where biological fluorophores typically do not

absorb, and by using cell growth media and imaging buffers that are free of

fluorophores.8

Figure 1-3 Excitation (A) and emission spectra (B) of the principal endogenous

cellular fluorophores that contribute to autofluorescence. The best relative

excitation/emission conditions have been considered. Derived from Wagnie`res et

al.11

Choosing the right emitter 1.2.2.2

As mentioned in 1.2.2, imaging single fluorescent emitters in living cells

introduces further challenges in addition to those that are present in other single-

molecule experiments. Fortunately, by making careful choices in sample-labeling

technique, sample preparation, and imaging conditions, one can extend the capabilities

of single-molecule imaging to live cells. One of the first considerations in preparing

live-cell samples for single-molecule imaging is choosing the right fluorescent emitter

to label the cellular construct-of-interest. Table1-1 compares several classes of

existing emitters for this purpose. In all cases, careful control experiments are

necessary to ensure that the emitter and labeling technique used do not alter the

physiology-of-interest. The following sections will briefly review quantum dots,

fluorescent proteins, and small-molecule emitters.

10

Table 1-1 Fluorescent emitters for single-molecule imaging in living cells.

Emitter Advantage Disadvantage

Organic

Fluorophores

Small molecular size

Synthetic flexibility

High photostability

10E6 photons emitted per

molecule)

Targeting Scheme required, see

1.2.2.2.3. for existing strategies

QDs and

Nanoparticles

Excellent source of

photons(>10E7 photons emitted per

particle)

Large

Cell delivery currently only

possible via microinjection

Targeting Scheme required, see

1.2.2.2.3. for existing strategies

Blinky

FPs Target specificity Less photostable (only

~100,000 photons emitted per molecule)

Fluorescent proteins 1.2.2.2.1

Compared to other existing options, fluorescent proteins (FPs) have the noted

advantage of being genetically-targetable. However, the major drawback to FPs is that

they are generally an order of magnitude less photostable than most small organic

fluorophores, which can emit millions of photons before photobleaching.12, 13

The

original green fluorescent protein (GFP) from the Aequorea victoria jellyfish is a

protein with a length of 238 residues, that covalently creates a fluorophore by

rearrangement and oxidation of Ser, Tyr, and Gly residues in the core of the 11-

stranded β-barrel upon folding and maturation.14, 15

Live-cell imaging took a step

forward following the introduction of the improved GFP that carries the point

mutation (S65T), in 1995 by Tsien et al.16

This point mutation red-shifted GFP λex,max

by ~90 nm, avoiding much of the cellular autofluorescence, and improved upon its

photostability. Since then, FPs have improved our ability to study protein function

directly in vivo by enabling individual proteins to be selectively labeled via genetic

fusion and expression of recombinant proteins.17

11

Since 1995, FPs have seen much optimization via techniques such as site-

directed mutagenesis to provide a well-stocked toolkit for cell imagers. In addition to

the S65Y mutation, other important mutations include: 1) S30R, V163A, and N149K

led to a significantly faster folding rate;18-20

2) Q69M improved photostability and

resistance to Cl- and low pH;

21 3) I167T reduced thermal-sensitivity.

22 Further, in

addition to the FPs derived from jellyfish, FPs from corals and anemones have also

been developed and studied.

Many recent single-molecule-based super-resolution imaging experiments (see

section 1.3.) have relied on photoactivatable fluorescent proteins (PA-FPs). PA-FPs

are FPs that “switch” to a new fluorescent state as a result of some stimulus/activation.

Over the last decade, more than 20 different PA-FPs have been developed to enable

both super-resolution imaging and pulse-chase experiments.23

To this date, PA-FPs

have enabled pulse-chase tracking of cellular organelles, 24-2624

tracking of cells in

tissues and embryos,27

and super-resolution tracking/imaging experiments in 2D and

3D.28, 29

For a comprehensive review please see references.14, 15

Experimentally, using the PA-FPs requires an additional ~400nm activation

laser other than the read-out imaging laser. PA-FPs belong to one of the three

following mechanistic categories: 1) irreversible conversion from a dark state to a

bright fluorescent state; 2) irreversible conversion from one fluorescent color to

another color (usually red-shift); 3) reversible conversion to a bright state with the

possibility of being converted back to the bright state again once photobleached. The

first two categories are generally termed “photoactivatable”, and the last category is

generally referred to as “photoswitchable”. The very first example of this

“photoswitching” behavior was observed in the enhanced yellow fluorescent protein,

eYFP, by Dickson et al. 30

Under continuous excitation at 514 nm, single eYFP

molecules first reversibly blink as expected, and then enter a transient dark state. Upon

re-activation using a pulse of 407nm light, the molecules can then be recovered from

the transient dark state. This photoswitching process can be repeated several times for

each molecule. This first demonstration of FP photoswitching illustrated the unique

ability of single-molecule spectroscopy to observe molecular behavior that had been

12

previously been obscured by ensemble measurements. In addition, this photoswitching

behavior of eYFP was later used to image the cytoskeletal protein MreB in supe-

resolution imaging (~40 nm)31

and the nucleoid-associated protein HU in Caulobacter

crescentus.32

Compared to choosing a regular FP, the process of choosing a PA-FP for an

experiment should include additional considerations of the following factors: 1)

brightness level after photo-conversion; 2) contrast ratio/on-to-off ratio (see sections

1.3.1.2-1.3.1.3 for more details); 3) likelihood of undesired spontaneous conversion to

the bright state; 4) photo-activation/switching cross-section. PA-FPs with high photo-

conversion cross-section convert readily and are more desirable as this reduces the

amount of UV/blue light to which the cellular sample is exposed. On the other hand,

the use of extremely sensitive PA-FPs may incur the additional complication of having

to prepare samples in the dark prior to imaging.

In addition to the stated uses and advantages, FPs and PA-FPs both exhibit

some limitations for use in more sensitive biophysical and mechanistic experiments.

As 30 kD proteins, (PA-)FPs may, especially at high expression levels, disrupt

cellular morphology and the assembly, interaction, or function of the proteins-of-

interest. Therefore, determining the optimal expression level of fusion proteins to both

achieve high density of labeling and to preserve cellular morphology is very

important. See Figure 1-4 for an example of changes in cellular morphology as a result

of over-expression of FP fusion.

Figure 1-4 Over expression of actin-GFP in a NIH3T3 cell results in severe

membrane blebbing. Data taken by Hsiao-lu Lee, during Bio-X Tissue Culture

workshop 2007.

13

Other limitations of FPs include their tendency for oligomerization (e.g. A.

victoria GFP is part of a heterotetrameric complex, DsRed is an obligate tetramer),

possible slow folding/maturation depending on the variant, and relative inaccessibility

for rational design and modification. Further, of import to sub-diffraction experiments,

the total number of photons emittable from FPs is generally about one-tenth that of

organic fluorophores. Other imaging probes including nanoparticles, quantum dot, and

NV-centers in diamond also exhibit much more favorable photophysical properties.

Therefore, chemistry-based tagging schemes have been developed to provide

alternatives for labeling proteins of interest with favorable probes directly in living

cells (see section 1.2.2.3.). In sum, while FPs will likely continue to be used widely,

complementary experiments with better emitters should also be used to achieve better

resolution.

Quantum dots 1.2.2.2.2

First discovered in the early 1980s, quantum dots (QDs) are semiconductor

nanocrystals that can be used like fluorescent molecules in imaging experiments.33

In

addition to QDs, other nanoparticles are also used in imaging, such as scattering

centers and nanoclusters.34, 35

While QDs and nanoparticles offer a plethora of

photons, they blink via processes that are ill-understood. Recent efforts to mitigate this

blinking behavior have helped, but occurred at the cost of broadening the emission

spectrum.36

Also, QDs are considerably larger than both organic fluorophores and

FPs.37

The large size of QDs may hinder motion of the analyte and obfuscate the true

underlying dynamics. Fortunately, this problem is ameliorated by the low Reynolds

numbers in the cellular fluids; at such low Reynolds numbers, drag forces scale

linearly with velocity and radius (instead of scaling with the square of both). The

current obstacle to using QDs and nanoparticles is the difficulty of delivery to targets

inside live cells. QDs and nanoparticles are usually endocytosed and remain trapped in

endosomes instead of entering the cytosol, so they have faced significant hurdles in

application to live cells.38, 39

This drawback limits the use of quantum dot to the

cellular surface and endocytic events40

unless microinjection is used for delivery.41

14

Small-molecule fluorophores 1.2.2.2.3

Small-molecule fluorophores have the advantage that a huge variety of such

molecules are currently available with good fluorescence quantum yield and various

functionalities are available that allow for protein/nucleotide attachments. The

experiments described in this dissertation were predominantly conducted using small

organic fluorophores for their synthetic flexibility, small size, and favorable

photophysics. Commonly used classes of organic fluorophores include the following:

carbocyanine dyes,42

rhodamines,43

fluoresceins,13

dicyanometheleydihydrofurans

(DCDHFs),44

terrylene and rylenes,45

etc..

Notably, the DCDHF fluorophores, developed by the Moerner Lab and the

Twieg Lab at Kent State University, have a central structure based on an electron

donor-conjugated network-electron acceptor motif. These linear “push-pull”

molecules fluoresce when excited to the charge transfer band created by having an

amine donor linked via a conjugated network to a dicyanomethylenedihydrofuran

acceptor group.46

Because of a twisted intra-molecular charge transfer state and other

internal conformations, these molecules are also sensitive to the viscosity of their local

environments. The DCDHFs are brighter in more viscous environments, making them

useful molecules for the study of membrane dynamics.47-50

Chapters 3 and 5 in this

dissertation make use of this environmental sensitivity. The unique two-part nature of

the DCDHF molecules renders them unique candidates for rational design to impart

photoactivatability, an attractive quality for super-resolution imaging (see 1.3.1.).

To make the DCDHF photoactivatable, Samuel Lord of the Moerner lab

identified a clever scheme involving first photo-caging the molecule. During

experiments in which click-chemistry was being explored to turn-ion fluorescence, he

realized that a DCDHF with an aryl azide in the electron donor position instead of the

original aryl amine itself have have useful photochemistry, without the rest of the

complexity of the click approach. Because the azide is a much poorer electron donor

than the original amine, the resultant fluorescence is very low. At the same time, it is

well known that aryl azides are photolabile.51

Upon irradiation with violet light, the

aryl azide converts to a nitrene, which then can form a amine with help from nearby

15

protons or C-C bonds, restoring the original fluorophore.52, 53

To date, the azido-

DCDHF class of molecules have been used to establish three-dimensional imaging,54

as well as utilized for target-specific imaging of both live bacterial and mammalian

cells (see Chapter 4 and 5).55

Also attractive for super-resolution imaging is reversible photo-switching. In

early super-resolution experiments performed by the Zhuang group at Harvard,56, 57

Cy3 and Cy5 molecules in close proximity exhibited photoswitching behavior in the

presence of high concentration of thiols.58

In these stochastic optical reconstruction

microscopy (STORM) experiments, the Cy5 emitter was first excited with 633 nm

light until photobleached, and then the nearby Cy3 was excited with 532 nm light to

restore the Cy5 fluorescence. Later, it was demonstrated by the Sauer lab that with just

high intensity blue light and high concentration of thiols, Cy5 alone can be restored

back to fluorescence without Cy3.42, 59

The turnoff of the Cy5 emission has been

shown to occur via a covalent reaction with the thiol, which can reverse either

thermally or with light.60

Other photoswitchable small molecules have been explored

including photochromic rhodamines 61

and spiropyrans. 62

The next section will explore chemistry-based site-specific labeling for

conjugating favorable probes (e.g. organic fluorophores, quantum dots) to proteins-of-

interest (POIs).

In vivo site-specific labeling with chemical tags 1.2.2.3

A key goal in biology and biochemistry is to understand biological processes

by probing proteins, lipids, and oligonucleotides within their native environment (i.e.

living cell). To be able to examine biomolecules in such a complex and noisy space

requires the ability to selectively label the molecules of interest with imaging probes

so that we may see them above background. Due to the complex chemical

environment inside cells, many bio-conjugation reactions used prevalently in vitro

(e.g. the reaction of N-hydroxysuccinimide ester with lysine) are not sufficiently

selective. Further, while derivation of monoclonal antibodies have led to great steps

toward elucidating the roles of specific proteins in dynamic cellular processes, the

16

sheer size of antibodies renders them largely not amenable to in vivo imaging.

Therefore, bio-orthogonal site-specific labeling schemes are necessary for targeting

fluorescent probes to biomolecules of interest in the cell.63, 64

An ideal bio-orthogonal site-specific labeling method includes the following:

1) high specificity for target; 2) minimal perturbation to the target protein; 3)

versatility in fluorophore choice; and 4) ease of labeling procedure. Generally

speaking, in site-specific labeling using chemical tags, the underlying process requires

a high specificity reaction between a peptide (which may be attached to a protein of

interest) and an organic functional group (which may be attached to a fluorophore). In

this approach, the target protein is expressed as a fusion to a peptide tag. In

practicality, this involves the construction of a DNA plasmid encoding a fusion of the

target protein and peptide tag, and transfection of this plasmid into the cell. The

transfected cells are then incubated with the fluorophore-tag, which diffuses into cells

and is then specifically bound to the peptide tag. In this manner, site-specific chemical

tags retain the specificity of FPs through genetic encoding but provide modular use of

organic fluorophores with favorable photophysics.65-77

In vivo labeling with chemical tags can be broken down into roughly 3

categories by the interaction of the peptide with the fluorophore: 1) covalent binding

between the fluorophore and the peptide tag achieved via an active site on the peptide

tag (i.e.,the peptide take itself is an enzyme); 2) covalent bonding between the

fluorophore and the peptide tag mediated by a third party enzyme; 3) noncovalent high

affinity binding. The peptide tag can be either an intact protein or a short peptide

sequence.

Table 1-2 Various tag-probe labeling methods. This table was constructed by

collecting and organizing information from recent reviews.68, 78-80

Interaction

with POI

Tag/substrate Tag Size Use in live cell imaging

Covalent CLIP-tag/cytosine derivative67

20kDa extra-/intraceullular

Covalent SNAP-tag/benzyl guanine

derivative81


Covalent HaloTag/alkylhalide derivative70, 82


17

Covalent Cutinase/ p-Nitrophenyl

phosphonate 67,70,83,81

22KDa extraceullular

Covalent Tetracysteine helix motif/FlAsH,

ReAsh( biarsenical) derivative84-86

6(12) amino acids extra-/intraceullular

Enzyme-

mediated

covalent

Biotin ligase/BirA (biotin

derivative)76, 87

15aa; but requires

expression of the

ligase protein

intraceullular / in vitro

Enzyme-

mediated

covalent

Lipoic acid ligase/LplA (lipoic acid

derivative)77

22aa; requires

expression of the

ligase protein

extra-/intraceullular

Enzyme-

mediated

covalent

Coumarin ligase/Lpla(W37V)74

22aa; but requires

expression of the

ligase protein


Enzyme-

mediated

covalent

Sortase A/ Lactobacillus plantarum

sortase 'LPQTSEQ' sequence72, 88

5aa; but requires

expression of the

ligase protein


Enzyme-

mediated

covalent

TGase labeling/PKPQQFM89

2kDa extracellular

Enzyme-

mediated

covalent

ACP-Tag/coenzyme A90

9kDa extracellul

Noncovalent,

1nM affinity

eDHFR(LigandLink)/ trimethoprim

derivatives91


Noncovalent,

0.1nM affinity

FKBP(F36V)/ SLF’-derivatives92


Noncovalent,

80pM

Texas red aptamer/Xanthene core of

Texas red derivatives93

38AA intraceullular

Noncovalent Cognate single chain antibody

(scFv)/malachite green94

11-28kDa extra-/intraceullular

While further improvement of existing principles and development of new

schemes will be actively studied and pursued, current tag-probe techniques for specific

labeling in living cells have greatly added to the elucidation of the behavior of

membrane proteins in vivo.95

Cellular delivery and wash-out of unbound fluorophore 1.2.2.4

The cell membrane evolved, in part, to be a significant barrier to entry of

exogenous agents. Therefore, to label targets in the cytosol, a probe must first be able

to pass through the hydrophobic lipid bilayer. Therefore, fluorescent probes for in vivo

imaging should be either genetically expressed or membrane-permeable, unless

microinjection or electroporation is performed.

18

After an exogenous probe has entered the cell, excess unbound probes

introduce other complications such as cellular toxicity, heightened background, and/or

spurious signal. Therefore, the unbound probes must be sufficiently washed out of the

cell. Unfortunately, sometimes excessive washing can introduce unwanted mechanical

stress and is deleterious for the sample; in these cases, one can choose to use

fluorogenic labeling reactions,44, 84

or to adjust the detector integration times to

average out the signal (spread over many pixels) from quickly diffusing unbound

fluorophores to mitigate the effects from unbound fluorophores.96-98

Finally, it is

imperative that careful controls are performed to ensure that the labeling technique,

sample preparation, and imaging conditions do not alter the physiology of interest.

Applications of single-molecule imaging to biology 1.2.2.5

Moerner and Kador’s seminal single-molecule experiment was performed at

cryogenic temperatures in solids. In this experiment, the optical absorption of single-

molecule dopant impurities was successfully measured in a transparent sample (i.e. p-

terphenyl crystal) by direct sensing of the absorbed light.99

A year later, Orrit showed

that the optical absorption can be detected more easily by using fluorescence.100

In the

early single-molecule experiments, optical saturation, spectral diffusion, photon anti-

bunching, resonant Raman, electric field effects, and magnetic resonances of single

molecules were explored and observed.5 Relevant to the experiments described in this

dissertation, room temperature optical detection of single molecules was successfully

realized in solution,101-103

in microdroplets,104

and by tracking of single emitters in

porous polymers.105

As single-molecule experiments are uniquely positioned to answer problems in

asynchronous and heterogeneous systems that are prevalent in biology, single-

molecule biophysics and single-molecule cell imaging quickly became burgeoning

fields.5, 106, 107

As mentioned previously, single GFPs were imaged and controllable

photoswitching was demonstrated.30

This property was later explored for super-

resolution in live cells (see 1.3.1.). Förster-resonance-energy transfer (FRET) at the

single-molecule level was first shown in 1996 by Ha et al.,108

was demonstrated for a

19

pair of cyan and yellow GFPs in a Ca++

sensing construct,109

and later used for many,

many applications such as Ras activation in vivo.110

Single-particle tracking

experiments have also seen much development. The diffusion of single fluorescent

emitters in phospholipid membrane was first observed in 1996 by Schmidt et al.;111

since then, single-molecule tracking has been extensively used to explore the

mechanisms underlying many trafficking and transport events.41, 112-118

Molecular

motor proteins have also received a great deal of interest.116, 119-127

Single-molecule

imaging has also seen use in virology, measuring viral DNA packaging kinetics and

motor conformations.128

Pointillist super-resolution imaging is an application of single-molecule

imaging included in this dissertation; the essential background and a more in-depth

review of super-resolution imaging can be found in the following section.

Super-resolution in live cells 1.2.3

Super-localization 1.2.3.1

In 1873, Ernst Abbé devised an expression that defined the smallest spacing in

a specimen that can be resolved. This expression, the Abbé diffraction limit, is

presented below as Equation 1-2. It states that the smallest possible resolvable distance

between two objects given an imaging wavelength, λ, depends linearly on the

wavelength and inversely on twice the numerical aperture (NA) of the objective.129

The NA is defined as n sin Θ, where n is the refractive index of the imaging medium,

and Θ is the half angular range accepted by the objective.

sin22 nNA

Equation 1-2

In the visible spectrum, this translates to a diffraction limit of approximately

200-300 nm. When the object of interest is small, the diffraction limit presents a

dramatic size mismatch between the object and its image; for example, although a

20

small organic fluorophore is on the order of 1 nanometer in size, its image relayed by

the microscope is limited by diffraction to ~300 nanometers in size.

Fortunately, information about the point spread function (PSF) of the

microscope can help us gain information about the molecular position beyond

diffraction. The PSF of a microscope is the image produced by a single point emitter.

The PSF is the response of the microscope to an infinitesimally small point source of

light, and is the Fourier transform of the circular back aperture of the microscope

objective. This diffraction pattern resulting from the circular back aperture has a

central bright spot, termed the Airy disk, which, together with the concentric bright

rings around it, is called the Airy pattern.130

Because each photon that arrives at the

detector is a sample of the position of the molecule, the PSF can be regarded as the

probability distribution function for the localization of a single molecule. This

probability distribution takes a Gaussian-like shape with mean/peak being the position

of the molecule. Therefore, one can then measure the position of the molecule just by

finding the position of the peak. This is a process we term “super-localization.” The

theoretical precision related to super-localization is given by the standard error of the

mean with a sample size of N (i.e. N is the number of detected photons). In a single

dimension (e.g. x), the standard error of the mean σx is given by

N

sxx

Equation 1-3

The σx in Equation 1-3 is the localization precision, the most important figure

of merit in super-resolution experiments. sx is the standard deviation of the point-

spread function, and N is the number of detected photons. The localization precision

represents the uncertainty associated with one measurement of the position of a

molecule extracted from an image with N detected photons. Notably, a smaller sx

(smaller PSF), and a bigger N (more photons collected) both contribute to a better

(smaller) σx, but since sx is limited by diffraction, it cannot be made smaller.

Therefore a molecule that emits more photons can be localized with more precision

21

(i.e., organic fluorophores such as DCDHF can be localized much more precisely than

fluorescent proteins such as eYFP).

Figure 1-5 Super-localization of a single fluorescent molecule

(A) Image of the fluorescence from a single DCDHF-N6 molecule embedded in a thin

PMMA film. (B) Cross section in the x direction through the center of the image in

panel A. Each bin is a pixel (160 nm in width), and the counts in the pixel are the

digital counts of photons coming from the camera. The data is fit to a Gaussian with a

standard deviation of 200 nm, which is roughly equivalent to the diffraction limit. (C)

Distribution of 50 localizations obtained plotted on the same spatial scale as the data

in B, showing a drastically smaller distribution (a standard deviation of 9 nm). Figure

courtesy of Michael A. Thompson.

In an imaging measurement, the finite size of the detector pixels and

background noise also contribute to σx. Thomspon et al. devised the following

expression in 2002 to include these effects.131

In this expression, σx, the localization

precision is written as a function of a finite pixel size, a, and root mean square

background noise, b, in addition to N and s. The best-case two-dimensional

localization precision taking into account these experimental parameters can then

written as,

22

422 2812/

Na

bs

N

as

Equation 1-4

Equation 1-4 is often referred to as the Thompson-Larsen-Webb equation and

has seen much use in many super-localization microscopy studies.32, 132, 133

Due to

exclusion of higher order terms in its derivation, the Thompson-Larsen-Webb equation

overestimates the precision of true measurements by at least 30%.131

Nevertheless, this

expression provides a useful quantity for bench marking and comparison.

22

Active control 1.2.3.2

The second idea that enables super-resolution imaging is “active control” of

the emitters; Single-Molecule Active Control Microscopy (SMACM) has been coined

as an umbrella acronym for techniques using this concept.134

That is, to enable super-

resolution experiments, the concentration of emitters need to be actively controlled

such that the emitters are separated by more than the diffraction limit at any given

time, and can then be individually super-localized using ideas detailed in 1.3.1.1..

This concept is an extension of the principles demonstrated in early single-

molecule experiments. For example, many molecules within the same diffraction-

limited volume were separated spectroscopically, even in the first experiments in

1991.135

This idea was then extended and generalized in 1995 by Betzig, who

proposed to separate the emission of the molecules by controlling variables other than

absorption wavelength.136

Experimental realization of this proposal at low temperature

was achieved in 1998, to produce imaging resolution below the optical diffraction

limit via spectral selection.137

In 2006, three groups independently developed and

published room-temperature SMACM methods called Photo-Activated Localization

Microscopy (PALM),138

Stochastic Optical Reconstruction Microscopy (STORM),56

and FPALM (Fluorescence Photo-Activated Localization Microscopy).139

All three methods listed above separated the emission of the fluorophores

temporally by photoactivation (PALM and FPALM, using PA-FPs) or by

photoswitching (STORM, based on photochemistry of Cy3-Cy5 in the presence of

high thiol concentration). In PALM/FPALM, all the fluorophores within the specimen

start out initially dark. Then a pulse of 405 nm light is used to photoactivate a small

subpopulation of the molecules that are resolvable; these well-separated molecules are

imaged till they photobleach and then another pulse of activation light is applied to

turn on a new subset of molecules. This process is repeated until enough molecules in

the specimen are sampled to produce an image. This strategy is illustrated in Figure 1-

6. It is important to make sure that enough molecules are localized to yield a filled-out

structure; the next section describes this requirement in more detail.

23

Figure 1-6 Schematic showing the key idea of super-resolution imaging of a

structure by single-molecule super-localization and active control. (A) It is not possible to resolve the underlying structure in a conventional widefield

fluorescence image because the fluorescent labels are in high concentration and the

images overlap. (B) Using controllable fluorophores, it is possible to turn on and

image a sparse subset of molecules which then can be localized with nanometer

precision (blue line is the underlying structure being sampled). Once the first subset of

molecules photobleaches, another subset is turned on and localized. This process is

repeated and the resulting localizations summed to give a super-resolution “image”,

actually a reconstruction, of the underlying structure. Figure courtesy of Michael A.

Thompson.

Sufficient sampling 1.2.3.3

The Nyquist-Shannon sampling theorem140, 141

requires a sampling frequency

at double the bandwidth of the underlying signal to be able to accrue enough data

points to reproduce the signal without misrepresentation. Experimentally, this

translates to the requirement that, in order to attain a certain resolution, a structure

needs to be sufficiently labeled with an appropriate number of emitters per spatial

distance. In addition, uniform labeling density and the subsequently uniform super-

localization density through the structure are required in principle, but these will also

have a stochastic component. In cell-imaging experiments, these criteria essentially

require that the probe targeting strategy (see 1.2.2.3.) be sufficiently reliable to yield

sufficient and uniform labeling to define the underlying biological structure. This is

not always easy, as fusion proteins may not be incorporated into natural structures in

the same way that the original protein does.

24

An illustration of the importance of satisfying the Nyquist-Shannon condition

can be found in Figure 1-7. In an example of under-sampling (Figure 1-7A), the helix

from Figure 1-6 was sampled randomly at half the number of samplings used in Figure

1-7C. This difference in sampling number can lead to aliasing. Based on Figure 1-7A,

one can misinterpret the underlying structure as a series of distinct, separate bands

rather than one continuous helix. The problem of under-sampling occurs frequently in

cell imaging when the biological system cannot sustain/incorporate sufficient number

of labels (e.g. when the required level of labeling is cytotoxic, see Figure 1-4).

The pattern in Figure1-7B illustrates the case of non-uniformity in

labeling/localization. Here, the specimen was well labelled with many fluorophores

but the distribution of the fluorophores/localizations along the helix is irregular. This

irregular localization density results in the false clustering apparent in the resultant

image. Both oversampling of certain subpopulations of fluorescent emitters, and

irregularity in probe labeling can contribute to non-uniformity of localization density.

Figure 1-6C illustrates the structure obtained when the labelling dense and

uniform enough to present correctly the underlying helix.

Figure 1-7. The importance of the Nyquist-Shannon criterion in interpreting super-

resolution structures.

(A) Thirty-four samples of a helical structure sampled at random intervals.

Undersampling causes aliasing in the structure and it appears to be banded rather than

continuous. (B) Irregular and oversampling causes the structure to look more clustered

than in the evenly sampled, completely resolved case in (C). Figure courtesy of

Michael A. Thompson.

25

The Nyquist-Shannon requirement adds a further criterion to choosing emitters

for super-resolution experiments. Namely, the turn-on ratio (i.e. ratio of the signal of

the bright state to the signal of the dark states of the molecule) must be very high. This

is necessary to prevent the collective signal from many “dark” molecules within the

same diffraction-limited volume from eclipsing the signal from the one “bright”

molecule in the vicinity. To enable super-resolution experiments, several papers have

measured the turn-on ratio of fluorophores, but this parameter is not measured as often

as one would like.53, 142, 143

PA-GFP has a turn on ratio of ~ 100,142

and the azido-

DCDHF has a turn on ratio of > 1000 (see Chapter 4 for more details). Higher turn-on

ratio leads to better (smaller) localization precision and structural determination.

Single-molecule imaging in cells: a timeline 1.2.4

While studying live cells can be significantly more difficult than studying fixed

cells, it offers understanding of actual dynamic biological processes free from artifacts

from fixation. See Table 1-3 for a selected timeline of single-molecule experiments

with relevance to cell imaging. Please see Ref.144 for an extensive review.

Table 1-3 Selected single-molecule experiments with relevance to cell imaging. For a

list that includes both experiments in fixed and live cells, as well as in vitro

experiments up to 2008, please reference Ref.140. Senior authors listed.

Year Milestone (in gray are related in vitro experiments) Authors References

1991 first imaging of a single molecule in condensed phase

(cryogenic)

Moerner 5, 135, 145

1995 single motor protein imaged Yanagida,

Kinosita, Vale

119-121

1996 3D nanoscale tracking of single emitters, using TIRF

evanescent field

Dickson,

Moerner

105

26

diffusion of single emitters recorded in a phospholipid

membrane

Schmidt,

Schindler

111

1997 first single-molecule imaging of fluorescent proteins Moerner, Tsien,

Vale

30, 146

first room-temperature SMS example of controlled

photoswitching, a fluorescent protein

Moerner, Tsien 30

1998 first single molecule localization of mRNA particle in

live yeast

Singer 147

2000 spFRET measured in living cells Yanagida,

Kusumi, S.

Webb

110, 148, 149

3D tracking of single fluorophores in living cells Schütz,

Schindler

150

transmembrane ion channels tracked Schmidt,

Schütz,

Schindler

150, 151

2001 binding kinetics to chemotactic receptors in the

membrane observed

Yanagida 152

infection pathway of singly-labeled viruses observed Bräuchle 153

2002 SMS in membranes

Moerner,

McConnell,

Kusumi, Dahan,

Triller

115, 154-160

2004 protein localization and movement in living bacteria

cells using FP labels

Moerner,

Shapiro,

McAdams

31, 97, 98, 161

molecules tracked through nuclear pore complexes Musser 162

2005 nerve growth factor tracked in living neurons Tani, Yanagida,

Chu, Mobley,

Cui

163, 164

2006 high-precision tracing of motions in living bacteria Moerner,

Shapiro

97

super-resolution SMS techniques PALM, STORM, and

FPALM introduced

Betzig, H. Hess,

Lippincott-

Schwartz,

Zhuang, S. Hess

56, 138, 139

27

single-molecule gene expression events studied in living

bacteria

Xie 6, 96, 165

singly-labeled molecular motors observed in living cells Meyhöfer,

Dahan,

Cappello,

Spudich

116, 122, 126

2007 super-resolution SMS images of living cells Betzig, H. Hess,

Lippincott-

Schwartz, S.

Hess, Moerner

31, 166-169

SMS used to count the number of subunits in membrane-

bound proteins

Isacoff,

Verkman

170, 171

2008 interaction of cell-penetrating peptides with membrane

observed

Moerner,

Wender

50

single monomers of the cytoskeletal protein

photoactivated and tracked

Yu 114

2009 development of single molecule-sensitive probes for

imaging RNA in live cells

Crowe 172

2010 Super-resolution imaging of targeted proteins in fixed

and living cells using photoactivatable organic

fluorophores

Moerner 55

Super-resolution of ParA protein, a spindle-like

apparatus guides bacterial chromosome segregation

imaged in supe-rresolution

Moerner 173

2011 observation of single amyloid-β(1-40) oligomers on live

cells: binding and growth at physiological concentrations

Steel 174

Super-resolution imaging of the nucleoid-associated

protein HU in Caulobacter crescentus.

Moerner 32

super-resolution study of integrin-mediated adhesions Waterman 175

28

Outlook 1.3

Single-molecule experiments and super-resolution microscopy have already

revolutionized the life sciences by probing processes such as bacterial chromosomal

segregation173

and eukaryotic endocytosis.176

These techniques will likely become

common practice in the next generation of cell biology experiments.

To further increase the feasibility for routine application of single-molecule

and super-resolution techniques, developments in the areas of probes and labeling

strategies would be very helpful. Notably, the development of better fluorescent

proteins with higher photon counts, narrower, more red-shifted emission, and lower

molecular weights is needed. Targeted fluorogenic small organic fluorophores would

add significantly to the current tool kit. In addition, cytosolic delivery of quantum dots

and related nanostructures would make them much more applicable to live cell

imaging.

Imaging schemes to circumvent the out-of-focus fluorescence and spherical

aberration stemming from the cellular thickness would also be helpful. Recent

advances that can overcome some of these problems include sheet illumination,177

adaptive optics,178

and axially confined temporal focusing.179, 180

Scope of the dissertation 1.4

This dissertation describes the development and application of single-molecule

imaging and sub-diffraction imaging to elucidate structure and dynamics of biological

systems. Chapter 2 describes specific experimental methods that enable the studies

presented in this dissertation. Chapter 3 presents an application of single-molecule

imaging and single-particle tracking to explore the cell-penetrating mechanism of

oligoarginine molecular transporters. Chapter 4 describes the development and

characterization of a novel targeted photoactivatable small molecule fluorophore for

sub-diffraction imaging in both live bacterial and mammalian cells. Finally, Chapter 5

details the characterization and application of highly specific fluorescent toxins for

both single-molecule tracking and sub-diffraction imaging of sodium ion channels in

live neuronal cells.

29

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2.EXPERIMENTAL

45

Abstract

The success of super-resolution imaging hinges on the proper preparation of

the sample. The enormous diversity of biological systems requires that careful

selection be made from the wide array of previously developed tools in both molecular

biology and tissue culture. This chapter details some of those techniques that have

been most successful in the projects described in later chapters, as well as the selection

criteria to allow the proper employment of a technique with particular attention given

to the caveats of each particular method. The chapter begins with an introduction to

the possible microscope configurations, and then covers biological sample preparation

from culturing mammalian and bacterial cells in the dish to measuring samples on the

slide.

46

Microscopy Instrumentation 2.1

The instruments and methods used for single-molecule fluorescence

experiments have been well described and extensively reviewed in a number of

comprehensive review publications and theses. Notably, Refs.1, 2 are particularly

good, and were used to write this chapter. The projects I have had the privilege to be

involved in were applications of single-molecule detection to cellular imaging. In the

following pages, I summarize the basics of cellular imaging. For more discussion,

please reference the following recent Moerner Group Theses:

“Fluorophores for Single-Molecule Imaging in Living Cells: Characterizing and

Optimizing DCDHF Photophysics “

Lord, S. J. Ph.D. Dissertation, Stanford University, March 2010.

“The Development of Techniques for Three-Dimensional Super-Resolution

Fluorescence Microscopy and Their Application to Biological Systems”

Thompson, M.A. Ph.D. Dissertation, Stanford University, June 2011

Microscope Configurations 2.1.1

The instrumentation details and manufacturer information for individual

projects are detailed in each chapter; here I present considerations for the common

components of the projects included in this thesis. The imaging experiments included

in this thesis all use inverted optical fluorescence microscopes, configured in the wide-

field illumination configuration. In a few examples, confocal microscopy was also

used (see Chapters 4 & 5).

The canonical wide-field method used is epifluorescence. In epifluorescence, a

telescopically-expanded excitation beam is focused by the “Köhler lens” at the back

focal plane of a microscope objective, producing a collimated illumination beam at the

sample. Fluorescence emission from the illuminated sample is then collected back

through the same objective. In this collection path, scattered excitation light is

removed using a dichroic mirror and long-pass or band-pass filters. Finally, the

fluorescent signal is imaged onto a detector (i.e. camera). Since the excitation beam

47

mostly propagates through the specimen and any returning scattering is twice-filtered,

it is possible to collect a high signal to background ratio. A noted advantage of

epifluorescence is also that a large region of interest can be imaged simultaneously.

This is particularly suitable for single-particle tracking experiments.

Figure 2-1 General microscope configurations for single-molecule imaging in

biological samples. From Ref. 2.

In epifluorescence the excitation beam propagates through a large volume of

sample, exciting out-of-focal plane emitters, contributing to an increased background

signal in thicker samples. While this problem can be mitigated by using thinner model

cellular systems and reducing background emitters when possible, the problem cannot

be entirely removed for all samples. Total-internal-reflection fluorescence (TIRF)

imaging is a useful method to circumvent this problem for a subset of cases.3-6

In a

through-objective TIRF experiment, the excitation beam, after being telescopically-

expanded as in the case of epifluorescence, is directed into the objective off axis,

causing the excitation beam to totally internally reflect at the coverslip-water high-

index to low-index boundary. The internal reflection produces a sharply decaying

evanescent field extending into the low-index medium, here water.. As a result, only a

very thin slice of the actual sample nearest the coverslip is excited by the impinging

beam. The resultant optical section is much thinner than in confocal or wide-field

imaging (50–100 nm compared to 700 nm).7 The fluorescence from the sample is then

collected through the objective, filtered, and imaged using a camera just as in the case

of epifluorescence. Cell imaging experiments often opt to achieve TIRF via a through-

48

the-objective configuration (as depicted in Figure 2.1). Through-objective TIRF

configuration is more flexible compared to prism-based TIRF because the imaging

modality can easily be switched between TIRF and epi-illumination by translation of

the Köhler lens used to focus the beam onto the objective. An important advantage

with through-the-objective TIRF on an inverted microscope is that it is easier to create

an interface with cell-holding sample chambered coverslips.

However, because the evanescent field falls off exponentially within ~100 nm,

the applicability of TIRF is restricted to experiments in which the region of interest is

very near the coverslip. Furthermore, a related method, quasi-TIRF (also referred to as

“pseudo-”, “leaky-TIRF”, or “Hi-Lo”) also sends in the excitation beam off-center, but

not far enough to produce an angle of incidence large enough to create total-internal

reflection. In the case of quasi-TIRF, an angled excitation beam escapes out the top of

the sample, illuminating it in a slice that is thicker than the TIRF evanescent field but

thinner than the epifluorescence focal plane.

Another method to reduce out-of-focus fluorescence and achieve high signal-

to-background is confocal microscopy.8-10

Confocal microscopy is a scanning point-

detection method. In confocal microscopy, a collimated excitation beam that overfills

the back aperture of the objective is directed into the microscope, producing a tiny

diffraction-limited spot at the sample. The confocal excitation spot is scanned across

the sample and the fluorescence emission from the sample is then collected through

the objective and filtered. The filtered emission signal is then focused through an

aperture to reject any emission from out-of-focus slices of the sample. The light

exiting through the pinhole is then collimated again, and focused onto a point photon

detector such as a Si avalanche photodiode (APD). Confocal imaging has the

advantage of not being constrained to only being able to image near the coverslip, so it

can be used to image deeper into a sample and to acquire three-dimensional

information. A primary limitation of confocal microscopy is that it requires scanning

the sample stage/excitation beam, and the use of a point photon detector. These two

requirements make imaging multiple parts of the sample simultaneously impossible,

although there are a number of new designs where many sources and samples

49

illuminate the sample at the same time using a spinning disk. In any case, the various

advantages and disadvantages may make the parallel wide-field epifluorescence

microscopy method more advantageous for experiments where the dynamics of

interest may be on par with (or faster than) the confocal scanning speed for the same

region of interest.

Detectors, in the form of Si charge-coupled device cameras (CCD) and APDs,

optical lenses, for focusing and collimating, and a light source, for excitation, are

common components to each of the methods mentioned above. In most studies, laser

sources of suitable coherence are used simply for the high brightness and diffraction-

limited focusing achievable, but when these properties are not needed, carefully

filtered lamps and even LED light sources can be used. Please see reference 1 for a

detailed discussion. In the following sections, I outline some of the basic

considerations for selecting each of these common components for cellular imaging.

Detectors 2.1.2

The type of detector used is a critical parameter in cellular imaging

experiments and particularly important in single-molecule imaging, as it partially

determines the sensitivity and, indirectly, spatial resolution of the obtained image. 11

To effectively detect the photon flux from single emitters, a detector must exhibit the

following characteristics: 1) low dark counts (expressed in electrons per unit of time at

a given temperature); 2) low noise (from reading, electron multiplication, or analog-to-

digital conversion); 3) high quantum efficiencies over visible wavelengths. For further

discussion and details regarding specific detectors and their characteristics and

capabilities (i.e. resolution, sensitivity, signal-to-noise ratios) please see references.1, 12

There are two classes of detectors most often used for cell imaging: single-

element/point detectors, and array detectors. Point detectors are used for confocal

imaging and include photomultiplier tubes (PMT) and avalanche photodiodes (APD or

SPAD). The advantage of PMTs is that they have a larger detection area (~1 cm2) with

very high temporal resolution (ps–ns). APDs have very low dark counts and much

higher quantum efficiencies and more easily detect single photons; moreover, APDs

50

have faster time resolution (relative to PMTs), and output a digital signal that allows

for ready interfacing with computers. The major drawbacks of APDs are (a) the small

detection area, which makes aligning onto the sensor more difficult, (b) the limited

photon detection rates, and (c) possible afterpulsing.

Wide-field imaging configurations allow the use of multi-detector arrays or

cameras as a region (versus a single point-per-unit-time in confocal microscopy) is

illuminated. For the experiments included in this thesis, the highly useful, but only

recently available, back-illuminated electron-multiplying (EM) CCD cameras are

used. These cameras typically have quantum efficiencies >80% for the visible

spectrum and frame-integration times of 10–100 ms, or faster for fewer pixels.

The EMCCD is essentially a standard CCD with a gain stage added between

the shift register (where the charge of each pixel is loaded to be read) and the output

amplifier (which converts charge to a digital voltage). Three fundamental steps occur

during conversion of an impinging photon into a digital count in the readout. Figure 2-

2 outlines the process that occurs when a photon excites a photoelectron in an

EMCCD pixel.13

The purpose of the third stage is to properly digitize the signal. Most

cameras allow the conversion gain to be adjusted in the software. Because the readout

of each pixel happens after the massive gain stage, the readout noise of the camera

will be very small compared to the signal that is fed into the conversion amplifier.

Figure 2-2 Illustration of the gain stages and ADC conversion of photons into

counts by the EMCCD camera.13

This process takes a 3-step scheme. In the first step,

n photons detected by the EMCCD excite n photoelectrons on chip. In the second step,

the cumulative charge from the photoelectron is loaded into an on-chip electon-

multiplying gain register, which converts n photoelectrons into α×n electrons by

impact ionization in a high electric field. This α parameter, the gain factor, typically

ranges from the hundreds to thousands. Third, the amplified photoelectrons are then

fed into an analog-to-digital buffer/converter, which takes G “gain” electrons and

converts them to a single count. The purpose of the third stage is to properly digitize

the signal. 13

51

Optics 2.1.3

As mentioned, high quality lenses, mirrors, and filters are especially important

for the ultrasensitive detection required for single-molecule imaging.1 In addition to

these considerations, some additional considerations may be required for imaging in

cells.

To achieve sensitive single-molecule imaging, filters (including dichroic

mirrors) must transmit as much of the desired fluorescence as possible, and must reject

as much of the Rayleigh scattering from the excitation light as possible. Thus, filters

must have sharp cut-on spectra, with optical density >6 at undesired wavelengths.13

The use of bandpass filters in place of longpass filters in the emission path can be

helpful to remove longer-wavelength background fluorescence or excess water Raman

scattering. If one choses to use a bandpass filter, it is important to compare the filter’s

transmission spectrum with the emission spectrum of the fluorescent label to avoid

rejecting too much of the emission as described in Ref. 1. Filters and lenses inside the

microscope should be anti-reflection (AR)-coated and aberration-corrected.

Plan (flat-field) microscope objectives with high numerical aperture (NA ~1.4)

that are well-corrected for chromatic aberrations over the visible spectrum are

necessary to collect as much of the emission as possible. A lens with an NA above the

refractive index of air (NA ~1) is designed to be used with an immersion medium of

higher refractive index such as immersion oil, water, or glycerol in the space between

the objective and the cover slip. The use of an immersion medium in the sample as

well decreases the optical refraction and total internal reflection at the glass–water

interface, resulting in an image with better contrast.

In live-cell imaging with high numerical aperture objectives, one needs to take

precaution to avoid spherical aberrations. Spherical aberrations come from refractive

index mismatch between the sample media (e.g. cell growth media or aqueous buffer

and the refractive index of the immersion media (e.g. immersion oil, typically equal to

the refractive index of the glass cover slip).7 Spherical aberration results in an

asymmetric spread in the focus along the optical axis such that the point spread

function of an emitter may become significantly elongated and distorted. Because the

52

signal is spread over a much larger volume in the presence of the aberration, the

important parameter of signal-to-noise ratio in a single-molecule image can be

significantly reduced. This problem can be especially debilitating in cell imaging, as

the cell sample may exhibit significant background autofluorescence. The spherical

aberration problem can be ameliorated by the use of relatively thin and adherent cells

(usually only several micrometers thick), and can be largely eliminated by using a

water immersion objective that is more closely matched to the refractive index of the

culture medium and carefully adjusted for a specific cover slip thickness.7 As an

alternative, the refractive index of the immersion medium or the thickness of the

coverslip can be adjusted. Further, refractive index changes with temperature, so the

optimal combination of immersion medium and coverslip thickness will differ slightly

between room temperature and 37 ° Celsius. This is a consideration whenever a stage-

top incubator is required.14

Excitation Light Source 2.1.4

In the early days of single-molecule imaging, single-frequency tunable dye

lasers were used in the cryogenic experiments.15

At room temperature, gas (e.g. argon-

ion, helium-neon, etc.), diode, or solid-state lasers are typically used. While lasers are

necessary for the tight focusing in confocal microscopy, epifluorescence excitation is

also possible using broadband sources such as arc lamps or light-emitting diodes. The

arc lamps produce a large amount of heat via infrared emission, which can damage

live cell specimens. A heat absorption filter (a filter the filters out infrared light)

placed in the light path could help to ameliorate this thermal-damage to the cellular

sample.7

White-light or fiber-based lasers can also provide broadband pulsed light,

produced by nonlinear optical effects when high-intensity very short pulses are

transmitted through special optical fibers. Broadband sources can be convenient

because they are tunable to a range of colors, but add the complication of having to

filter carefully to obtain a reasonably monochromatic excitation source. In all cases,

the excitation source used in the single-molecule experiment should be filtered prior to

53

reaching the sample to reject unwanted leakage of any other colors, including long-

wavelength broadband emission from plasma discharges. In live cell imaging,

significant effort should be taken to avoid exposure of the sample to unwanted blue

light to avoid pumping cellular autofluorescence (see Chapter 1 for further discussion

on this topic).16

Cell Culture 2.2

Since the early 1900s, cell culture has been a pivotal tool for studies of biology

and biochemistry.17-20

The term cell (or tissue) culture refers to the cultivation for

growth of cells apart from the host organism. Accompanied by aseptic techniques,

cell/tissue is generally facilitated by the use of liquid, semi-solid, or solid growth

media of different materials, such as broth or agar to sustain cellular survival and

growth. This body of knowledge and techniques enable the use of in vitro models of

the primary tissue in a well-defined environment, which can then be controlled,

changed, and analyzed. Cell culture techniques have also been important in facilitating

single-molecule and super-resolution cellular imaging experiments.21-32

The rest of

this chapter details bacterial cell culture, mammalian cell culture, and protein handling

as related to the cellular studies used in this thesis. Culturing these vastly different

systems provides the cell imager with a rich repertoire of models to work with for a

variety of experiments; while culturing the different systems can be quite different in

detail, the recurrent motifs of cultivation, freezing, and thawing of the cell stocks are

common in all culturing practice. The following sections are organized as such, and

include additional sections on transfection/transformation, and fixation.

Bacterial Cell Culture 2.2.1

As mentioned in 2.2., bacterial cultures can be grown with a solid nutrient

support or with liquid broth-based media.33

Specifically, bacterial cultures can first be

grown in petri dishes containing a layer of agar-based growth medium. Once the

growth medium in the petri dishes has been streaked (inoculated) with the bacterial

strain of interest, the petri plates are then incubated at suitable temperature for the

54

growth of the selected bacteria. When the plates are inoculated properly, bacterial

strains grow in distinct colonies, with each colony a uniform phenotype.

The second method of bacterial culture is suspension liquid culture. In this, the

bacteria of interest do not grow in distinct colonies unless the initial inoculation is

started from a single colony. In the following, I detail the cultivation of Caulobacter

crescentus, which can also be adjusted to be used for Escherichia coli when steps are

taken to use the appropriate reagents.

Caulobacter crescentus cultivation 2.2.1.1

Caulobacter crescentus is a gram-negative, oligotrophic, aquatic bacterium

that divides asymmetrically at the end of each cell cycle. This asymmetry renders

Caulobacter crescentus an interesting system for studying cell division, cell cycle

regulation, and cellular differentiation, a decades-long area of interest in the laboratory

of Lucy Shapiro, a close collaborator of the Moerner lab.26, 34-37

Despite its unusual

asymmetric cell division, much like other bacteria, Caulobacter crescentus’s growth

rate is determined largely by several factors including nutrient availability, cell

density, temperature, and the environmental pH value. As mentioned previously,

bacteria can be cultured on agar plates or in liquid suspension. In preparing samples

for wide field single-molecule experiments, samples of well-separated cells are

desired. For this reason, we generally grow our bacteria culture in liquid suspension.

We use both nutrient rich and minimal nutrient media. Nutrient rich media provide a

source of amino acids and nitrogen, as well as carbon such as glucose. Whereas

minimal growth media, containing only the minimum nutrients possible for colony

growth, do not provide amino acids; rather, minimal growth media provide carbon and

various salts (to provide magnesium, nitrogen, phosphorus, and sulfur) to allow the

bacteria to synthesize proteins and nucleic acids. Minimal growth media is used during

imaging to lower background fluorescence.26, 31, 38-40

Figure 2-3 details the general flow of bacterial cultivation for experimentation.

In this section, I describe the techniques necessary for bacterial cell culture, as

outlined by the figure.

55

Figure 2-3 General scheme for bacterial culture cultivation. Further details about

each step can be found below.

When bacterial cells are inoculated in a closed and fresh medium, the number

of cells monitored over a period of time (typically assessed by optical density

measurements of the medium corresponding to cell density) generally follows the

curve shown below (growth curve, Figure 2-4). More specifically, bacterial growth in

batch culture can be roughly divided into four disparate phases: lag phase, log

(actually exponential growth) phase, stationary phase, and death phase. Experiments

on bacteria should generally be conducted when cultures are in the log phase to ensure

proper cell cycle progression and function. For Caulobacter crescentus, cultures must

have an OD at 660 nm <0.4 to be in log phase.

56

Figure 2-4 A typical bacterial culture growth curve. After a certain period of time to

activate the metabolism (lag phase), cells grow exponentially (log phase) until they

reach the “stationary phase.” Ultimately, due to the limitations of nutrients, space,

etc., cells cannot grow further, and the number of viable cells declines.41

The first phase of bacterial culture growth is the lag phase, during which

bacteria adapt themselves to the growth conditions. In this phase, individual bacteria

mature, proceed through their cell cycle, but are not yet able to divide. It is important

to note that, in this phase of bacterial culture growth, typical cell cycle progression,

synthesis of RNA, and synthesis of different proteins are still in progress. Therefore, it

is important to recognize that microorganisms are not dormant in the lag phase. 41

The second phase is the log phase, characterized by facile and steady cell

division. In this phase, the rate of new bacteria generation is proportional to the

present population, yielding exponential growth. If we maintain the same culturing

condition, not replenishing nutrients or diluting cell suspension concentration, the log

growth phase will lead into the stationary phase as the new cells will soon deplete the

nutrients in the growth media and replace with waste products.

The third phase is the stationary phase. During the stationary phase, cell

growth plateaus as a result of the cell starvation from nutrient depletion and

accumulation of waste products. This phase is reached as the bacteria begin to use up

all the nutrient resources, and leads into the death phase. Finally, during the death

phase, the bacteria run out of nutrients and die.

57

To prepare Caulobacter crescentus for imaging, cells were streaked from

DMSO stocks onto a PYE (Peptone-Yeast Extract) agarose plate using a sterile plating

stick. To obtain well-isolated discrete colonies, one should use the quadrant streak

technique. This method provides a means to achieve sequential dilution of the original

bacterial material. This is achieved by successively streaking over quadrants, so that

the density of bacteria in each streak decreases. This technique is detailed in the

following.

Figure 2-5 Schematic of inoculating an agarose plate for bacterial culture. See the

following steps for streaking sequence.

1. Using a sterile plating stick (or autoclaved pipette tip, or flamed metal stick),

touch the tip of the stick to either a broth culture or a frozen aliquot. Immediately

streak the inoculating stick gently over a quarter of the agarose plate using a

streaking motion (see area 1 in Figure 2-5).

2. Using a new sterile stick (or after flaming the original one) gently go over the

edge of the first quadrant, extending the streaks into the next quadrant (see area 2

in Figure 2-5).


edge of the second quadrant, extending the streaks into the next quadrant (see

area 3 in Figure 2-5).


edge of the third quadrant, extending the streaks into the next quadrant (see area

4 in Figure 2-5).

58

After 4 days at 30 ºC, a single well-isolated colony is selected from the

streaked plate and placed into PYE media and incubated overnight. The colony

selection is done with either a sterile pipette tip or a sterile plating stick, taking care

not to touch neighboring colonies. On the next day, cells are diluted into M2G buffer,

and incubated at 28 ºC to allow two full cell cycles with OD 660<0.4. PYE media has

nutrients from bactopeptone and yeast extract, as well as several salts necessary for

Caulobacter growth. Since PYE media had considerable optical impurities

contributing to spurious signal, especially high background fluorescence, for

fluorescence microscopy experiments we always use the M2G (Minimal to Growth)

buffer instead. In contrast to the high-nutrient PYE medium, M2G buffer has only

glucose and ammonium salts as nutrients. The doubling time of Caulobacter in the

M2G media at 30 ºC is approximately 140 min.

Freezing cells, and re-establishing growth from frozen samples 2.2.1.2

It is well known that many living organisms are capable of tolerating

prolonged periods of time at temperatures below water’s freezing point. This makes

cryopreservation possible. For many purposes, including preserving experimental

continuity and reproducibility, it is important to store cells for future use.

Cryopreservation ensures that we will always have back-up cell aliquots in the

unfortunate event of contamination or culturing instrument malfunction.

Cryopreservation can be achieved by keeping frozen cell stocks at -80°C for several

months-long storage, or <-120°C for long-term storage for important samples. At

these temperatures, biological activities that eventually lead to cell death are slowed

down substantially. To achieve this in nature, living organisms accumulate

cryoprotectants such as anti-nucleating proteins, polyols, and glucose to protect

themselves against frost damage by sharp ice crystals. In particular, Herminiimonas

glaciei have reportedly been revived after surviving for thousands of years frozen in

ice.42

Instead of relying on the microorganisms to produce polyols, we use

cryoprotectant solutions to help the cells from suffering ice crystal damage during

thawing.

59

Freezing down bacterial aliquot stocks demands careful sterile techniques to

ensure that only the intended bacteria are frozen. When starting with a plate of

bacteria, first make sure it is clear and fresh (i.e. with isolated colonies visible). Using

a sterile stick, choose a single colony and inoculate 5ml of sterile broth. Grow bacteria

overnight (~16 hours) at its appropriate temperature to reach the appropriate density.

Do not overgrow since that may overcrowd cells into the stationary phase. In sum, it is

best to cryopreserve cells when they are at their maximum growth rate (i.e. log phase)

but at high density to ensure efficient thawing later on.

It is common practice to create a master bank consisting of 10 vials of the cell

line. Then create one or two working banks from this with 10 vials in each (depending

on how often the cells will be required). When the working bank is used up, a new

working bank can be cultured and created from one vial of the original master bank.

Careful freezing and budgeting of the cell bank can ensure that one will always have

stocks at lower passage numbers. This is helpful as sometimes phenotypes change

over time (i.e. at high passage number) and it may be necessary to go back to a lower

passage for experimental continuity.

Preparing Samples for Imaging 2.2.1.3

To be able to image bacteria in a wide field set-up, it is important to

immobilize them while preserving a nutrient-rich environment. In the following we

describe the preparation of Caulobacter crescentus and Escherichia coli for imaging.

Agarose immobilization of live cells 2.2.1.4

The Moerner laboratory has had success with imaging on agarose pads from

some years,43

which follows this procedure: Add approximately 500 µl high-purity,

low-fluorescence and low-melting point, 1.5% agarose (made with M2G buffer for

Caulobacter crescentus, or other minimal growth buffer for other bacterial strains) on

a microscope slide. Place a second slide on top, avoiding air bubbles, and press gently

to obtain a thin agar slab with an even thickness.

60

After about 1 minute, pull the two slides apart (pull in a direction parallel to

the plane of the slide). With some practice and care, this will leave a thin layer of

solidified agarose on one of the slides. Let the agar dry briefly with gaseous nitrogen,

add 2 µl of well-separated cell suspension (as little as possible) and place a coverslip

on top. Using a razor blade, excess agarose gel is then cut off. The edge of the agarose

and glass is then sealed with melted paraffin wax. We then invert this resulting

sandwich of coverslip-agarose-coverslip to image. See Figure 2-6 for schematic.

It is strongly advised that 2 transmission (or DIC) images be taken for every

region of interest that an epifluorescence image stack is taken; one taken before the

epifluorescence image stack, and one taken after. This gives us information about cell

mobility, morphological change, and possible drift during the course of the

experiment. In super-resolution imaging, addition of fluorescent fiduciary beads to the

sample is strongly recommended to correct for stage/sample drift.

Figure 2-6 Schematic for preparing bateria containing agarose gel pads on overslips

for imaging.

Cell synchrony 2.2.1.5

In a bulk batch bacterial culture, the four phases detailed in 2.2.1.1 do not

directly correspond to phases in the cell cycle. The cells do not divide in synchrony

and their log phase growth is often not ever purely exponential, but instead slows

down over time, a constant stochastic response to pressures both to reproduce and to

go dormant in the face of declining nutrient concentrations and increasing waste

concentrations. As a result, in order that we might be able to experiment and image

61

cells within the same stage of cell cycle, extra steps must be taken to synchronize the

bacterial sample. Caulobacter is especially nice in this respect, as the Shapiro lab has

developed a useful procedure for cell synchronization. The following steps take

advantage of the fact that cells in different stages in the cell cycle have different

densities and yields a sample of synchronous swarmer cells.44

1. Start with 14-15 ml of a culture in log phase, with plenty of swarmers present.

Spin at 6000 rpm for 10 minutes at 4C.

2. Pour off the supernatant and resuspend the pellet in 1 ml of cold 1X M2 media,

transferring to a 2 ml eppendorf tube, to rinse off residual media. Spin for 3

minutes at 13,000 rpm.

3. Aspirate off supernatant and put the pellet on ice immediately. Carefully

resuspend the entire cell pellet in approximately 900 ul of 1X M2.

4. Add 900µL of Percoll. Percoll (Pharmacia) consists of 15-30 nm PVP coated

silica particles. It has a low viscosity, which allows cell separations to be

completed in only a few minutes using low centrifugal forces. It is sterile, non-

toxic to cells, and adjustable to physiological ionic strength and pH. It is more

expensive than the alternative, Ludox reagent, and is therefore used only for

mini-synchronies.

5. Spin for 20 minutes at 11,000 rpm, 4C.

6. Aspirate off the top stalk cell band and collect the swarmer band.

7. Wash the swarmers two times in 1 – 1.5 ml of cold M2. Centrifuge at 7000-8000

for 3 minutes.

8. Resuspend the pellet in 5 to 10 ml of desired media to allow cells to progress

through the cell cycle.

Transformation 2.2.1.6

Transformation is defined as the genotype alteration of biological cells

resulting from the direct incorporation and expression of exogenous genetic material

(i.e. DNA and RNA). Transformation is a very useful technique in constructing model

systems that can express proteins and fusion proteins of interest (see Chapter 4 for an

62

example). A natural occurrence, transformation can also be achieved by human means.

In molecular biology, bacteria that can be transformed, whether naturally or

artificially, are called competent. Please see reference for bacterial transformation

strategies.45-47

Introduction of foreign DNA into eukaryotic cells is usually called

"transfection." Please see 2.2.2.6. for more details on mammalian cell transfection.

Bacterial Fixation 2.2.1.7

Fixation, the dehydration/cross-linking of cellular proteins, can be useful in

conducting immunofluorescence experiments, and in the exclusion of protein

dynamics from structure. The protocol described here is sufficient for simple bacterial

cell fixation (i.e. for visualization of fluorescent fusion proteins). In this protocol,

methanol or formaldehyde is used as a fixative. Formaldehyde and other aldehydes are

by far the most commonly used fixatives in histology, and are considered strong

fixatives. Formaldehyde fixes tissue by cross-linking the proteins, primarily using the

residues of the basic amino acid lysine. In addition to formaldehyde, glutaraldehyde is

another popular crosslinking aldehyde. Glutaraldehyde functions by causing

deformation of the alpha-helix structures in proteins. In comparison, glutaraldehyde is

a larger than formaldehyde in size, and so the glutaraldehyde fixation process

generally needs a longer time to allow glutaraldehyde enough time to diffuse into the

cell. A noted advantage of glutaraldehyde fixation is that it may offer a more rigid or

tightly-linked fixed sample— its greater molecular size and two aldehyde groups

allow it to connect more distant pairs of protein molecules compared to formaldehyde.

On the other hand, glutaraldehyde is fluorescent, so it is not ideal for

immunofluorescence staining unless extensive washing of the sample is done prior to

imaging. A combination of both formaldehyde and glutaraldehyde is also sometimes

used, taking advantage of both fixatives.

Methanol, on the other hand, functions via a different reaction from the

aldehydes. Methanol and other alcohols are precipitating (or denaturing) fixatives.

Precipitating fixatives operate by reducing the solubility of protein macromolecules

and frequently also by disrupting the hydrophobic interactions necessary for protein

63

tertiary structure. The precipitation and aggregation of proteins is a very different

process from the crosslinking that occurs with the aldehyde fixatives and care should

be taken to make sure that the fixation process does not affect the cellular structure of

interest.

Lastly, if immunofluorescence or other labeling protocols are desired in

conjunction to cell fixation, care need to be taken to remove as much of the fixative as

possible. Also, please see the separate protocol for mammalian cells (2.3.1). I detail

the bacterial fixation protocol in the following:

1. Grow cells as usual in M2G. Harvest cells at ~ OD660 0.3–0.4.

2. Prepare mixture of 135 L methanol in 5 mL cold 1 M2 salts (or 135 L 37%

formaldehyde in 5 mL cold 1 M2 salts).

3. Spin 5 mL cell culture in a 15-mL falcon tube (7830 rpm for 90s) at 4 C.

Remove supernatant.

4. Re-suspend the cell pellet in the cold methanol or formaldehyde mixture. Be

careful to pipet gently; no bubbles should form (this is particularly difficult with

the formaldehyde).

5. Incubate cells in the formaldehyde or methanol solution at room temperature for

10 min, shaking gently once or twice to make sure that the cells stay suspended

and do not pellet.

6. Incubate cells on ice for >30 min.

7. Spin cells for 15 min at 7830 rpm at 4 C to pellet.

8. Remove supernatant and resuspend in 5 mL chilled 1 PBS buffer (pH 7.4) or

chilled 1 M2 salts. Spin for 15 min at 7830 rpm at 4 C.

9. Repeat step 7 two more times for a total of three washes.

The fixed cells can now be stored in the falcon tube for days. Prepare samples

for imaging on agarose pads as usual. If bacteria remain alive, double the

concentration of formaldehyde (to 270ul) and use the same protocol as above.

64

Mammalian Cell Culture 2.2.2

Tissue culture techniques have found wide application in many fields such as

cell biology, medicine, biochemistry, and lately in interdisciplinary fields.48

Offering

control over physiochemical condition as well as the constancy of physiological

conditions of biological samples, cell cultures are utilized in a variety of research

settings for diagnostic and research studies. In the experiments detailed in this thesis,

the access and ability to culture a variety of model cell lines have been essential. For a

list of milestones in cell culture technique history, please see Table 2-1.

Table 2-1 Important timeline in tissue culture technique development 19, 20, 49

Year Event

1878 Proposal by Claude Bernard states that physiological systems of an organism can be

maintained in a living system after the death of an organism.

1885 Embryonic chick cells were successfully kept alive in a saline culture by Roux. 50

1907 The father of cell culture, Harrison, cultivated frog nerve cells in a lymph clot held by

the 'hanging drop' method and observed the growth of nerve fibers in vitro for several

weeks.

1910 Burrows succeeded in long-term cultivation of chicken embryo cells in plasma clots;

his success led to the first observation of mitosis.

1911 The first liquid media, consisting of seawater, serum, embryo extract, salts, and

peptones was developed by Lewis and Lewis.

1913 Carrel introduced strict aseptic techniques so that cells could be cultured for long

periods.

1916 Rous and Jones introduced trypsin for the subculture of adherent cells.

1923 Carrel and Baker developed the T-flask as the first specifically designed cell culture

vessel.

1940 Antibiotics, penicillin and streptomycin, used in culture media decreased the problem

of contamination in cell culture.

1952 Establishment of a continuous cell line from a human cervical carcinoma known as

HeLa cells by Gey et al. from the cancerous tissue of Henrietta Lacks.

1955 Eagle studied the nutrient requirements of selected cells in culture and established the

first widely used chemically defined medium.

1961 Hayflick and Moorhead isolated human fibroblasts and showed that they have a finite

lifespan in culture.

65

1965 Ham introduced the first serum-free medium, which was able to support the growth

of some cells.

1973 F. L. Graham and A. J. van der Eb developed calcium phosphate transfection.

1978 Establishment of serum-free media from cocktails of hormones and growth factors by

Sato et al.

Both primary and secondary (i.e. immortalized or continuous) mammalian cell

lines can be used as models in microscopy experiments. Primary cell lines are taken

directly from host organisms in the form of tissue. Cells can then be isolated from

animal tissues for ex vivo culture in several ways. From blood, white blood cells can

easily be purified and cultured as a suspension culture. Epithelial and fibroblast cells

are excellent adherent flat microscopy model systems that can be extracted from soft

tissues by dissection, followed by digestion with enzymes such as collagenase, trypsin,

or pronase.48, 51

These enzymes break down the extracellular matrix and release cells

out of their confined from. Once re-plated to a suitable surface (e.g. tissue-culture

polystyrene petri dishes with surface treatment to stimulate growth), healthy

fibroblastic and epithelial cells will secrete extracellular matrix proteins and

proteoglycans that the cells can use to adhere to via cell surface receptors. With the

exception of certain tumor-derived cell lines, these primary cell lines have finite life

span. During the life span of the cell lines, primary culture offers scientists access to

cells that do not divide in culturing conditions (e.g. neurons, macrophages). Notably,

when working with primary cell lines like mouse embryonic fibroblasts (MEFs), it is

particularly important to note the passage number of specific samples as certain

cellular functions change over time.

On the other hand, secondary cell lines can be cultured for a long time in

optimal culturing conditions. These cell lines have acquired the ability to proliferate

indefinitely either through genetic mutation or deliberate genetic modification, such as

induced expression of the telomerase gene via transfection (details in 2.2.3.7.). There

are numerous well-established cell lines representative of particular cell types; they

generally divide much faster, and provide the advantage of culturing ease. 17-19, 51-56

See Table 2-2 for a list of common continuous cell lines.

Table 2-2 Commonly used continuous cell lines for microscopy.

66

Cell

Line

Host Origin Tissue Morphology Ploidy Required coating for

cell adhesion to glass

NIH-

3T3

Mouse Embryonic

Fibroblast

Fibroblast Aneuploid none

CHO Hamster Ovary Epithelium Diploid Fibronectin

COS-7 African

green

monkey

Kidney Fibroblast Diploid none

HEK-

293

Human Cervical

Cancer

Epithelium Aneuploid Fibronectin, or poly-L-

lysine

HeLa Human Cervical

Cancer

Epithelium Aneuploid none

Vero

cells

African

green

monkey

Kidney Fibroblast Aneuploid Poly-L-lysine

BSC-1 African

green

monkey

Kidney Epithelium Aneuploid none

PC12 Rat Medulla fibroblast Diploid Collagen, fibronectin,

poly-L-lysine, matrigel

In most cases, cells must be grown in culture for weeks or months to sustain

the length of research projects. Maintenance of cells in culture for this length of time

requires strict and careful adherence to aseptic techniques to avoid contamination and

potential loss of valuable cell lines. In the following sections aseptic techniques, sub-

culturing techniques, freezing and thawing of cell storage, as well as transfection will

be discussed.

Aseptic techniques 2.2.2.1

As mentioned in 2.2.2, since Carrel et al.’s development of aseptic techniques

in 1913, the use of aseptic technique has been standard for successful tissue culture.

Cell cultures can be contaminated by bacteria, mycoplasma, yeast, or other cells at any

time during handling, so precautions must be taken to minimize the possibility of

contamination. In general, all supplies and reagents that come into contact with cell

67

culture and enter the incubator must be sterile. It is also strongly advised that work

surfaces should be kept clean and free from unnecessary clutter. The following

procedure details the sequential steps that one must follow to maintain sterility in

tissue culture areas. These steps relate primarily to the Biosafety Level 1 cells used in

the Moerner lab; much more complicated procedures are essential for cell lines of

higher Biosafety Level.

1. Sanitize the laminar flow/biosafety cabinet using 70% ethanol and 1%

benzalkonium chloride before commencing work. Benzylalkonium chloride is a

cationic surfactant that maintains sterility by preventing microbial growth. Note

that the physical process of ethanol evaporation is essential in establishing

sterility; therefore it is important to allow the ethanol solution to evaporate before

moving on to the next step.

2. Sanitize gloves by spraying them in 70% ethanol and 1% benzalkonium chloride

and allowing to air dry for 30 seconds before commencing work.

3. After putting all materials and equipment into the cabinet spray them down with

70% ethanol and 1% benzalkonium chloride prior to starting work.

4. In the vertical laminar flow hood, the air is directed down toward the surface, so

the airflow pushes contaminants such as bacteria and yeast down. Care should be

taken to avoid blocking the airflow, and the grills at the front and back of the

hood should always remain unobstructed.

5. While working, do not contaminate hands or gloves by touching anything outside

the cabinet (especially face and hair). If gloves become contaminated re-sanitize

with 70% ethanol and 1% benzalkonium chloride as above before proceeding.

6. Discard gloves after handling contaminated cultures and at the end of all cell

culture procedures.

7. Equipment in the cabinet or that which will be taken into the cabinet during cell

culture procedures (media bottles, pipette tip boxes, pipette aids) should be wiped

with tissue soaked with 70% ethanol prior to use.

8. Movement within and immediately outside the cabinet must not be rapid. Slow

movement will allow the air within the cabinet to circulate properly.

68

9. Speech, sneezing and coughing must be directed away from the cabinet so as not

to disrupt the laminar airflow.

10. After completing work inside the laminar flow hood, disinfect all equipment and

material before removing from the cabinet. Spray the work surfaces inside the

cabinet with 70% ethanol and wipe dry with tissue. Dispose of tissue by

treatment with bleach solution.

11. Sanitize the cabinet with 10 – 30 min UV light daily. Warning – plastics will

crack and become brittle over time with repeated exposure to UV light. Only

some cabinets have timed UV lights. Ensure they are not left on for extended

periods.

12. Discard waste by first treating with 10% bleach solution for 30 min., then

disposing of the waste down the drain. For cells with different Biosafety Levels,

consult EH&S.

Decontamination of the incubator is required at least yearly. To conduct this,

remove all racks from incubator for autoclaving, and heat the cell incubator up to 80°C

for 3 hours. It may also be necessary to fume the incubator with formaldehyde if

fungal contamination is a repeated problem. To do this, place an open container of

10% formaldehyde solution in the incubator for 30 min at 80°C.

Cell line passaging and plating for imaging 2.2.2.2

Passaging (also known as sub-culturing or splitting cells) involves transferring

a small number of cells into a new vessel and media. Standard mammalian cell growth

media include the following components: amino acids, vitamins, salts (Na+, K

+, Mg

2+,

Ca2+

, Cl-, SO4

2-, PO4

3-, HCO3), glucose, and animal sera as a source for minerals,

lipids, and hormones. It is important to note that sera from the same host organism and

same manufacturer may differ from batch to batch; therefore, records of the batch of

serum used should be kept carefully, batch-matching may be necessary to ensure

continuity of experiments, and using chemically serum-free media should be

considered when planning experiments. 48, 51

When possible, ordering a large quantity

of a particular batch can ensure repeatability.

69

Phenol red is often included in media as a pH indicator, but to lower the

background fluorescence, cell culture in the Moerner group is performed without

phenol red. To further lower the background, live cells are imaged in supplemented

buffer containing only salts and sugars wherever possible. In addition, antibiotics are

also frequently used in growth media to prevent contamination of cell culture.

It is important to split cells often, as cells can be cultured for a longer time if

they are split regularly, as splitting avoids the cellular senescence associated with

prolonged high cell density. Some cells are contact-inhibited, and are especially

susceptible to this. Suspension cultures are easily passaged with a small amount of

culture containing a few cells diluted in a larger volume of fresh media. For adherent

cultures like epithelial cells or fibroblasts, cells first need to be detached; this is

commonly done with a mixture of trypsin-EDTA, however other enzyme mixes are

now available for this purpose. A small number of detached cells can then be used to

seed a new culture.

Adherent cell lines will continue to divide until they have covered the surface

area available or the medium is depleted of nutrients. Prior to this nutrient depletion

the cell lines should be sub-cultured in order to prevent the culture dying. To

subculture the cells, it is necessary that the adherent cells are brought into suspension.

The degree of adhesion depends largely on the distinct extracellular matrices that vary

from cell line to cell line, but in the majority of cases proteases, e.g. trypsin, are used

to release the cells from the flask as mentioned above. However, this may not be

appropriate for some lines where exposure to proteases is harmful or where the

enzymes used remove membrane markers/receptors of interest. In these cases cells

should be brought into suspension into a small volume of medium either with EDTA

solution or mechanically with the aid of cell scrapers.57, 58

Before starting to passage each time, it is important to examine the culturing

dish/flask under the microscope. This practice is also important for familiarizing

oneself with the cell shapes, division speed, and degree to which cells carry debris

with different cell lines so that contamination (e.g. yeast, bacteria, fungus),

spontaneous differentiation and changes to the morphology can be detected.

70

After passaging, pay attention to the length of time it takes for the cells in the

newly established cultures to attach and spread out. Cellular attachment within an hour

or two is indicative of a successful passage and longer attachment times are suggestive

of problems. Nevertheless, good cultures may result even if attachment does not occur

for four hours. The following details the general procedure for passaging.

1. Remove used medium.

2. Wash the cell monolayer with enough PBS without Ca2+

/Mg2+

to cover the

monolayer.

3. Pipette trypsin onto the washed cell monolayer using 1ml per 25cm2

of surface

area (e.g. 1 ml – T25, 3 ml T75). Swirl the culturing flask to cover the monolayer

with trypsin.

4. Incubate the flask in the hood for 2-10 minutes. Too long of a period of

trypsinisation may be harmful to the cells.

5. Once the cells start to sheet and the media becomes cloudy, move on to the next

step. Examine the cells using a microscope to ensure that many (~80%) the cells

are detached and floating. It may help to “slap or tap” the flasks gently to release

any remaining attached cells.

6. Re-suspend the cells in a small volume of fresh pre-warmed serum-containing

growth media to inactivate the trypsin. Generally speaking, for every 2 mL of

trypsin used, use 8 mL of media to inactivate the trypsin.

7. Disaggregate clumps or sheets of cells by running the suspended cells in the

medium three to four times through the tip of the pipet on the bottom corner of

the flask. Be very careful not to aspirate growth media into the pipet aid. If this

happens, the filter MUST be replaced.

For cell passaging go to step 8. For plating for imaging go to step 9.

8. Transfer the required number of cells to a new labeled flask containing pre-

warmed medium. The name of the cell line, date of passage, passage number, and

the name of the passage should all be on the flask.

9. Transfer the required number of cells to a new appropriately-coated cell

chambers with #1 borosilicate coverslip bottoms (LabTeK, Nunc). Refer to Table

71

2-2 for the proper glass surface coating, and section 2.8 for associated

procedure). Approximately 400µL of cell suspension from step 7 is needed for

each chambered well. The name of the cell line, date of passage, passage

number, and the name of the passage should all be written on the chambered

coverslip. Approximately 2- 6 hours is needed for the cells to adhere properly to

the surface for imaging.

Counting Cells 2.2.2.3

For accurate cell counting, the hemacytometer slide should be clean, dry, and

free from lint, scratches, fingerprints, and watermarks. The coverslip supplied with the

hemacytometer should always be used because it has an even surface and is specially

designed for use with the counting chamber. Use of an ordinary coverslip may

introduce errors in cell counting. If the cell suspension is too dense or the cells are

clumped, inaccurate counts will be obtained. If the cell suspension is not evenly

distributed over the counting chamber, the hemacytometer should be washed and

reloaded.

Figure 2-7 A. Hemacytometer, B. Loading a hemacytometer channel, C.

Hemocytometer grid: red square = 1.0000 mm2, 100.00 nl green square = 0.0625

mm2, 6.250 nL yellow square = 0.040 mm

2, 4.00 nl blue square = 0.0025 mm

2, 0.25

nL at a depth of 0.1 mm.59

1. When lifting the cells with trypsin/EDTA, reserve approximately 50 μL of the cell

suspension. A 96 well plate is well suited for working with these small volumes.

2. Gently mix 20μL of the cell suspension with 20 μL of the 0.4% Trypan blue

solution.

72

3. Place the coverslip on top of the hemocytometer, and pipette <20 μL of the Trypan

blue cell suspension into the hemocytometer channel. The solution should coat the

hemocytometer.

4. Count the number of unstained cells in the four corner squares of the

hemocytometer. The sick or dead cells are permeable to Trypan blue and will

appear heavily stained.

5. Calculate the number of cells in the centrifuge tube according to:60

(

) (

) ( )( )

6. When finished, gently wipe the coverslip and hemocytometer with lens tissue and

ethanol.

Freezing cell lines 2.2.2.4

Cells in culture will undergo changes in growth, morphology, and genetic

characteristics over time. Such changes can adversely affect reproducibility of

laboratory results. Non-transformed cells will undergo senescence and eventual death

if passaged indefinitely. The time of senescence will vary with cell line, but generally

at between 40 and 50 passages fibroblast cell lines begin to senesce. Cryopreservation

of cell lines will protect against these adverse changes and will provide a backup in

case of contamination.

Cultures selected for cryopreservation should be in good condition(i.e. clean

and intact membranes, and free from excessive debris), in the log-phase growth, and

free of contaminants. Cells should be harvested as in the case of passaging,

concentrated, and flash frozen at a concentration of 106 to 10

7 cells/ml in the

appropriate cryoprotectant solution for each cell line.60

Cells should be frozen slowly

and thawed quickly to prevent formation of ice crystals and osmotic shock that may

cause cells to lyse, respectively. Cell lines can be thawed and recovered after long-

term storage in liquid nitrogen. The top of the freezing vial should be cleaned with

70% alcohol before opening to prevent introduction of contaminants as discussed in

the next section. Careful records on the date, cell identity, cell passage number, the

73

personnel involved in the freezing, and characteristics of frozen cells as well as their

location in the freezer should be maintained to allow for easy retrieval.

1. Remove medium from one dish / flask, wash and trypsinize as described in

2.2.2.2.

2. Once cells are detached, add back 5-10 ml media and transfer to centrifuge tube

(15 ml sterile centrifuge tube).

3. Count the cells using trypan blue for a viable cell count. The viability should be

over 90% to ensure the cells are healthy enough for freezing.

4. Spin down at 1500 rpm for 5 minutes and remove medium.

5. Slowly re-suspend cells in enough cold freezing medium to create a cell

suspension of 1x106 cells per ml. Pipette up and down to ensure even mixture

and aliquot about 1 ml into storage vials. This will provide 1×106 cells per

cryovial.

6. Transfer cells immediately to -20°C for one hour, followed by -80°C overnight

before permanent storage in liquid nitrogen. For cells that will be used in a time

frame of months, storing in -80°C would be sufficient.

Thawing frozen cells 2.2.2.5

It is vital to thaw cells correctly in order to maintain the viability of the culture

and enable the culture to recover more quickly. Some cryoprotectants, such as DMSO,

are toxic above 4ºC, therefore it is essential that cultures are thawed quickly and

diluted in culture medium to minimize the toxic effects. To ensure good recovery of

cultures, thawed cells should be reseeded at a higher concentration than that used for

passaging.

1. Remove cryovial from liquid nitrogen and place in a water bath at 37ºC for

mammalian cells. Submerge only the lower half of the ampule. Allow to thaw

until a small amount of ice remains in the vial - usually 1-2 minutes. Transfer to

biosafety laminar flow cabinet.

2. Wipe the outside of the ampule with a tissue moistened (not excessively) with

70% alcohol; hold tissue over ampule to loosen lid.

74

3. Slowly, dropwise, pipette cells into pre-warmed growth medium to dilute out the

DMSO.

4. Incubate cells overnight. Change media the next morning. Removal of DMSO is

critical.

5. Examine cells microscopically (phase contrast) after 24 hours and sub-culture as

necessary.

Transfection 2.2.2.6

The definition of transfection is "infection by transformation". For most

applications of transfection, it is sufficient if the transfected genetic material is only

transiently expressed. Since the DNA introduced in the transfection process is usually

not integrated into the nuclear genome, the foreign DNA will eventually be diluted

through mitosis or degraded. If it is desired that the transfected gene actually remain in

the genome of the cell and its daughter cells, a stable transfection must occur. To

accomplish this, a selection marker gene is usually co-transfected, which gives the cell

some selectable advantage, such as resistance towards a certain antibiotic. Very few of

the transfected cells will, by chance, have integrated the foreign genetic material into

their genome. If the selection antibiotic is then added to the cell culture, only those

few cells with the marker gene integrated into their genomes will be able to

proliferate, while other cells will die. After applying this selective pressure for some

time, only the cells with a stable transfection remain and can be cultivated further. We

then call the resultant cell line stably transfected.

There are various methods of introducing foreign DNA into a eukaryotic cell;

we detail a few commonly used chemical-based methods below.61-66

Physical means

such as heat shock, electroporation, sonoporation, and microinjection are also

sometimes used.67-69

It is worthwhile to note that high levels of expression of certain

proteins (e.g., GFP) may be cytotoxic for some cell types. Therefore, when

transfecting, one should pay attention to possible changes in cell morphology, division

speed, and general health of the resultant cell culture.

Calcium Phosphate transfection 2.2.2.6.1

75

As mentioned in Table 2-1 in 2.2.2, F. L. Graham and A. J. van der Eb first

developed calcium phosphate transfection in 1973.70

This technique takes buffered

saline solution (HBS) containing phosphate ions and combines it with a calcium

chloride solution containing the DNA to be transfected.62, 71

When the two

components are combined, precipitation of the positively charged calcium and the

negatively charged phosphate occurs, and thus binding the DNA to be transfected on

its surface. The suspension of the precipitate is then added to confluent, adherent cells.

By a process not entirely understood, the cells take up some of the precipitate, and

with it, the DNA. Generally speaking, this technique is very economical, as alternative

liposomal carriers (see 2.2.1.3.3. for more info) tend to be more expensive. It is also

important to know that this technique is highly DNA sensitive, and some trial-and-

error may be required to find the optimum DNA concentration. The following details a

general set of procedures.

1. To prepare the calcium phosphate–DNA co-precipitate, combine 10 µl of 2.5 M

CaCl2 with 2 µg of plasmid DNA in a sterile 500µl plastic tube and, if necessary,

bring the final volume to 0.1 ml with TE buffer (10mM Tris and 1mM EDTA,

pH 7.6). Mix one volume of this calcium phosphate–DNA solution with an equal

volume of HEPES-buffered saline (HBS) at 15–25 °C. Tap the side of the tube to

mix and incubate the solution at 15–25 °C for 1 min.

2. Immediately transfer the calcium phosphate–DNA suspension into the medium

above the cell monolayer. Use 0.1 ml of suspension for each 1 ml of medium in a

well. Rock the plate or dish gently to mix the medium, which will become

turbid/cloudy.

3. Return the cells to the incubator for incubation at 37 °C for 2–6 hrs; then remove

the medium and DNA precipitate by aspiration. Add pre-warmed (37 °C)

complete growth medium to each dish of cells and return the cells to the

incubator for 24–48 hours prior to imagining.

DNA-liposome complex transfection 2.2.2.6.2

76

A very efficient method for transfection is the inclusion of the DNA to be

transfected in liposomes, i.e. small, lipid-bilayer-bounded spherical shells (vesicles)

that are in some ways similar to the structure of a cell and can actually fuse with the

cell membrane, releasing the DNA into the cell. For eukaryotic cells, transfection can

be achieved using cationic liposomes (or mixtures), because the cells are more

sensitive to this vesicle charge.61, 63, 72-75

2.2.2.6.2.1 Lipofectamine transfection

The following protocol is for a 1 cm2 surface area culture vessel (e.g. 1 well in

the chambered coverslip). Also see the Invitrogen documentation for protocols for

different versions of Lipofectamine complex.76, 77

1. Heat OPTI-MEM(Invitrogen) to 37ºC in a water bath.

2. Observe the cells under microscope – check for contamination and note cell

confluence.

3. Place the cells in the hood.

4. Remove the cell culture media, and add 2 mL of media (without serum or

Pen/Strep).

5. Aseptically place lipofectamine and DNA in the hood.

6. Pipette 92 uL of OPTI-MEM into a sterile polystyrene tube.

7. Pipette 8 uL of lipofectamine directly into the OPTI-MEM media. Avoid

direct contact between lipofectamine and the tube wall.

8. Tap tube to mix.

9. Incubate for 5-10 min at room temperature.

10. Add 1 µg of DNA vector to the dilute lipofectamine. The amount of DNA

may be changed depending on the system.

11. Tap to mix.

12. Incubate for 30 min at room temperature.

13. Remove the dish of cells from the incubator.

14. Add the DNA-containing lipofectamine complex to the cells in a drop-wise

fashion.

15. Gently swirl dish to cover the cells.

77

16. Label the petri dish (DNA vector used, date and time of transfection).

17. Place the cell-containing petri dish back in the incubator (replace with serum

containing medium in 6 hours).

2.2.2.6.2.2 Fugene6 (Promega) transfection

The following protocol is for a 1 cm2 surface area culture vessel (e.g. 1 well in

the chambered coverslip). Also see the Invitrogen documentation for protocols for

different versions of Fugene6 complex.

1. Heat phenol-red free DMEM to 37ºC in a water bath.

2. Observe the cells under a microscope – check for contamination and note cell

confluence.

3. Place the cells in the hood.

4. Remove the cell culture media, and add 2 mL of media (without serum or

Pen/Strep).

5. Aseptically place FuGENE 6 and DNA in the hood.

6. Pipette 91 uL of DMEM into a sterile polystyrene tube.

7. Pipette 9 uL of FuGENE 6 directly into the DMEM media. Avoid direct contact

between FuGENE and the tube wall.

8. Tap tube to mix.


10. Add DNA vector (1 ug) to the dilute FuGENE. Use a plasmid vector

concentration between 0.02 and 2.0 µg/µl.

11. Tap to mix.


13. Remove the dish of cells from the incubator.

14. Add the FuGENE/DNA complex to the cells in a drop-wise fashion.

15. Gently swirl dish to mix.

16. Label the dishes (DNA vector used, date and time of transfection).

17. Place the cells back in the incubator.

18. Replace with serum containing medium in 6-24 hrs. This time depends on the

cell line.

78

Immunofluorescence 2.3

Immunofluorescence experiments use the specificity of antibodies to their

antigen to target fluorescent dyes to specific bio-macromolecule targets within a cell,

and allows visualization of the distribution of the target protein within the cell sample.

The general flow of an immunofluorescence experiment is depicted in the following

figure (Figure 2-8). Overall, the process starts with a wash to remove cell debris,

fixation to cross-link cellular components, permeabilization of the cells, pre-blocking

to prevent unspecific binding, application of primary antibodies, a second pre-

blocking to prevent unspecific binding, application of fluorescently labeled secondary

antibodies, and finally a wash to remove any unbound secondary antibodies to remove

background.

Figure 2-8 A. Schematic for immunofluorescence experiments B. Antibody

schematic

The binding affinity of primary and secondary antibodies used is an extremely

important parameter that determines the overall signal-to-background, and can vary

79

widely. Figure 2-8B depicts the simplified structure of and antibody . Composed of a

heavy chain and two light chains, an antibody is a immunoglogulin protein (e.g., IgG

or IgM), generally with epitope binding sites, called paratopes. For most

immunofluorescence experiments, there is a primary and a secondary antibody.

Secreted by plasma cells, primary antibodies are raised against a target antigen of

interest (e.g. protein, peptide, carbohydrate, or other small molecule) and are most

frequently unconjugated (not fluorescent). These antibodies are useful not only to

detect specific biomolecules but also to measure changes in their expression level and

specificity of modification by biochemical processes such as phosphorylation,

methylation, or glycosylation. The paratope of the primary antibody is the portion of

the antibody that changes depending on the antigen of interest, but the rest of protein

remains unchanged from target antigen to target antigen. This allows a use of a

secondary antibody to recognize the part of the primary antibody that is unchanged

(i.e. heavy chain). The secondary antibody is an antibody that binds to the primary

antibodies. The secondaries are labeled with fluorophores probes for detection,

purification, and cell sorting applications.78-81

Multiple immunofluorescence labeling can be done with the same procedure as

single labeling. In these experiments, antibodies derived from different host animals

can be mixed and incubated simultaneously as a cocktail (e.g. rabbit anti-tubulin +

mouse anti-caveolin). In applying the secondary, this is also true (e.g. goat anti-rabbit

IgG Alexa488 + goat anti-mouse IgG Alexa647). Careful consideration should be

taken to ensure there is no significant spectral overlap amongst the different

fluorophores used to avoid loss of signal due to FRET (unless FRET is a desired

readout for protein proximity).

Different chemical reagents and their concentrations for fixing and blocking

can be tried to optimize the immunostaining of cellular samples. However, since the

parameter space is enormous it is advisable to start with an established protocol for the

same or closely related protein targets. For single-molecule imaging, it is strongly

recommended that one starts with a protocol that works for ensemble-level imaging,

and then work to increase signal-to-background by increasing specificity and

80

decreasing non-specific binding. In choosing a chemical fixative, one should consider

the subcellular localization of the target antigen (e.g. soluble, transmembrane,

cytoskeleton-associated) as certain fixatives introduce artifacts into the system of

interest. For example, methanol fixation is easy and also permeabilizes the membrane,

providing time economy; however, methanol may solubilize transmembrane antigens,

allowing them to diffuse away. As this approach relies on simple precipitation of the

protein, methanol only provides low structural preservation. If a high degree of

structural preservation is necessary (e.g. if immunoelectron microscopy is intended),

aldehydes such as paraformaldehyde and glutaraldehyde should be used as a fixative.

Since glutaraldehyde is fluorescent, proper washing and quenching is necessary. Also,

when using paraformaldehyde, using a freshly-made fixing solution is imperative as

paraformaldehyde oxidizes over time.80-82

Using detergents to permeabilize the membrane should be done with caution,

especially when imaging membrane-associated antigens. Lastly, the quality of pre-

block solution is extremely important to reduce undesired non-specific binding of

antibodies. Some commonly used pre-block reagents include host sera (e.g. use goat

serum for antibodies raised in goat or horse serum for antibodies raised in horses),

bovine serum albumin, and casein, but as always, autofluorescence from the preblock

must be tested.

While immunofluorescence experiments require many washing steps,

mechanical stress to the sample should be kept to a minimum to avoid compromising

the quality of the final sample. To aspirate or add solution to the sample, one should

gently touch the pipette tip to the side of the chambered coverslip and pipette gently.

Care should be taken to never aspirate or apply solutions directly onto the sample.

Also, the cells should be kept hydrated throughout the process of sample prep.

Lastly, visualization of the cytoskeleton has been an important proof of

principle for a variety of single-molecule imaging schemes. The cytoskeleton is very

dynamic and sensitive to both chemical and mechanical pressure. Also, high levels of

actin monomers and tubulin (i.e. subunits for microtubules) in the cytosol can

compromise the signal-to-background of cytoskeletal filaments and render it difficult

81

to analyze the detailed localization of filamentous skeletal proteins. To overcome this

complication, monomers can be briefly extracted from the cells before fixation to

achieve a filament stabilizing condition. This extraction of monomers, when

performed appropriately, removes free subunits/unbound binding proteins from the

cellular cytoplasm without compromising polymeric structure.

In the following sections, I include protocols for immunofluorescence and

antibody conjugation. Protein isolation from bacterial and mammalian cells (i.e. cell

lysate production) procedures can be found in the sections following. Cell lysate can

be very helpful to ascertain the specificity of antibody conjugates.

Immunofluorescence protocol 2.3.1

1. 8+ hours before immunostaining, plate cells on #1 chambered coverslips.

Depending on the cell line, the coverslip may have to coated. Refer to Table 2-2

for details.

2. Fix cells with 4.0% paraformaldehyde in PBS (pH 7.4) at room temperature for

20min.

3. Wash the sample three times with PBS. Each time, soak the sample in PBS for 10

minutes.

4. Permeabilize and pre-block the cells with 0.2% Triton X-100 in 1% BSA

containing PBS on ice for 30min. This solution is the blocking solution. Note

here, the 1% BSA can be substituted by sera, or casein as mentioned above in

2.3.

5. Dilute primary antibody in the blocking solution described above (dilutions will

vary depending on the specific antibody; start with the manufacturer’s

recommendation).

6. Add diluted primary antibody solution to the cell sample. Place the sample in a

humidified chamber.

7. Incubate 1 hour at room temperature, or overnight at 4°C.


minutes.

82

9. Pre-block the cells with 0.2% Triton X-100 in 1% BSA containing PBS on ice

for 30min.

10. Dilute secondary antibody in the blocking solution described above (dilutions

will vary depending on the specific antibody; start with the manufacturer’s

recommendation).

11. Add the secondary antibody dilution to the cell sample. Place the sample in a

humidified chamber.

12. Incubate cells at room temperature in the dark for 1 hour.


minutes.

Antibody conjugation 2.3.2

Given that the desired fluorophore-conjugated antibodies may not be available

commercially, it may be at times necessary to label antibodies with a fluorophore of

choice. The following details this protocol. It is important to verify the specificity of

antibody for target antigen after this labeling procedure via Western blot.

Labeling antibodies with amine-reactive molecules (NHS-Esters) 2.3.2.1

1. Dissolve 2 - 10 mg of protein in 1 ml of sodium bicarbonate buffer. Reaction

Buffer = pH 8.15, Sodium bicarbonate 0.15M. Adjust pH to desired value with

HCl.

2. IgG protein solutions must be free of any amine-containing substances such as

Tris, free amino acids, or ammonium ions.

3. Dissolve 1.0 mg of amine-reactive dye in 100 to 500 μl of anhydrous, amine-free

DMSO at 1.0 mg/ml.

* Steps 1 and 3 can be modified based on the reaction yield- basically, a ~ 3-10

molar excess of the amine-reactive dye in the final reaction solution is desired.

4. Prepare the dye solution immediately before starting the labeling reaction to

prevent hydrolysis of the NHS-ester.

83

5. Slowly add, while stirring with your pipette tip, a threefold molar excess of

reactive dye to the protein solution.

6. Incubate the reaction mixture for 3 hour at room temperature. To increase the

degree of labeling a higher ratio of NHS-ester to protein has to be used.

7. Use a Bio-Spin p30 column for purification of labeled protein from reaction

mixture and free dye. I usually purify ~3 times via the p30. Alternate

purification protocols, such as dialysis, may be used in some cases.

Storage of the protein conjugate 2.3.2.2

1. For storage in solution at 4 °C, sodium azide (2 mM final concentration) can be

added as a preservative.

2. The conjugate should be stable at 4 °C for several months.

3. For long term storage, divide the solution into small aliquots and freeze at -20 °C.

Determining the degree of labeling (DOL) 2.3.2.3

The degree of labeling (DOL, fluorophore-to-IgG ratio) can be determined by

absorption spectroscopy making use of the Lambert-Beer law:

( )

( )

( )

Simply measure the UV-VIS spectrum of the conjugate solution as obtained

after gel filtration in a quartz (UV-transparent) cell.

[ ]

[ ]

[ ] [ ]

84

Total protein isolation from cells/cell lysis 2.4

Cell lysates are created by lysing the cell membrane, usually via osmotic or

enzymatic means, and are composed of the entire contents of the live cells. This

provides access to cellular proteins in an in vitro environment that includes all cellular

contents. Cell lysates are often important in trouble-shooting in small molecule

method development, as by using lysates, one can study interaction of small molecules

with cytosolic proteins without having to solve the problem of cell entry. Enclosed in

the following are procedures for both bacterial and mammalian cells.

Total protein isolation from bacterial cells 2.4.1

Bacterial cell lysate is an important diagnostic tool to assess bacterial protein-

small molecule interactions. I include the following procedure for total protein

isolation from bacterial cells.

1. Harvest cells by centrifugation of cellular suspension for 10 min at 7500 × g, and

wash in a buffer containing 25 mM Tris/HCl (pH 7.9) and 5 mM MgCl2.

2. Add 270 L 37% formaldehyde (1% final),

3. Incubate at room temp for 10 min with occasional shaking, then 30 min on ice.

4. Spin cells for 15 min at 8000 rpm at 4oC.

5. Pour off supernatant and wash in the 10 mL chilled 1x PBS, pH 7.4, to remove

the formaldehyde. Do this step on ice.

6. Repeat steps 4 and 5 for a total of 2 washes and spin again.

7. If cells are ready to be used, proceed to step 8. If a stopping point is needed,

resuspend the pellet in 10mL PBS, spin for 5 min., and discard the supernatant.

Flashfreeze the cells in liquid nitrogen and store at -80oC until needed.

8. Resuspend the cells in 1 mL lysis buffer (10 mM Tris, pH 8, 20% sucrose, 50

mM NaCl, 10 mM EDTA) with 20 mg/ml of lysozyme freshly added. **Make

sure that the lysozyme is not DNase contaminated. Flame the spatula before

scooping.

9. Incubate at 37oC for 30 minutes.

85

10. Add 1 mL IP wash buffer (100 mM Tris, pH 7.0, 300 mM NaCl, 2% Triton X-

100) with 1 mM PMSF added fresh. (PMSF stock soln – 100 mM in EtOH)

11. Incubate the cells for 10 min. at 37oC and then put on ice.

12. Divide the mixture into 4 500 L aliquots for sonication.

13. Sonicate the cells with a Branson sonifier set at “hold”, “constant”, and about

20% output in 10 second pulses. The tubes should be packed in ice and straight

during sonication. In between pulses, let the sample rest on ice.

14. If the solution foams, spin down for 1 min max speed at 4oC.

Total protein isolation from mammalian cells 2.4.2

Mammalian cell lysate is an important diagnostic tool to assess protein-small

molecule interaction. I include the following procedure for total protein isolation from

mammalian cells.

1. Prepare RIPA Lysis buffer

a. 1X RIPA :150 mM NaCl, 0.5% sodium deoxycholate, 0.1% SDS, and

50 mM Tris, pH 8.0.

b. Add 10 µl PMSF solution, 10 µl sodium orthovanadate solution and 10

µl protease inhibitor cocktail solution to 1ml of 1X RIPA buffer to

prepare complete RIPA lysis buffer,

2. Pour off media from tissue culture dish into waste container (100mm dish is most

often used for this).

3. Wash cells twice with PBS pouring excess off into waste beaker.

4. Carefully soak up any extra PBS with a Kimwipe.

5. Add 150ul of RIPA lysis buffer to the culture dish.

6. Use cell scraper to scrape cells from the bottom of the dish.

7. Pass cell lysate through pipette 20 times to form homogeneous lysate.

8. Transfer lysate to 1.5 ml microcentrifuge tube.

9. Allow samples to stand for 5 minutes at 4ºC (in cold room or on ice).

10. Centrifuge the resulting mixture at 14,000g for 15 mins at 4ºC to separate cell

debris from protein.

86

11. If storage of cell lysate is required, transfer supernatant to a new tube and store at

-20ºC.

Coverslip coating 2.5

Immobilization and coating of glass coverslips are necessary for observation of

different biomolecules and cell lines. Refer to Table 2-2 for cell lines and their

preferred surface coating for proper cell adhesion. In the following, I include the

procedures for making poly-L-lysine, collagen and poly-electrolyte multilayer-coated

coverslips.

Fibronectin surface coating 2.5.1

1. Place trays in a Petri dish, with lens paper underneath.

2. Warm a 25 L aliquot of frozen fibronectin until it melts with hands.

3. Pipette the fibronectin into 3.2 mL of Dulbecco’s PBS, and mix gently.

4. Add 200 L of the fibronectin solution to each well, coating the bottom of each

well by tilting gently.

5. Let the fibronectin solution sit in the trays for 1 hour at room temperature.

6. Harvest the cells as normal. Obtain final 16 x 104 cells in 3.2 mL total volume.

7. Aspirate the fibronectin solution from each well, and replace with 200 L of

diluted cell solution in each well, swirling to ensure each well is coated.


minutes.

Collagen surface coating 2.5.2

1. Place trays in a Petri dish, with lens paper underneath.

2. Take a 50 L aliquot of collagen.

3. Add 1.6mL of 0.02N acetic acid into collagen gently (do not mix).

4. Add 400 L of the collagen solution to each well, coating the bottom of each

well by tilting gently.

5. Let the collagen solution sit in the trays for 1 hour at room temperature.

87

6. Harvest the cells as normal. Obtain final 16 x 104 cells in 3.2 mL total volume.

7. Aspirate the collagen solution from each well, and lines with 500uL of PBS, then

replace with 500 L of diluted cell solution in each well, swirling to ensure each

well is coated.

8. Place the trays in the incubator. Image the trays 12-20 hours after plating the

cells.

Polyelectrolyte multilayer surface coating 2.5.3

Plasma-etched glass coverslips can be coated with polyelectrolytes of

alternating charge: 2 mg/mL poly (ethyleneimine) (PEI), prepared from a stock

solution of 50% w/v in water, MW 750,000; and 2 mg/mL poly(acrylic acid, sodium

salt) (PAcr) solution, prepared from a stock solution of 40 wt % in water, average

MW~30,000 (Aldrich, standard reagent-grade polymers), using the procedure and

buffers described by Kartalov and coworkers.83

This procedure will produce a

coverslip surface terminating in a negatively charged carboxylate layer, useful for

immobilizing positively charged samples. To obtain the opposite charge, one need

only stop one step short.

1. Plasma-etch # 1 coverslips for 30 minutes.

2. Load plasma-etched glass coverslips into microslide mailers from Thomas

Scientific Inc.

3. Immersed in solutions of the positive (PEI or PAll) polyelectrolytes for 10 min.

4. Immersed in solutions of the positive (PAcr) polyelectrolytes for 10 min.

5. Repeat steps 3 and 4 ten times.

6. Rinse coverslip with nanopure water. Slides can be stored in nanopure water for

weeks.

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95

3.SINGLE-MOLECULE MOTIONS OF OLIGOARGININE

TRANSPORTER CONJUGATES ON THE PLASMA

MEMBRANE OF CHINESE HAMSTER OVARY CELLS

96

Abstract

To explore the real-time dynamic behavior of molecular transporters of the

cell-penetrating-peptide (CPP) type on a biological membrane, single fluorescently

labeled oligoarginine conjugates were imaged interacting with the plasma membrane

of cultured Chinese hamster ovary (CHO) cells. The diffusional motion on the

membrane, characterized by a single-molecule diffusion coefficient and residence

time(R), defined as the time from the initial appearance of a single-molecule spot on

the membrane (from the solution) to the time the single molecule disappears from the

imaging focal plane, was observed for a fluorophore-labeled octaarginine (a model

guanidinium-rich CPP) and compared with the corresponding values observed for a

tetraarginine conjugate (negative cell entry control), a lipid analog, and a fluorescently

labeled protein conjugate (Transferrin-Alexa594) known to enter the cell through

clathrin-mediated endocytosis. Imaging of the oligoarginine conjugates was enabled

by the use of a new high-contrast fluorophore in the dicyanomethylenedihydrofuran

(DCDHF) family, which brightens upon interaction with the membrane at normal

oxygen concentrations. Taken as a whole, the motions of the octaarginine conjugate

single molecules are highly heterogeneous, and cannot be described as Brownian

motion with a single diffusion coefficient. The observed behavior is also different

from that of lipids, known to penetrate cellular membranes through passive diffusion,

conventionally involving lateral diffusion followed by membrane bilayer flip-flop.

Furthermore, while the octaarginine conjugate behavior shares some common features

with transferrin uptake (endocytotic) processes, the two systems also exhibit very

dissimilar traits when diffusional motions and residence times of single constructs are

compared. Additionally, pretreatment of cells with cytochalasin D, a known actin

filament disruptor, produces no significant effect, which further rules out unimodal

endocytosis as the mechanism of uptake. Also, the involvement of membrane potential

in octaarginine-membrane interaction is supported by significant changes in the

motion with high [K+] treatment. In sum, this first study of single transporter motion

on the membrane of a living cell indicates that the mode by which the octaarginine

transporter penetrates the cell membrane appears to either be a multi-mechanism

97

uptake process or a mechanism different from unimodal passive diffusion or

endocytosis.

This work was originally published in J Am Chem Soc. 2008 Jul 23;130(29):9364-70,

by H-L. Lee, E. A. Dubikovskaya, H. Hwang, A. N. Semyonov, H. Wang, L. R. Jones,

R. J. Twieg, W. E. Moerner, and P. A. Wender. The work represented a collaborative

effort between the Moerner and Wender labs, using a fluorophore synthesized by A.

N. Semyonov and H. Wang in the Twieg lab at Kent State. The fluorophore was

covalently attached to the cell-penetrating peptides by Elena Dubikovskaya in the

Wender lab. The initial imaging was performed by Dr. H. Hwang in the Moerner

laboratory, and H.-L. Lee performed the cell culture, subsequent imaging, and all

analysis.

http://www.ncbi.nlm.nih.gov/pubmed/18578528

98

Introduction to cell penetrating peptides 3.1

Biological membranes have evolved in part to protect the cellular interior by

preventing xenobiotics from passively entering cells.1 The cellular plasma

membrane that envelops the cell is highly effective as a controlled and selectively

permeable barrier; the intact cellular plasma membrane is essential to cell survival and

function. However, the plasma membrane also represents a major obstacle for

intracellular delivery of various cargo of interest. Since therapeutics, reporter

molecules, and imaging agents often require intracellular access to function

effectively, understanding and developing strategies for membrane penetration is

necessary.

Indeed, small-molecule drug design and selection have historically been

dominated by the conventional but reasonable view that only compounds that adhere

well to Lipinski’s “Rule of Five” can be bioavailable, i.e., cross the membrane barrier

to enter the cytoplasm. The five (four) requirements are: 1) Molecular mass smaller

than 500 Da; 2) High lipophilicity, expressed as cLogP greater than 5 (see below); 3)

More than 5 hydrogen bond donors, 4) More than 10 hydrogen bond acceptors. In

Lipinski’s Rule of Five, the partition coefficient of a molecule in octanol versus water

is noted to be particularly important.2, 3

⁄ (

[ ] [ ]

)

Equation 3-1

Compounds with a certain log P should be appropriately solubilized in the

mostly polar, water-based biological fluids of the animal body and at the same time

would also be able to pass through the relatively non-polar membrane of a cell. By the

log P metric, the use of many polar (e.g., nucleic acids, siRNA, etc.) and non-polar

(e.g., taxol, camptothecin, etc.) drug molecules would thus be prohibited or require

that additional effort be taken to tackle formulation, distribution or bioavailability

problems arising from their inherent inability to pass membranes.

While chemical agents can sometimes be synthetically modified to improve

their membrane permeability and cellular uptake, this approach often requires much

99

molecular alteration before membrane penetration is achieved. Alternatively, if a

modular synthetic molecular transporter can be utilized to promote cellular uptake of

cargos without other serious alternations to the original molecule, this may potentially

be employed for a wide range of applications.

Nature has evolved a number of ways for plasma membrane translocation;

numerous organisms have developed proteins, many of which are transcription factors,

that breach the eukaryotic plasma membranes through a variety of mechanisms.4 The

human immunodeficiency virus type-1 (HIV-1) transactivator of transcription (Tat)

protein, for example, when used in vitro, rapidly enters the cytosol (and nucleus) of a

wide variety of cells by endocytosis.5 Since 1988, numerous cell penetrating peptides

(CPPs) have been designed based on the Tat-protein, including polyarginine,

penetratin and other arginine-rich peptides.6 This rich class of molecular transporters

is an attractive toolkit for intracellular delivery of substances with low membrane

permeability, such as larger proteins, oligonucleotides, liposomes, certain classes of

small molecule drugs, and non-covalent supra-molecular complexes.7-10

Interestingly,

the nine amino acid peptide required for the cellular uptake of HIV tat, residues 49–57

(RKKRRQRRR), appears to utilize an additional mechanism compared to the

complete Tat protein.11

The uptake process for the 9 amino acid sequence remains

effective at 4 °C, and is therefore is cell energy-, and endocytosis-independent. This

uptake process has been shown to be sensitive to the attached cargo size and

composition and even cell type. The promise of Tat-like proteins as drug delivery

vectors has inspired a community of scientists to further study and develop cell

penetrating peptides based on the Tat template.

100

Table 3-1 Amino acid sequences of a sampling of arginine-rich cell penetrating

peptides

Immediately clear from Table 3-1, which summarizes recent efforts in CPP

development, is that the amino acid arginine is a key motif in CPPs. Studies by

Wender et al. demonstrated that the positively-charged guanidinium head groups of

Tat 49–57 are the key functionalities required for its entry into cells.11

Replacement of

all non-arginine residues in Tat 49–57 with arginines provides molecular transporters

that exhibit superior rates of uptake compared to the original sequence. Further,

Wender et al. also showed that while the positive charges from the guanidinium

groups are necessary, they are not solely sufficient, as is evident from the

comparatively poor cellular uptake of lysine nonamers.8, 9

These prior studies point to

the importance of bi-dentate hydrogen bonding of the cationic guanidinium groups to

anionic cell surface groups (phosphates, carboxylates, sulfates on heparins and

proteglycans). This theory is further corroborated by the important finding that mono-

or dimethylation of the critical guanidinium groups, while preserving charge, reduces

or completely eliminates cellular uptake. Lastly, it has been shown that the specific

number of intact arginines, and therefore guanidinium groups, is also important, with

7–15 arginines being the optimal length range.8, 12

Synthetic ease is a major consideration in strategizing drug delivery design. In

this respect, the CPP class of molecular transporters show great promise. For one,

natural peptide backbone chirality is not critical for cellular uptake of CPPs, as the

artificial peptides show increased level of cellular uptake relative to the natural

peptides. In addition to this, CPP class of molecular transporters also exhibit several

Cell-penetrating peptide Amino acid sequence

Tat49–57 RKKRRQRRR

Polyarginines RRRRRRRRR (R9)

R9F2RRRRRRRRRFF

Decalysine KKKKKKKKKK (K10)

Penetratin RQIKIWFQNRRMKWKK

Transportan GWTLNSAGYLLGKINLKALAALAKKIL

HIV-Tat derived PTD4 YARAAARQARA

101

other advantages for molecular design. First, studies have demonstrated that the

position of cargo attachment on guanidinium-rich peptids and length of the side chains

can be altered significantly without disrupting the efficiency of cellular uptake.13

Second, further studies also point to parameters that could be optimized to increase

cellular uptake. For example, changes in the backbone composition and in the side

chain spacing between residues can both lead to higher efficiency in cellular

delivery.14

These efforts report highly-branched guanidinium-rich oligosaccharides

and dendrimers as excellent molecular transporters.8, 9, 15-17

The molecular

conformational flexibility in highly branched dendrimeric structures has been

speculated to be responsible for the improved cellular delivery efficiency.

Interestingly, this finding is in direct contrast to receptor-mediated endocytotic uptake

(i.e. clathrin-mediate endocytosis), where specific structural conformation and specific

cellular receptors are necessary for receptor recognition and endocytosis. In fact, given

the potential and interest centered around CPP molecular transporters, the cellular

internalization mechanism of these peptides in cells still remains largely controversial,

and specific measurements to clarify some of the issues involved is a key goal of this

research. In the following, we summarize several leading models of CPP association

with and translocation through the plasma membrane.

102

Figure 3-1 Mechanisms of peptide uptake across the cellular membrane. A variety

of internalization mechanisms have been proposed to explain cellular uptake of CPPs.

These mechanisms include well-characterized energy-dependent pathways, based on

vesicle formation and collectively referred as endocytosis, and direct translocation or

cell penetration models, which involve the formation of hydrophilic clusters/pores or

charge-dependent cell potential-mediated translocation.18

Reprinted with permission

from Ref.18.

Figure 3-1 illustrates several proposed mechanisms that could accommodate

the above structure-function relationships for arginine-rich cell-penetrating peptides

(CPPs) and, more generally, guanidinium-rich molecular transporters (GRTs).9, 19

These mechanisms have been reported mostly separately but in principle may operate

concurrently depending on, for example, the properties of the CPPs and cargos, CPP

concentrations, the choice of selected model cell lines, and specific assay conditions.

Energy-driven clathrin-mediated endocytosis, the predominant form of cellular

endocytosis, and macropinocytosis, cellular drinking of a large volume of fluid

involving f-actin polymerization-driven membrane ruffling, have been proposed and

appear to be pertinent to entry involving high molecular weight CPP conjugates.13, 19,

20 On the other hand, lipid-like passive diffusion through the plasma membrane as a

103

membrane interaction has also been proposed, but is seemingly difficult to reconcile

with the polarity of the arginine oligomers (which are highly water-soluble) and the

dependency of cellular uptake on the specific number of positive charges.

Additionally, prior studies have suggested that an “adaptive translocation” mechanism

may be operative for low molecular weight guanidinium-based conjugates.8, 14, 21

In

this proposed model, positively charged guanidinium oligomers, which alone are too

polar to enter a membrane, form bi-dentate hydrogen-bonded, ion pair complexes with

complementary-charged (negative) cell surface functionalities of membrane-

embedded groups (carboxylates, sulfates, phosphates on cell surface heparins and

proteoglycans).21, 22

These polyarginine-glycan complexes are then driven inward

across the membrane under the influence of the membrane potential. It is especially

noteworthy with respect to this mechanism that water-soluble arginine oligomers can

be completely solubilized in octanol (a membrane dielectric mimic) by treatment with

an equivalent of a fatty acid salt (sodium laurate).21

104

Figure 3-2 Representative associations of a polycationic guanidinium transporter

with anionic cell membrane constituents. Depiction of adaptive translocation of

polyarginine after membrane association (bottom left), and octanol–water partitioning

(bottom right).23

Reprinted with permission from Ref.23.

Design and application of several CPPs have been reported to be fruitful and

these CPPs have successfully carried exogenous molecules into cells.8, 12, 17, 19

Understanding the internalization mechanism of peptide transporters is fundamental to

their use as delivery vectors for drugs and probes especially in view of the increasing

105

interest in the delivery of biological macromolecules (peptides, proteins, nucleic acids,

etc.). In view of the standing questions surrounding the uptake mechanism of arginine-

rich CPP peptides,13, 20, 24-28

we decided to examine by direct observation in real time

their membrane interactions on a single-molecule level in living cells.

Single-molecule imaging with single-particle tracking has been used to probe

the dynamics of molecules as well as their local environments in liquids, solids,

surfaces, and living cell membranes.29-35

Unlike assays used in previous studies, where

every polyarginine molecular transporter is assumed to be undergoing exactly the

same process of cellular uptake, and where the inference drawn is based on the

ensemble average, single-molecule techniques can be used to directly observe the

possibly unique motion of individual oligoarginines in real time. Oftentimes,

attempting to indirectly define population behavior based solely on an average can be

extremely difficult and complex. Mechanistic conclusions extracted from ensemble

averages alone would be open to a number of different interpretations particularly

when multiple mechanisms may be operating concurrently.20, 26, 28

This work is the

first report describing the direct observation of the motion of CPPs on the plasma

membrane in real time on a single-molecule level in living cells. The Zhuang group at

Harvard has explored the related question of virus cell-entry and cell-surface

proteoglycan endocytosis. 36, 37

In this investigation, epi-fluorescence imaging is used to study the movement

of single fluorophore-labeled octaarginine molecular transporters (Arg8-DCDHF-V)

using the cell membrane of living Chinese Hamster ovary (CHO) cells as a model

biological plasma membrane.34, 38-4041

The diffusional motions of single octaargine

transporters were then compared to the diffusional motions of transferrin-Alexa594, a

fluorescently labeled protein that undergoes clathrin/receptor-mediated endocytosis,

DCDHF-V-12, a fluorescent lipid mimic that intercalates into the phospholipid bilayer

and passively diffuses, and tetraarginine molecular transporters (Arg4-DCDHF-V), a

negative control for cellular entry. We also explored the effects of different treatments

on the diffusional motions of single octaarginines. These treatments include the

following: alteration of the cellular potential (extracellular high [K+] treatment),

106

inhibition of macropinocytosis (amiloride treatment),25, 26, 42

inhibition of cytoskeleton

polymerization to eliminate endocytosis (cytochalasin D), and facilitation of

octaagarinine cluster formation to promote endocytosis (presence of excess unlabeled

octaarginine).

Herein, we used the DCDHF-V fluorophore as a fluorescent tag for the

octaarginine transporters; DCDHF-V belongs to a class of single-molecule

fluorophores consisting of an amine donor and dicyanomethylenedihydrofuran

acceptor linked by a conjugated unit (e.g., benzene, naphthalene, styrene, etc.). More

on the molecules in the DCDHF class can be found in Chapter 4; briefly, DCDHF

fluorophores exhibit useful properties for single-molecule studies such as high

quantum yields, photostability, charge neutrality, and environmental reporter

functions.43-48

DCDHF-V, with the central phenyl conjugation extended by an

additional vinyl group, absorbs and emits at relatively long wavelengths (λex = 610

nm, λem=630 nm in water), thereby avoiding cellular autofluorescence. In this work, a

maleimide derivative of DCDHF-V is covalently attached to the octaarginine

backbone through an N-terminal cysteine and, as such, it reports on the mobility of the

resultant octaarginine peptide conjugate (structure, see Figure 3-3A). Similar to other

fluorophores in the DCDHF class, DCDHF-V is brighter when the molecule is

interacting with the more constrained environment of the membrane (and the cell

interior) compared to the aqueous buffer outside the cell, hence contrast in single-

molecule imaging is enhanced (see Chapter 4). Moreover, from DCDHF-V we

obtained good single-molecule signal-to-background ratios on the plasma membrane

under physiological oxygen concentrations, and the image contrast is observed to be

superior to that of the Cy3 fluorophore with or without oxygen scavenging systems

(see Section 2.5.1 for more details). From the obtained single-molecule trajectories we

obtain two quantities: the translational 2D diffusion coefficient, and the residence time

(the time that a fluorescent single molecule remains visible in the plane of the plasma

membrane.)

Our results indicate that the behavior of octaarginine on the cell surface is

different from that of lipids, known to associate with cellular membranes through

107

passive diffusion, exhibiting lateral diffusion sometimes followed by membrane flip-

flop. Furthermore, while the octaarginine behavior shares some common features with

protein uptake (endocytotic) processes, in that a subpopulation exhibits very low

diffusion coefficients, it also exhibits dissimilar diffusional properties in that another

subpopulation exhibits much larger diffusion coefficients. From our observations, we

conclude that the mode by which octaarginine penetrates the cell membrane appears to

either be a multi-mechanism uptake process or a mechanism different from passive

diffusion and endocytosis. These results have relevance to the understanding the

mechanism and optimization of cellular uptake of guanidinium-rich transporters

conjugated to small molecules, drugs and probes (MW ca. <3000).

Experimental 3.2

Fluorescent Conjugates 3.2.1

Arg8-DCDHF-V, Arg4-DCDHF-V, DCDHF-V-12 (further abbreviated as D-

V-12), and Transferrin-Alexa594 are the fluorescent conjugates used for this single-

molecule study. Arg8-DCDHF-V is the primary octaarginine molecular probe under

scrutiny; Arg4-DCDHF-V is the negative control, as Arg4 has been shown to be

insufficient for cellular uptake; D-V-12 is a fluorescent lipid analog, used for

mimicking passive lipid diffusion; Transferrin-Alexa594 is known to enter the cellular

interior via receptor- and clathrin-mediated endocytosis. The preparation of the

DCDHF-V maleimide, its Arg8 and Arg4- conjugates, and the lipid analog D-V-12 are

detailed in the supplementary online material of the published manuscript.49

The

synthesis of DCDHF-V has been previously reported.41, 46, 50

The transferrin-Alexa594

conjugate was purchased from Molecular Probes.

108

Figure 3-3 A. Structures of DCDHF-V labeled octaarginine (Arg8-D-V) and

tetraarginine (Arg4-D-V) B. Structure of DCDHF-V labeled lipid analog (D-V-12). Reprinted with permission from Ref.49. Copyright 2011 American Chemical Society.

Epi-illumination for single-molecule imaging 3.2.2

Both white-light transmission images of the cells and epi-fluorescence images

of single molecules were acquired using an inverted microscope (IX71, Olympus,

Center Valley, PA). Bright field white light illumination from a condenser lens

allowed the direct visualization of the edges of the cells. The fluorescence imaging of

the cells was performed with wide-field epi-illumination in an area of ~ 15 µm 15

µm (see Figure 3-4). For more details on the specific requirements and

recommendations for instrumentation, please see Chapter 2.

He-Ne laser illumination at 594 nm provided an intensity of ~ 0.14 kW/cm2 at

the sample focal plane. The resulting epi-fluorescence was collected with a 100

magnification, 1.3 NA, oil-immersion objective (UPlanApo, Olympus, Center Valley,

PA) and imaged through a 620 nm long-pass filter and a 610 nm dichroic mirror

(Omega Optical Inc., Brattleboro, VT) on an EMCCD-camera (IXonDV887, Andor,

South Windsor, CT). The pumping beam intensity (~ 140 W/cm2) was selected to

achieve acceptable signal-to-background ratio (~4-6) while extending the time before

photobleaching of the fluorophores to obtain the longest single-molecule trajectories

possible. The total number of detected photons from a single molecule was determined

by integrating all detected signal counts from a single spot and then computing the

total detected photons taking into account the known detector quantum efficiency and

measured multiplication gain.

109

Figure 3-4 Schematic of the imaging arrangement. A. Epifluorescence imaging

setup, B. CHO cells were cultured in fibronectin-coated imaging chambers and

examined in an inverted epifluorescence microscope. The focal plane of the

microscope is indicated. Reprinted with permission from Ref.49. Copyright 2011

American Chemical Society.

Cellular sample preparation 3.2.3

For details of cell culture and cellular sample preparation see Chapter 2.

Briefly, Chinese Hamster Ovary (CHO) cells were grown in RPMI 1640 phenol red-

free media (Gibco BRL, Grand Island, NY) supplemented with 10% fetal calf serum

(HyClone, Logan, UT), 10mM HEPES. (4-(2-hydroxyethyl)-1-

piperazineethanesulfonic acid), 1mM sodium pyruvate, and 0.1mM nonessential

amino acids, 100units/ml penicillin, 100µg/ml streptamycin, 10µg/ml gentamicin,

0.5mg/ml geneticin (Gibco, BRL, Grand Island, NY), and kept in 5% carbon dioxide

incubators at 37 ºC. Cells were seeded at 30% confluency and grown on chambered

coverglass for 1 day before imaging (Nalge Nunc International, Naperville, IL).To

facilitate optimum adhesion of cells, the coverglass was coated with 50ug/ml

fibronectin (human plasma, CalBiochem, San Diego, CA) in Dulbecco phosphate

buffered saline, DPBS, pH 7.4 (Gibco BRL, Grand Island, NY) for 1hr at room

temperature prior to deposition of cells to the chambered coverslips.

Cells were imaged at 22 ºC in a supplemented PBS buffer (see Chapter 2 for

details), while treatments with different small-molecule drugs or buffers, prior to

110

imaging, were done at 37 ºC. Unless mentioned otherwise, 1 nM of Arg8-D-V

conjugates in PBS buffer was applied to the CHO cells for observing single molecules.

Imaging was performed within 1 hour after removing the cell tray from the 37 ºC

incubator to ensure the viability of the cells. During imaging, both the fluorescent

conjugates and the inhibitors/drugs remained in solution.

Imaging Arginine8-D-V tethered to polyelectrolyte multi-layers 3.2.4

Polyelectrolyte multilayers (PEM) were used to immobilize Arg8-D-V in an

aqueous environment in order to mimic the binding of Arg8-D-V on the CHO plasma

membrane. Sequential deposition of polyacrylic acid (PAcr, Aldrich Chemical,

Milwaukee, WI, USA) and polyethyleneimine (PEI, St. Louis, MO, USA) has been

shown to reduce non-specific binding of proteins to glass slides51

while the uppermost

negatively charged PAcr layer immobilizes the cationic Arg8-D-V via electrostatic

interactions without dissociation or movement during imaging. The immobilized

Arg8-D-V on PEM surface were then imaged in imaging buffer (See Chapter 2 for

details on imaging buffer) exactly the same way as the cellular sample. This

experiment, designed as a control, was used to show that the time to photobleaching,

at the excitation conditions used, was longer than the time we were able to observe

single Arg8-D-Vs associated with the cellular plasma membrane. See Section 2.5.3 for

results and analysis.

Passive diffusion inquiry 3.2.5

D-V-12 (Figure 3-3B), the didodecylamine analog of the diethylamine

containing DCDHF-V, was stored in chloroform (1 mg/ml, stock) has been used

extensively in previous plasma membrane diffusional studies as a lipid analog.34

Here

we use D-V-12, as a model probe for passive diffusion compared to other types of

endocytosis. Immediately before use, 1-5 µL of D-V-12 stock solution in DMSO was

dried into a film and then reconstituted in 20 – 100 µL of ethanol. CHO cells were

incubated with a final concentration of 100 nM-1 µM of D-V-12 for 10-20 min at 37

ºC in supplemented RPMI 1640 media with FCS. The maximum concentration of

111

ethanol during incubation was 1% v/v to avoid any deleterious effect on the cellular

morphology.

Adaptive translocation inquiry 3.2.6

The cellular membrane potential stems directly from the net effects of ion

channels/pumps in the membrane, which, by establishing different concentrations of

ions (i.e. electrical charge) on the two sides of the membrane, is responsible for the

membrane potential. In non-excitable cells such as CHO (epithelial) cell, the

membrane potential is held stably, at what is termed the resting potential.

Since the cellular membrane potential is strongly implicated in the adaptive

translocation mechanism for internalization of polyarginines, we sought to investigate

the effects of membrane potential on the diffusional behavior of single polyarginines.

High potassium concentration buffer incubations (high [K+], 140 mM in PBS) were

used to reduce the membrane potential, a treatment which prevented poly(arginine)

cellular entry in previous bulk experiments.19

Receptor-mediated endocytosis inquiry 3.2.7

Transferrin is an iron-binding blood plasma glycoprotein, with an 80kDa

molecular mass. When a transferrin protein encounters a membrane-spanning

transferrin-receptor on the surface of a cell, binding of the transferrin to the receptor

occurs readily. After this binding occurs, a signal is sent through the membrane,

leading to clathrin coating the membrane receptor, and formation of a membrane

invagination. The transferrin and its receptor are then opsonized in clathrin-coated

vesicles. Once opsonization occurs, the clathrin coating surrounding vesicle disperses

(a prerequisite for the vesicle to fuse with other membranes) and individual vesicles

fuse to form the early endosome.52

The topic of receptor-based, clathrin-mediated endocytotis as a cellular uptake

mechanism is a highly controversial one in the CPP community. It has been reported

that in HeLa cells, fluorescently labeled CPPs co-localize with transferrin, indicating

endocytosis. Others have reported that the opposite was true- fluorescently labeled

112

octaarginine conjugates do not co-localize with transferrin. Other experimental

attempts to tease out this information were also met with disparate results. For

example, inhibition of endocytosis affects the process of CPP internalization to vastly

different degrees. For example, under inhibition of endocytosis, a Tat-avidin conjugate

shows only a small decrease in uptake.53

This result suggests that other mechanisms,

in addition to endocytosis, might be at play. In other studies, authors report that

treatment with the endocytosis inhibitor chlorpromazine results in a 50% decrease of

uptake of fluorescently-tagged Tat, also implicating other cellular uptake mechanisms

at play.54

In our experiments, Transferrin, as fluorescently-labeled by Alexa594, are used

on the CHO plasma membrane under conditions that permitted endocytosis (i.e. 22ºC,

and no use of endocytotic inhibitors). The single-molecule trajectories from

Transferrin-Alexa594 were compared with those of Arg8-D-V molecules. Imaging

conditions were the same as for Arg8-D-V on CHO cells, except that the concentration

of transferring-Alexa594 was 0.01 nM for tracking individual molecules. It should be

noted that it would have been helpful to perform experiments at 4°C where

endocytosis is suppressed, but this imaging condition would have been difficult to

implement and should be left for future studies.

Macropinocytosis inquiry 3.2.8

Other than receptor-based, clathrin-mediated endocytosis, a great deal of

current attention is also being focused on macropinocytosis as a specific pathway for

cellular update of octaarginines. Macropinocytosis is a particular type of endocytosis

that is mediated by lipid rafts and is clathrin-, caveolae-, and receptor-independent.

The specific process involves the generation of F-actin-containing membrane

protrusions which sweep inward and form fluid-filled vesicles called

macropinosomes.55

These vesicles are quite sizable and can often be greater than 1 μm

in diameter. Exploration of this endocytotic pathway is usually achieved via two major

means of negative control : 1) by pretreatment of cells with an F-actin

elongation/polymerization inhibitor, such as cytochalasin D, to inhibit

113

macropinocytosis; 2) by pre-treatment of cells with amiloride, an inhibitor of the

Na+/H

+exchange that is required in the process of macropinocytosis.

42

A number of research groups have proposed macropinocytosis as the main

mechanism of uptake for CPPs. Dowdy et al., have recently reported that both a Tat–

Cre fusion protein and FITC-labeled Tat enter CHO and HeLa cells through

macropinocytosis,56, 57

which is supported by the dose-dependent inhibition of uptake

observed when cells are pretreated with amiloride. In another mechanistic study,

Futaki et al. observed a considerable role for macropinocytosis in peptide uptake into

HeLa cells, finding that both cytochalasin D and the macropinocytosis inhibitor

amiloride significantly suppress uptake of the peptide into HeLa cells.26

In contrast to

the aforementioned groups, Shen et al. reported that the translocation of CPPs into

HeLa cells is not inhibited by amiloride pre-treatment or incubation at 16 °C, which

argues for an uptake mechanism distinct from macropinocytosis.25

This view is

supported with a subcellular fractionation method they developed to separate the

vesicular from the cytosolic compartments. Additionally, the authors found that co-

incubation with EGF, a known stimulator of macropinocytosis, does not significantly

increase the amount of oligoarginine found in the cytosol. 25

In this study we employ both of these treatments to query the role of

macropinocytosis in octaarginine internalization. The specific effect of cytochalasin D

has been described elsewhere.26, 28

In the cytochalasin D treatment, CHO cells were

treated by incubation with 10 µM cytochalasin D (Sigma) with for 30 min at 37˚C

prior to Arg8-D-V addition and imaging. Also, inhibition of macropinocytosis with

pre-treatment of amiloride was also used. Incubation with serum-free cell growth

medium containing 100 µM amilioride for 60 min at 37ºC followed by washing with

PBS buffer was employed prior to Arg8-D-V addition and imaging.25, 26

High octaarginine concentration inquiry 3.2.9

In another model for the cellular uptake of CPPs, proponents postulate the

formation of octaarginine clusters prior to association with the plasma membrane at

high CPP concentration. It is suggested that the size of these putative clusters, similar

114

to larger biological macromolecules (such as EGF) may induce nonspecific energy-

and receptor- dependent endocytosis by their sheer size. To further investigate this

model mechanism, we conducted an experiment with a high concentration of

octaarginines. At the much higher concentration of octaarginines used, to enable

single-molecule imaging, only a small fraction (0.01%) of the octaarginines used were

fluorescently labeled. Specifically, unlabeled octaarginines (Arg8 peptide, at 10 µM)

were mixed with Arg8-D-V (1 nM) in buffer, and were applied together to the CHO

cells for imaging.

Effects of addition of oxygen scavenger on imaging 3.2.10

In optimizing the imaging conditions to yield the best signal-to-background

ratio and longest length of single-molecule trajectories, the use of an oxygen

scavenging system was investigated. A standard enzymatic oxygen scavenger system

was prepared: 1% v/v glucose (Sigma; 500 mg/ml stock), 1% v/v glucose oxidase

(Sigma; 5000 U/ml stock), 1% v/v catalase (Sigma; 40,000 U/ml stock) and 0.5% v/v

2-mercaptoethanol (Sigma; 14.3 M stock). Covalent attachment of the DCDHF-V-

NHS fluorophore to the chaperonin protein GroEL (using methods described in the

Chapter 2) enabled observation of singly labeled proteins in an agarose gel, where the

average photobleaching time was observed to increase by roughly a factor of 3 in low

oxygen. At the same time, greatly increased blinking was observed with 100 ms time

resolution. More importantly, cellular imaging of Arg8-D-V with the enzymatic

oxygen scavenger present in the solution above the cell (Figure 3-5B) showed a far

higher fluorescence background than without the oxygen scavenger system (Figure 3-

5A), implying that quenching of the emission from Arg8-D-V molecules in solution

was suppressed in anoxic conditions. In other words, at normal oxygen

concentrations, the Arg8-D-V complexes on the cell membrane were much more

visible than those in the solution above the cell. Therefore, all the data reported in this

study were obtained without the use of an oxygen scavenger system.

115

Figure 3-5 Effect of the enzymatic oxygen scavenger system for single conjugate

cellular imaging showing high contrast from the DCDHF label at normal oxygen. Reprinted with permission from Ref.49. Copyright 2011 American Chemical Society.

Single-molecule fluorescence imaging of positional trajectories 3.3

The fluorescence from individual molecules was recorded as described in

3.2.2. Except in the case of D-V-12 (20 ms) and transferrin-Alexa594 (100 ms), all

movies were recorded at 50 ms integration time with continuous illumination. Laser

light at 532 nm was used for imaging of D-V-12, while 594 nm laser light was used

for imaging of Arg8-D-V, Arg4-D-V, and transferrin-Alexa594.

CHO cells adhere well to the glass surface of the fibronetin-coated chambered

coverglass, and become spindly-shaped with dimensions of ~30 x 10 x 5 (L×W×H)

µm. Thus, the bottom and the top portions of the plasma membrane are roughly

parallel to the focal plane of the microscope and can be treated as two-dimensional

planes (see Figure 3-4B). Near the edges of the cell, out-of-focal-plane diffusion can

occur, and could be detected by an increase in the spot size. It is well-known that the

bottom membrane of the cell contains adhesions that could restrict or perturb the

trajectories of single molecules. Therefore, only single molecules located on the

upper surface of the cell and away from cell edges were included in the analyses.

The successive (x,y) positions of the single molecules on the plane of the cell

surface were recorded as a function of time. In order to investigate the behavior of

116

single conjugates in a consistent fashion, only the fluorescent spots which appeared

after the start of illumination and disappeared before the end of recording were

considered. The residence time (τR) is defined as the time of visibility from the initial

appearance of a single-molecule spot on the membrane (from the solution) to the time

the single molecule disappears from the imaging focal plane in a single frame. The

diffusion coefficients for each trajectory were extracted as described in section 3.4

below.

Analysis of single-molecule motion 3.4

Trajectories of single conjugates were extracted by using a Single Particle

Tracking (SPT) program from ETH-Zurich.58

In this approach, particle positions were

iteratively refined by using the intensity centroid for sub-pixel interpolation.

Trajectories were computationally extracted from the recorded movie sequences using

a feature-point tracking algorithm. All trajectories were also visually inspected to

ensure that the tracking program was operating properly.

The mean-squared displacement from each individual single-molecule

trajectory (truncated to 10 time steps, typically at 0.050 s per time step) was used to

extract an observed diffusion coefficient D using

1

1

4][][][][m

Dtmnynymnxnx

Equation 3-2

where ][nx x position at time n, ][ny y position at time n, and

]1[][][ nxnxnx , ]1[][][ nynyny , with the fixed time lag t = 50 ms.

This method has the virtue that it removes statistical bias from the exposure time as

well as the error from the positional measurement. See Dr. Adam Cohen’s dissertation

for more detail. Each single-molecule trajectory analyzed yields an apparent diffusion

coefficient, D, which can be regarded as an estimate of the true diffusion coefficient of

the particles.59

An expected distribution of apparent diffusion coefficients for the

individual trajectories assuming an underlying homogeneous diffusion process was

generated to test for heterogeneity in the motion. This approach takes into account the

error in the estimates of D arising from the limited number of position measurements.

117

Under the homogeneity hypothesis that all molecules are moving as unconstrained

Brownian diffusers, the distribution of measurements of D follows:60

( )

( ) (

) ( ) (

)

Equation 3-3

where D is the diffusion coefficient extracted from an individual trajectory, D0 is the

underlying Brownian diffusion coefficient, and N is the number of time steps in each

trajectory. This expression was used to generate the smooth curves in the figures

below by fixing N=10 and varying Do in a nonlinear least-squares fit.

Results 3.5

Acquired as described in Section 3.2.2, the experimental data consist primarily

of high-speed microscopic imaging of the motion of single fluorescently-labeled

molecules on the plasma membrane of CHO cells. A schematic of the epi-

fluorescence imaging arrangement is shown in Figure 3-4A, emphasizing that the

focal plane of the microscope was most often located at the upper plasma membrane

to avoid artifacts arising from interactions with the borosilicate glass surface.

118

Fluorescently-labeled Arg8 conjugates are readily internalized, and 3.5.1

single copies can be observed interacting with the plasma membrane

Figure 3-6 Representative images of Arg8-D-V on the CHO plasma membrane with

decreasing concentration of conjugate (A B, and C). Representative trajectories of

single Arg8-D-V conjugates on the CHO plasma membrane (D). Reprinted with

permission from Ref.49. Copyright 2011 American Chemical Society.

Figure 3-6A shows a wide-field epifluorescence image of two CHO cells

which have been incubated with a high concentration of Arg8-D-V (5 µM). The focal

plane was placed near the center (equatorial plane) of the cell for this image only. The

bright fluorescent regions confirm that many internalized, labeled peptides are easily

seen in the cell interior, in various stages of interaction with organelles. The

constructs are also concentrated at the plasma and nuclear membranes, and do not

enter the nucleus. The ease of uptake of the Arg8 construct is consistent with earlier

reports.4-19

Figure 3-6B shows the upper plasma membrane where the cell has been

119

incubated with a lower concentration of Arg8-D-V (10 nM). Now the fluorophore-

labeled peptides appear as resolved single fluorescent spots. In Figure 3-6C, at 1 nM

incubation, resolved and isolated, individual molecules are clearly observed. These are

seen to disappear in a single frame and have brightness representative of single copies

of the DCDHF-V emitter. It is worth noting that the high contrast of this image is a

result of the use of the DCDHF-V label, which exhibits lower fluorescence intensity in

the lower viscosity environment of the buffer solution above the cell, yet becomes

bright in constrained and hydrophobic environments,34, 61

for example, when the Arg8-

D-V constructs are interacting with the plasma membrane.

The behavior of the DCDHF-V label also becomes apparent when the oxygen

concentration is lowered by an enzymatic oxygen scavenger system with β-

mercaptoethanol, often used to extend the photobleaching time of fluorophores (see

above). In experiments exploring the behavior in gels, mentioned in 3.5.1, the

photobleaching time of DCDHF-V increases, but severe blinking also occurs on the

100 ms time scale. More importantly, at reduced oxygen concentration the fast-

diffusing labeled peptides in solution above the cell now become visible as a

background haze (see section 3.2.10. and Figure 3-5). This is likely due to an

alteration in photophysical parameters (e.g. triplet lifetime) which should be examined

in a separate study. The net effect is that with lowered oxygen concentration, the

contrast of the single-molecule spots in the membrane is greatly reduced due to out-of-

focus fluorescence. Similar low-contrast images were also observed when the Arg8

peptide was labeled with Cy3, with or without oxygen scavenger. While background

fluorescence could be reduced by confocal imaging or other methods, by using the

DCDHF-V label, the simplicity, parallelism, and speed of wide-field epifluorescence

imaging is maintained under conditions of normal oxygen concentration, with higher

contrast than is available with Cy3.

By acquiring images like Figure 3-6C at the frame exposure time of 20-100 ms

per image and extracting the positions of the single molecule spots as a function of

time, single-molecule position trajectories are obtained. Figure 3-6D shows some

representative examples, placed at different positions in the plane for clarity. This

120

motion is not due to molecules in solution above the cell, because it is much slower

than free conjugates. It is also well-known that cytoplasmic diffusion is in general

faster than in-membrane diffusion. For example, a cytoplasmic diffusion coefficient

for GFP has been reported62

to be ~ 8 µm2/s, much faster than the diffusion

coefficients we observe (vide infra). For these reasons, it is reasonable to regard the

trajectories in Figure 3-6D as in-membrane motion. However, since the focal depth of

the microscope is ~300 nm, we cannot completely exclude the possibility that some of

the observed motion represents motion in the cytoplasm below the membrane.

A goal of this work is to characterize these motions in a quantitative fashion

by analysis of the individual trajectories, and to compare these characteristics across

the panel of various transporters (octaarginine, tetraarginine, lipid, and the protein

transferrin). Qualitative examination of Figure 3-6D suggests that the motions

observed for single molecules of the octaarginine transporter are quite heterogeneous.

Some molecules move very little during the observation period, others appear to

undergo random motion over larger distances, and some make large initial

displacements before more random explorations (e.g. the molecule at x=9 µm and y=

2.5 µm). While this last behavior is quite intriguing and seemingly relates to models

of picket-fence membrane organization, it was relatively rare, and thus was not further

analyzed.

Single Arg8-D-V molecules disappear before photobleaching 3.5.2

After obtaining the single-molecule trajectories, the first quantity of interest is

the residence time τR in or near the plasma membrane, defined above. A molecule

might disappear for several reasons; namely, the destruction of the DCDHF-V label

due to photobleaching, motion out of the depth of focus of the microscope of ~300 nm

which can occur due to dissociation away from the cell surface, or internalization into

the cell.

To quantify the photobleaching behavior, single molecules of Arg8-D-V were

immobilized and imaged on a poly(electrolyte) multilayer (PEM) in (normally

oxygenated) buffer as an approximate model for the confined and charged membrane

121

environment. Under these conditions, the time-to-photobleaching, τB was recorded,

yielding the τB distribution in Figure 3-7A. For the same molecules, the total number

of detected photons from each emitter was determined by integration of the intensity

over a small area containing all the photons from the molecule, with background

subtracted; the average total detected photons for 46 molecules is 8.3+/- 1.1 x104. A

single exponential fit to the τB data, as is appropriate for (first-order) Poisson statistics,

is a reasonable fit, yielding an average τB of 6.9 +/- 1.2 s. While the DCDHF-V

molecules may last longer in a completely oxygen-free environment or in a polymeric

host,38

this measurement provides a reasonable assessment of the photo-stability of

Arg8-D-V under our aqueous, oxygenated cellular imaging conditions.

In contrast, Figure 3-7B shows the observed residence time τR for single

molecules of Arg8-D-V on the CHO cell membrane. Each single molecule was

followed frame-by-frame as it moved in order to record the total number of detected

photons and to determine the total time before disappearance. A double exponential

fit to the τR distribution yields characteristic times of 1.16 ± 0.07s and 3 ± 1 s; under

these conditions the average number of detected photons per emitter is 4.4 ± 0.61

×104. The trajectories that contribute to the long-time tail of the τR distribution may

arise from photo-bleaching, but the faster component cannot be, and it is reasonable to

interpret the disappearance of single molecules in the fast population as dissociation or

internalization rather than photo-bleaching. While the fraction of CPPs actually

internalized was not quantitatively determined in this work, Figure 3-6A shows that

internalization readily occurs, which is consistent with earlier reports.8, 12, 13, 15-17, 19-21,

24-28, 63

122

Figure 3-7 Comparison of photobleaching time and residence time. A. Histogram of

photobleaching times for single Arg8-D-V on PEM. B: Histogram of residence times

for Arg8-D-V on CHO cells. The pump wavelength is 594 nm with a pumping

intensity of 140 W/cm2. Reprinted with permission from Ref.49. Copyright 2011


Length of single-molecule trajectories yields fluorescent probe residence 3.5.3

time

Residence time distributions, as shown in Figure 3-8, were also obtained for D-

V-12, transferrin-Alexa594, Arg4-D-V, and Arg8-D-V on CHO cells treated with high

[K+] buffer, and single-exponential behavior was observed for all cases except the last.

Given that the residence time of Arg8-D-V does not seem to be perturbed significantly

by the presence of high [K+] concentration buffer, one may infer that the association

of Arg-8-D-V with the plasma membrane is not perturbed by a change in the cellular

resting potential. On the other hand, a change in the number of arginines (i.e.

guanidinium groups) from eight (Arg8-D-V) to four (Arg4-D-V), reduces the

residence time by a factor of 2. This suggests that the positive charge of the

guanidinium head groups plays a significantly role in membrane interaction of the

arginine-rich CPPs. The charge-neutral lipid analog, D-V-12, only associates with the

membrane briefly, averaging a residence time of 0.54±0.01s. Tranferrin, the receptor-

mediated endocytosis model probe, maintains persistent close association with the

plasma membrane for up to 4.4s, much longer than the residence time of Arg8-D-V.

Lastly, an intact actin cytoskeleton is necessary in macropinocytosis. Interestingly,

123

treatment with cytochalasin D, the actin polymerization inhibitor, lengthens the

residence time of Arg8-D-V on average; this implicates actin elongation as a

contributor in octaarginine cellular uptake. Further discussion on the comparison of

single-molecule trajectories of the fluorescent probes on residence time and diffusion

coefficient can be found in Section 3.5.5.

Figure 3-8 Residence time distributions and fitted residence times of Arg8-D-V,

Arg8-D-V on high [K+] treated CHO cells, Arg4-D-V, D-V-12, Transferrin-

Alexa594, and Arg8-D-V on cytochalasin D treated CHO cells.Reprinted with

permission from Ref.49. Copyright 2011 American Chemical Society.

0 2 4 6 8 10

0

10

20

30

40

0 2 4 6 8 10 12 14

0

2

4

6

8

10

12

14

16

0 1 2 3 4 5 6

0

20

40

60

80

100

0 1 2 3 4 5 6

0

10

20

30

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50

0 2 4 6 8 10

0

20

40

60

80

100

120

0 2 4 6 8 10

0

20

40

60

80

100

120

R1: 1.16±0.01sec

R2: 3.03±0.85sec

Arg8-D-V Arg8-D-V, High [K+]

R1: 1.15±0.01sec

R2: 2.62±1.06sec

Arg4-D-V

R : 0.79 ± 0.01 sec

Transferrin-Alexa594

R : 4.04 ± 0.22 sec

Arg8-D-V, Cytochalasin D

D-V-12

R : 0.54 ±0.01sec

Occu

rren

ce

s

Residence Time (sec)

R1: 1.58±0.24sec

R2: 5.10±1.87sec

124

Diffusion coefficients of single-molecule trajectories 3.5.4

The positional trajectories of single molecules allow analysis of the motion to

determine if the constructs move randomly on the surface by unconstrained Brownian

motion, or move in some other way. To do this in an unbiased fashion, each trajectory

was truncated to 10 time steps so that all molecules would have the same trajectory

length, and diffusion coefficient values were extracted from mean-squared

displacements as described in Section 3.4, displayed in Figure 3-9. Since the

trajectories are finite in length due to experimental limitations, naturally there is a

statistical error in each D determination. To be as unbiased as possible, we compare

the measured D distributions with the homogeneity hypothesis described above in

Section 3.4, i.e., that all measured values arise from underlying random Brownian

motion with a single diffusion coefficient Do. Under this homogeneity hypothesis, the

measured single-molecule D values should scatter according to a well-known

probability distribution,33, 34, 60, 64

the smooth black curve in each panel. Departures

from the distribution indicate heterogeneous motion. Whether or not this hypothesis

was upheld, we also computed an ensemble average diffusion coefficient Davg from the

measurements.

125

Figure 3-9 Distributions of observed single-molecule diffusion coefficients (D) for

A: Arg8-D-V, B: D-V-12, C: transferrin-Alexa594 conjugate, and D: Arg8-D-V in

high [K+] on the CHO plasma membrane. Part C (inset) shows the fraction of

molecules below and above a cutoff of 0.2 µm2/s, for Arg8-D-V (red) and

transferrin-Alexa594 (green). Part D (inset) shows the fraction of molecules

below and above a cutoff of 0.08 µm2/s, for Arg8-D-V (red) and Arg8-D-V in high

[K+] (blue). Reprinted with permission from Ref.49. Copyright 2011 American

Chemical Society.

The membrane-associated motion of the Arg8-D-V construct was examined by

extracting distributions of single-molecule diffusion coefficients and residence times,

and these values were compared to those obtained from the following cases: (a) a

lipid-like derivative of the same fluorophore (D-V-12), expected to enter cells via

lateral passive diffusion followed by membrane flip-flop mechanism; (b) a labeled

protein (transferrin-Alexa594 conjugate) known to penetrate cells via clathrin-

mediated endocytosis; 19

and (c) Arg4-D-V, a water-soluble tetraarginine conjugate

that in contrast to octaarginine is incapable of penetrating or only poorly penetrates the

intact plasma membrane.8, 14, 21

Treatments of cells prior to imaging were also

126

performed with high [K+] PBS buffer (140 mM) that is known to reduce membrane

potential to close to zero and consequently to reduce uptake of octaarginine

conjugates. 8, 19

Treatment with amilioride, known to shut down macropinocytosis,25, 26,

42 and addition of excess unlabeled Arg8 were also performed.

Figure 3-9 compares observed diffusion coefficient distributions for the four

primary cases. The difference between the histograms and the solid curves indicates

that the motion of the molecules at our imaging rate is inconsistent with homogeneous

Brownian diffusion for all cases except for the lipid analog. Table 3-2 collects key

quantities for facile comparison in the following sections.

Table 3-2 Table of Values. Tabulated values for residence time, average diffusion

coefficients, and N (number of particles tracked) for transferrin-Alexa594 conjugate,

Arg8-D-V, Arg8-D-V in high [K+], Arg4-D-V, and D-V-12 on the CHO plasma

membrane. Reprinted with permission from Ref.49. Copyright 2011 American

Chemical Society.

The motion of Arg8-D-V on CHO cells differs from that of an endocytotic 3.5.4.1

protein conjugate, Transferrin-Alexa 594.

For the transferrin conjugate Figure 3-9C, the motion is clearly heterogeneous

indicating that a variety of diffusional behaviors are present, perhaps due to

endocytotic vesicles in various stages of formation. Heterogeneous motion occurs

even more dramatically for the Arg8-D-V conjugate (Figure 3-9A), where a long tail

to high D values is present. This may reflect the binding of the positively charged

guanidinium groups to varying numbers of lipids or proteoglycans with negatively

127

charged head groups. The average diffusion coefficient for Arg8-D-V (0.15± 0.20

µm2/s) is similar to that of MHCII transmembrane proteins in the CHO cell membrane

at 22C of ~0.2 µm2/s.

31

Comparing diffusion coefficients and τR values in Table 3-2, both the single

transferrin proteins and the Arg8-D-V molecules move much more slowly than the

lipid D-V-12. Transferrin shows even slower diffusion on the membrane with large

residence times, and is known to internalize by formation of a receptor-mediated

endocytotic vesicle.65

Moreover, the motion of Arg8-D-V is different from transferrin

in two ways. First, the residence time distribution shows shorter values for Arg8-D-V,

and second, the Arg8-D-V molecules have a longer tail to larger diffusion coefficients.

To show this more directly, we chose an arbitrary D value of 0.2 µm2/s, and show the

fraction of molecules below and above this value in Figure 3-9C inset. The CPP

conjugate has a much larger fraction of molecules with high D values.

The motion of Arg8-D-V on CHO cells differs from that of lipid analog, D-3.5.4.2

V-12.

In contrast, the motion of the lipid analog (Figure 3-9B) is closer to

homogeneous Brownian diffusion, as was observed earlier for other lipid-like DCDHF

derivatives.34

The average diffusion coefficient of 1.9 µm2/s is similar to that observed

for other lipids in the CHO cell membrane at 22 C.34

Because the lipids move

quickly, the residence times are quite small, mostly due to motion of the molecules out

of the observation region.

The Arg8-D-V construct associated with the plasma membrane more 3.5.4.3

strongly than Arg4-D-V

In previous studies, tetraarginine (Arg4) was shown to be much less efficient in

facilitating cell entry compared to octaarginine.8, 19, 21

In comparison to Arg8-D-V, the

Arg4-D-V residence times are much shorter when compared to that of Arg8-D-V. This

is consistent with a more rapid dissociation of Arg4-D-V from the cell surface due to

the fewer number of possible ionic interactions.

128

Changes in cellular potential affect Arg8-D-V diffusion 3.5.4.4

High [K+] in the extracellular solution is used to reduce the membrane

potential and cellular uptake of polyarginine.8, 19

The D distribution for Arg8-D-V

with this treatment is changed compared to that for the cells with normal cellular

membrane potential (Figure 3-9A, D). To explore the changes more carefully, we

have constructed normalized probability distributions from the two data sets, which

show a relative reduction of the species with lower diffusion coefficients and an

increase in the species with higher diffusion coefficients with high [K+] treatment

(plotted on the same figure in Figure 3-10). To illustrate this effect approximately, if

an arbitrary cutoff is selected, for example at 0.080 µm2/s, the fraction of molecules

above and below this value are shown in Figure 3-9A inset for the normal cell and for

high [K+]. It appears that the reduction in membrane potential causes Arg8-D-V to

move faster laterally on the membrane surface, and skews the distribution of Arg8-D-

V away from that for the endocytotic transferrin construct. However, the average

residence times of Arg8-D-V on the CHO cell plasma membrane are not changed

dramatically with high [K+] (Figure 3-10) suggesting that membrane potential does not

play a role in sustaining CPP-membrane association.

129

Figure 3-10 Normalized probability distributions for Arg8-D-V (black) and Arg8-D-

V on high [K+] buffer treated CHO cells (gray). Reprinted with permission from

Ref.49. Copyright 2011 American Chemical Society.

Endocytosis and macropinocytosis do not contribute significantly to CPP-3.5.4.5

membrane interaction

Treatment with a commonly-used suppressor of macropinocytosis, amilioride,

was also performed as previously discusses in addition to the cytochalasin D

treatment. However, multiple attempts to observe single molecules on the CHO

plasma membrane after amilioride treatment were not successful. This is not too

surprising, because amilioride treatment affects macropinocytosis through interference

with cellular processes, but also various ion channels, morphology, and membrane

potential.13

On the other hand, treatment of cells with cytochalasin D, a known actin

filament disruptor, was also performed to investigate the possible involvement of

macropinocytosis.25, 26

Under this treatment, the Davg and D distribution of Arg8-D-V

do not change significantly (Table 3-2). Also, the residence times are relatively close

to one another. To further explore the possible effects of cytochalasin D, normalized

distributions from the two data sets were also constructed and compared (Figure 3-11)

0.0 0.1 0.2 0.3 0.4 0.5

0.7

Fra

cti

on

No

rma

lize

d p

rob

ab

ilit

y d

istr

ibu

tio

n

Diffusion Coefficient (μm2/s)

Arg8-D-V Davg = 0.17 ± 0.01

Arg8-D-V, high [K+] Davg = 0.22 ± 0.01

Arg8-D-V Davg = 0.17 ± 0.01

Arg8-D-V, high [K+] Davg = 0.22 ± 0.01

0 0.08D (μm2/s) 0.08 m2/s0

130

as in the case of high [K+] buffer. Using an arbitrary cutoff at 0.050µm

2/s, the relative

fractions above and below this cutoff remain unchanged within statistical error by the

cytochalasin D treatment, unlike the case for high [K+] buffer. From the

aforementioned results, we infer that actin polymerization, a vital step in endocytosis,

does not play a major role in Arg8-D-V interactions with the plasma membrane in the

regime of nanomolar concentrations.

Figure 3-11 Normalized probability distributions for Arg8-D-V (red) and Arg8-D-V

on cytochalasin D treated CHO cells (purple). Reprinted with permission from


Changes in cellular potential affects Arg8-D-V diffusion 3.5.4.6

Finally, treatment of cells with an excess level of unlabeled Arg8 peptides

(~10µM), known to stimulate probe aggregation and possibly induce both endocytosis

and macropinocytosis,66

was performed. With this treatment, the D distribution for

Arg8-D-V skews slightly to lower diffusion coefficients (Figure 3-12). Also, the

residence time of Arg8-D-V is reduced almost by a factor of 2 by the treatment with

excess Arg8 peptide. In light of these findings, we infer that high concentration of

Arg8 ( ~10µM) may magnify one, or more, specific mode(s) of interaction with the

plasma membrane compared to nanomolar concentrations of Arg8. However, it is not

possible to distinguish at this point between possibly enhanced macropinocytosis or

0.0 0.2 0.4 0.6 0.8 1.0

Arg8-D-V Davg = 0.17 ± 0.01

Arg8-D-V , Cytochalasin D Davg = 0.16 ± 0.01

Arg8-D-V Davg = 0.17 ± 0.01

Arg8-D-V , Cytochalasin D Davg = 0.16 ± 0.01

Fra

cti

on

0

No

rma

lize

d p

rob

ab

ilit

y d

istr

ibu

tio

n

Diffusion Coefficient (μm2/s)

0.05µm2/s

0.8

0 0.05D (μm2/s)

131

more efficient penetration by complexes of Arg8 peptides. This effect should be

examined in future studies.

Figure 3-12 Distributions of single-molecule diffusion coefficients and residence

times. On the left, distribution of single-molecule diffusion coefficients (D) of Arg8-

D-V in presence of 10 µM of excess unlabeled Arg8 peptides; on the right, the

distribution of residence time (R) of Arg8-D-V in presence of 10 µM of excess

unlabeled Arg8 peptides. Reprinted with permission from Ref.49. Copyright 2011


Conclusions 3.6

Our results represent the first report where plasma membrane interactions of a

CPP are examined on a single-molecule level in living cells. This study was enabled

by the use of a new DCDHF high-contrast fluorophore, which brightens upon

interactions with the membrane and provides good contrast under oxygenated

conditions. The primary quantities extracted were the residence times of single

molecules on the membrane and the single-molecule diffusion coefficients; these

quantities cannot be extracted from previous ensemble measurements. Our findings

indicate that the motions of the octaarginine conjugate are highly heterogeneous, and

cannot be described as Brownian motion with a single diffusion coefficient. The

observed motions for the octaarginine constructs are different from that of lipids,

known to interact with cellular membranes through passive diffusion. Furthermore,

while the octaarginine behavior on the plasma membrane shares some common

features with transferrin uptake (energy-dependent endocytotic) processes, it also

132

exhibits very dissimilar traits when diffusional motions of single constructs are

compared. Not only were much larger diffusion coefficients observed for populations

of Arg8-D-V, but also their residence times were far smaller than those for transferrin.

On the other hand, suppression of the membrane potential by the high [K+] buffer

skews the Arg-8-D-V diffusion coefficient distribution to higher numerical values.

Furthermore, pretreatment of cells with cytochalasin D, a known actin filament

elongation inhibitor, does not change the diffusion coefficient distribution which rules

out unimodal endocytosis as the sole mechanism of uptake. Additionally, treatment

with excess unlabeled Arg8 peptides (10µM) favors macropinocytosis or aggregation-

enhanced uptake processes. In summary, our real-time, single-molecule analysis

suggests that the mode by which octaarginine conjugates penetrate the cell membrane

appears to either be a multi-mechanism uptake process or a mechanism different from

passive diffusion and endocytosis. These results have relevance to the mechanism of

cellular uptake of guanidinium-rich transporters conjugated to small molecules, drugs

and probes (MW ca. <3000). Specific membrane components that may aide in cellular

association of oligoarginines, and cellular treatment that might heighten cellular

uptake remain to be identified.

3.7 Acknowledgements

I thank Dr. Andrea Kurtz-Pomerantz for assistance with PEM slide

preparation, and Dr. Adam Cohen, Dr. Stefanie Nishimura, and Dr. So Yeon Kim for

helpful discussion and assistance. This work was supported in part by the National

Institutes of Health through the NIH Roadmap for Medical Research Grant No.

HG003638 (W.E.M., R.J.T.) and by Grant Nos. CA31841 and CA31845 (P.A.W.).

133

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139

4.SUPER-RESOLUTION IMAGING OF TARGETED

PROTEINS IN FIXED AND LIVING CELLS USING

PHOTOACTIVATABLE ORGANIC FLUOROPHORES

140

Abstract

Super-resolution imaging techniques based on sequential imaging of sparse

subsets of single molecules require fluorophores whose emission can be

photoactivated or photoswitched. Because typical organic fluorophores can emit

significantly more photons on average than fluorescent proteins, organic

fluorophores have a potential advantage in super-resolution imaging schemes,

but targeting to specific cellular proteins must be provided. We report the design

and application of HaloTag-based target-specific azido DCDHFs, a class of

photoactivatable push–pull fluorogens which produce bright fluorescent labels

suitable for single-molecule super-resolution imaging in live bacterial and fixed

mammalian cells.

This work was originally published in J. Am. Chem. Soc., 2010, 132 (43), pp 15099–

15101. The work represented a collaborative effort from the Moerner, Twieg, Rao,

and Shapiro labs, using a fluorophore synthesized by H. Wang in the Twieg lab at

Kent State University. The fluorophore was covalently attached to the HaloTag

targeting moiety by H. Xie in the Rao lab. Genetic constructs in bacterial and

mammalian cells were made possible by G. Bowman in the Shapiro group, and K.

Zhan in the Rao group, respectively. H.-L. Lee performed the cell culture, imaging,

and analysis assisted by S. Lord and S. Iwanaga in the Moerner Group.

141

4.1. Introduction

Requirements for pointillist super-resolution imaging 4.1.1

As mentioned in Chapter 1, fluorescence microscopy is an extremely powerful

tool for biological studies as it can noninvasively light up different subcellular

structures simultaneously with a wide spectrum of labels, high signal-to-background

ratio, and extremely precise specificity. On the other hand, the use of visible light

subjects the experiment to the cost of relatively poor spatial resolution when compared

side-by-side to other high-resolution microscopy techniques such as X-ray or electron

microscopy. More specifically, this is because optical diffraction limits the resolution

of an optical microscope to roughly the optical wavelength divided by twice the

numerical aperture of the imaging system, λ/(2NA).1 Since the highest values of

numerical aperture for state-of-the-art, highly corrected microscope objectives are in

the range of 1.3-1.6, the spatial resolution of optical imaging has, until recently, been

limited to ~180 nm for visible light.

The aforementioned resolution limit in far-field imaging has recently been

overcome in a number of experiments using two main groups of methods:

deterministic super-resolution, and stochastic (pointillist) super-resolution microscopy.

Deterministic super-resolution methods can be described as utilizing the nonlinear

response in fluorophore emission to a strong pumping excitation to improve

resolution. This includes the likes of STED (STimulated Emission Depletion)

microscopy. Comparatively, stochastic pointillist super-resolution imaging takes a

different approach. To achieve super-resolution in a stochastic single-molecule

pointillist-fashion, both super-localization and active photo-control over fluorescent

emission are required. In recent years, these two ideas have been married in many

stochastic super-resolution studies to enable super-resolution fluorescence imaging,

effectively circumventing diffraction. We now briefly describe both super-localization

and active control in the following. Please reference Chapter 1 for more in-depth

discussion on super-resolution (SR) imaging.

142

The idea and process of super-localization of a single emitter in an image is

based on our knowledge of the shape of the point spread function (PSF) of the

microscope. The fluorescent signal emitted by a single fluorescent molecule appears in

the image relayed by a microscope as a broad, diffraction-limited point spread

function. (Technically the single molecule emitter is not a point source of light, but

rather a dipole emitter, but that subtle distinction is out of the scope of this discussion.)

Even though size of the point spread function in the sample plane may be large

(diffraction-limited to ~200 nm) compared to the molecule emitting the light (1-2 nm),

it is possible to localize the molecule to nanometer precision by numerically fitting the

observed PSF with a model function, such as a 2-D Gaussian or an Airy function.

This seminal idea was first explored and used in the first recordings of single-molecule

images, in which the fluorescence excitation signal from one molecule was used to

map out the size of the focused pumping laser beam spot.2 This concept of fitting the

measured PSF to achieve ‘‘super-localization,’’ or spatial localization beyond the

diffraction limit, is well-known in many areas of science and in one early biological

experiment was applied to the super-localization of single nanoscale fluorescent beads

with many emitter molecules.3 The accuracy with which a single molecule can be

localized depends fundamentally upon the signal-to-noise ratio of the single-molecule

image. Since this image is generated by the Poisson process of photon detection, the

most important factor affecting accuracy is the total number of photons detected above

background, with a weaker dependence on the size of the detector pixels and

background noise.4;

5, 6

Active control is the second concept necessary to achieve super-resolution

imaging. To use a superposition of super-localizations of single molecules to construct

a super-resolution image of a structure, it is necessary to obtain positional information

for at least two emitters within each resolution unit, according to the Nyquist–Shannon

theorem.7, 8

However, in theory this is impossible to accomplish, as super-localization

can only be achieved if adjacent molecules are clearly distinguishable (further apart

than the diffraction limit) so that they can be fitted individually. Therefore, a strategy

needed to be devised to handle high concentrations of fluorophores, where PSFs

143

would overlap, and still be able to achieve super-localization. Active control of single

emitters presents a solution. In the year 2006, three separate research groups

independently applied direct photo-control of single emitters to achieve super-

resolution imaging in biologically relevant systems. Hess et al. and Betzig et al. both

used photoactivated GFP fusions for fluorescence photoactivation localization

microscopy (FPALM),9 and photoactivated localization microscopy (PALM)

10,

respectively. In the Zhuang group, Rust et al. made use of photo-switching produced

by Cy5 emitters in close proximity to Cy3 activator molecules in the presence of thiols

for a method termed stochastic optical reconstruction microscopy (STORM).11

Since

2006, other photo-control schemes have been developed, including ones based on (a)

photoreactivaiton of fluorescent proteins,12

(b) dark state shelving (ground state

depletion microscopy followed by individual molecule return to ground singlet state

[GSDIM], 13

blink microscopy, 14

and ROXS15

and (c) collisional flux of environment-

dependent fluorophores to membranes from solution (points accumulation for imaging

in nanoscale topography [PAINT]).16

No matter what mechanism is selected for the

photocontrol, all these acronyms use the same basic underlying idea, shown in Figure

4-1. To have a mechanism-independent term for this kind pointillist approach to

super-resolution imaging, the Moerner lab has been using the umbrella term “Single-

Molecule Active Control Microscopy,” or SMACM.17

The success of these

experiments, based on the sequential imaging of sparse subsets of photo-

activatable/photo-switchable single-molecule fluorophores, has enabled optical

imaging beyond the diffraction limit (DL), providing unprecedented visual details into

the sub-diffraction world.9, 11, 13, 16, 18, 19

Finally, it is essential to note that since these

methods are time-sequential, the underlying structure should not change during the

time required to obtain the multiple sparse images. For this reason, many studies use

fixed cells; if live cells are used, the structure should be quasi-static (reference Chapter

1 for more in-depth discussion on super-resolution).

144

Figure 4-1 Schematic showing the key ideas of super-resolution imaging of a

structure by photo-control. (A) It is not possible to resolve the underlying structure in

a conventional wide-field fluorescence image because the fluorescent labels are in

high concentration and the images overlap due to diffraction. (B) Using controllable

fluorophores, it is possible to turn on and image a sparse subset of molecules which

then can be localized with nanometer precision (blue line is the underlying structure

being sampled). Once the first subset of molecules photo-bleaches, another subset is

turned on and localized. This process is repeated and the resulting localizations

combined to give a reconstructed super-resolution image of the underlying structure.20

Figure courtesy of Michael A. Thompson.

The successes of these stochastic single-molecule super-resolution techniques

have charged the chemistry and biology communities with the challenge and impetus

for the development of new actively controllable fluorophores. What this means it that

in addition to being able to satisfy the specific needs and requirements for single-

molecule fluorescence imaging in live cells, the new fluorophores need to be able to

be switched on and off reliably. Both fluorescent proteins and small-molecule

fluorophores have been explored to fulfill this need.

Aside from STORM and GSDIM, which imaged fixed immune-stained cells,

other super-resolution experiments in living cells have used photo-controllable

fluorescent proteins.21-25

To further assemble a toolkit of controllable fluorescent

protein emitters, several groups have been using complex mutational strategies to

generate improved photo-switchable fluorescent proteins.25-27

While these fluorescent

proteins enjoy the advantage of genetic target specificity, fluorescent proteins on

average provide 10-fold fewer photons before irreversible photo-bleaching than good

organic fluorophores.20, 28

Additionally, small organic fluorophores have the benefit of

synthetic design flexibility for developing specific target specificity, spectral

wavelength, solubility, and other desirable properties for live cell imaging. Therefore,

145

photo-controllable targeted bright organic fluorophores that are compatible with the

live-cell environment would be advantageous.

Finally, as added benefits beyond super-resolution imaging, the ability to

photo-chemically control the fraction of emitting molecules has additional applications

in pulse-chase experiments, single-molecule tracking, or in any situation where the

number of emitting molecules at a given time must be kept low. In the following, I

present the development of a class of targeted small-molecule photoactivatable

fluorophores, the HaloTag azido-DCDHFs for super-resolution imaging in cellular

systems. Section 4.1.2 details the initial design of photoactivatable azido-DCDHFs,

and Section 4.1.3. discusses our strategy to install target-specificity.

Small-molecule photoactivatable fluorophores: the azido-DCDHFs 4.1.2

In 2003, K.A. Willets and O. Ostroverkhova in the Moerner lab discovered29

that the dicyanomethylenedihydrofuran (DCDHF) class of chromophores, originally

designed for photorefractive polymers,30

have very high fluorescence quantum yields

in glassy polymer films and emit millions of photons before photo-bleaching. Since

then, a productive collaboration between the Moerner laboratory and the Twieg

laboratory at Kent State University has developed and explored the properties of this

class of charge neutral fluorescent molecules. The DCDHF molecules are electron

density push–pull fluorophores characterized by the motif of an amine electron donor

covalently linked to an electron acceptor dicyanomethylenedihydrofuran group via a

π-conjugated central extension core. The basic structure of this class of molecules is

shown in Figure 4-2. This work has seen success in a number of areas, including the

direct use of several DCDHF molecules in a number of cellular environments, both

eukaryotic and prokaryotic.31

146

Figure 4-2 Schematic structure of the DCDHF fluorophores. The amine donor and

DCDHF acceptor are connected by a π-conjugated linker. The R groups can be

modified (usually without affecting the photophysics) in order to add reactive

functionality or increased solubility. The naming scheme used in this Dissertation

specifies the π system: “DCDHF-(π unit closest to acceptor)-…-(π unit closest to

donor)” with the π units denoted P = phenylene, V = vinyl, T = thiophene, N =

naphthalene, A = anthracene; the amine donor is not specified because it is present in

all structures.

Recently, the Moerner-Twieg joint venture developed a novel class of

photoactivatable single-molecule fluorophores. Photo-activatable (or “photo-caged”)

donor–π–acceptor push–pull chromophores can be designed by disrupting the original

charge-transfer band,32, 33

and significantly blue-shifting the absorption to the extent

that it is no longer resonant with the red imaging laser. In these cases, the restoration

of fluorescence or “photoactivation” requires a photoreaction that converts the

disrupted charge transfer band to a substituent that is capable of restoring the spectrum

of the charge-transfer band. For example, if the electron-donating half of DCDHF is

removed from DCDHF to photocage the parent fluorophore and produce a fluorogenic

molecule, photoactivation must restore electron donation, red-shifting the absorption

back.

In the approach pioneered by former Moerner Lab graduate student Samuel J.

Lord, the electron-donating amine group in DCDHF is replaced by a weakly electron

withdrawing azido group, disrupting the push-pull motif, and significantly blue-

shifting the charge transfer band of the fluorophore. 34-37

When excited with long

wavelengths that are resonant with the parent fluorophore, the modified azido

fluorogenic molecules are dark. Fortunately, aryl azides have been extensively studied

147

and are known to be very photolabile.38

When fluorescence from the fluorogenic

molecule is desired, fluorescence can be restored by applying low-intensity blue light

to photochemically convert the azide to an amine via an intermediate nitrene. This

restores the donor–acceptor motif, red-shifts fluorophore absorption, and regenerates

the bright fluorescent emission. While this photochemical reaction by itself by may

also produce azepine products, which would be insufficient to restore the push-pull

motif, Soundararajan and Platz39

demonstrated that directly installing electron-

withdrawing substituents near the intermediate can stabilize the nitrene intermediate in

the photochemical reaction and thereby promote the formation of the desired amine. In

sum, photoactivatable molecules designed with the aforementioned strategies can be

used in pointillist super-resolution imaging.

Figure 4-3 Photo-conversion of dark azido fluorogens produce fluorescent emitter. Photo-conversion of dark azide-substituted fluorogens produce fluorescent amine-

substituted fluorophores, which may involve insertion into C–H or C–C bonds. A dark

photoproduct is also possible. Reprinted with permission from Ref.40. Copyright 2011


Furthermore, unlike the photo-switching or blinking processes used in other

super-resolution schemes, photoactivation of these azido fluorogens to produce

fluorophores is an irreversible chemical reaction. Therefore, reversible photoswitching

is not possible. This may be a drawback in cases that require cycling between bright

and dark forms, such as in STED or when using SMACM to image dynamics. On the

other hand, for quantitative imaging of static structures, a probe that activates, emits

millions of photons, and then disappears permanently is preferable. Otherwise, the

reactivation of fluorophores causes some portions of the structure to be repeatedly

localized, thus complicating the counting of labeled molecules and altering subsequent

image analysis and reconstruction. Photoactivation of azido DCDHFs as well as other

148

push–pull chromophores has been previously demonstrated.34-37, 41, 42

Figure 4-3

shows the primary photoactivation reaction of an azido push–pull DCDHF fluorogen.

Detailed syntheses by colleagues in the Twieg laboratory have been reported in

previous papers and in supporting online material.37, 40, 43

Figure 4-4 Chemical structure and spectra of an azido DCDHF-V photo-activatable

flurogen. (A) Photochemical activation of the azido DCDHF-V fluorogen from dark

azido state to bright amino state. (B) Absorption curves in ethanol (bubbled with N2)

showing photoactivation of azido DCDHF-V (λabs = 424 nm) over time to fluorescent

product amino DCDHF-V (λabs = 570 nm). Different colored curves represent 0, 10,

90, 150, 240, 300, 480, and 1320 s of illumination by 3.1 mW/cm2 of diffuse 407-nm

light. Dashed line is the absorbance of pure, synthesized amino DCDHF-V. (Inset)

Dotted line is weak pre-activation fluorescence of azido DCDHF-V excited at 594 nm;

solid line is strong post-activation fluorescence resulting from exciting amino

DCDHF-V at 594 nm, showing >100-fold turn-on ratio. Reprinted with permission

from Ref.40. Copyright 2011 American Chemical Society.

A common property among the azido push–pull fluorogens is their high

sensitivity to photo-activating illumination, as measured by the photo-conversion

quantum yield ΦP. Two prior publications and Sam Lord’s thesis from the Moerner lab

discuss various strategies for characterizing the photo-conversion quantum yield of

photoactivatable fluorophores, in addition to other photophysical figures of merit. We

discuss here in brief.

149

Photo-conversion from a dark precursor fluorogen to the emissive form can be

monitored in bulk ensemble-averaged measurements by measuring changes over time

in absorbance or emission of the reactant or photoproduct of interest. We can define

the quantum yield of photo-conversion ФP is defined as:

hcIσ

RR

R

λλ

λ

11

PabsPabs

PP

Equation 4-1

where RP is the rate of photo-conversion; Rabs is the rate of photon absorption; τP as the

average decay constant from the exponential fit of the decaying absorption values for

the starting material; the absorption cross-section is related to the molar absorption

coefficient by the equation σλ = (1000)2.303ελ/NA ≈ 10–16

cm2; Iλ is the irradiance at

the sample; λ is the excitation wavelength; h is Planck’s constant; and c is the speed of

light.

ФP, photoreaction yield, is the probability that the starting material will react

for each photon absorbed. The higher the value of ФP, the more the more readily-

convertible the fluorogen is to the activating light, such that less potentially cell-

damaging blue or UV irradiation is required to activate fluorescence. Generally

speaking, only a fraction of the precursor fluorogenic molecules become fluorescent

because the photoreaction yield to fluorescent product is usually less than unity.

Table 4-1 Photophysical properties of various photo-switchable molecules. This

table includes information regarding whether the fluorophore can be cycled between

bright and dark states multiple times, absorption and emission peaks, and molar

absorption coefficient, fluorescence quantum yield, photo-conversion quantum yield,

turn-on ratio, photo-bleaching quantum yield, and total photons emitted. All values are

reported for the photo-converted form except photo-conversion quantum yield. The

first three rows are fluorophores we have designed and characterized; the data for the

other rows were extracted from the literature. Table adapted from Ref.20.

λabs/λem

(nm)

εmax

(M–1

cm–1

) ФF ФP

turn-on

ratio[b]

ФB

Nphotons

DCDHF-V-P-

azide37, 40

570/613 54,100

0.025–

0.39[c]

good

(0.0059)

excellent

(325–

1270)[d]

4.1×10–6

2.3×106

Cy3/Cy5+thiol28

, 44-46

647/662[a]

200,000 0.18 very good excellent

(≤1000)[e] ~670,000

150

EYFP26, 28, 47-51

514/527[a]

83,400 0.61 moderate

(1.6×10–6

) moderate

5.5×10–5

~140,000

PAGFP24, 28, 49, 52

504/517 17,400 0.79 moderate

(1.1×10–6

)

moderate

(100)

~6×10–5

~140,000

mEosFP52-54

559/581 37,000 0.62 good

(1.6×10–5

) very good

3.0×10–5

21,000

[a] Reversible photo-switch. Some fluorophores listed as not reversible may be, but have yet to be

reported as such. [b] Ratio of the fluorescence after and before photoactivation (see definition

above). Some papers report a “contrast ratio” of red to green fluorescence, which is the product of

the fold increase in red fluorescence and fold decrease in the green fluorescence;55

therefore, those

reported contrasts are many times higher than the turn-on ratio, which is the relevant parameter for

super-resolution imaging. Other papers report “contrast ratios” without definition, so I cannot

confidently compare these values directly to turn-on ratio. [c] DCDHFs become brighter when

rigidized.56, 57

[d] This range corresponds to Reff–R. [e] In the SI of reference 46

is reported only

0.1% spontaneous turn-on at ideal conditions (e.g. very high thiol and oxygen-scavenger

concentrations). This value does not take into account the inherent on-off ratio of a single Cy5, so

it is an upper limit. [f] Value estimated from photo-conversion wavelengths, intensities, times, and

spectra reported previously.58

Table 4-1 tabulates and compares important photo-physical quantities of some

of the most popular fluorophores for super-resolution imaging, including both

fluorescent proteins and small-molecule fluorophores. The photo-conversion quantum

yield of DCDHF-V-P azide compares well with the other molecules. Another noted

benefit of using azido-DCDHFs is that the photo-conversion quantum yield is tunable.

For example, adding electron-withdrawing fluorine atoms to the azido-phenyl group

dramatically increases the photoactivation quantum yield.37

For some fluorogens,

only tens or a few hundreds of photons need be absorbed before the fluorogen

converts to a fluorescent product. This is particularly important in live cell imaging

because high doses of blue or UV light can kill cells or alter cellular morphology and

physiology. This benefit comes with the requirement that sample preparation be

carried out in the dark or under red light to avoid undesired preactivation of the

fluorogenic molecules.

Figure 4-4 and Figure 4-5 demonstrate the utility of the azido DCDHF as a

fluorogen in living systems: before photoactivation, live Chinese Hamster Ovary cells

incubated with DCDHF-V-P-azide are dark; after photoactivation, the cells light up as

DCDHF-V-P-azide converts to DCDHF-V-P-amine. It is important to note that the

intensity of activating light required to generate fluorescent DCDHF molecules is

extremely low, at least three orders of magnitude lower than the intensity of the

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imaging light. This helps ensure that the high-energy blue or UV activating light does

not kill the cells of interest or alter their morphology. Furthermore, photo-bleaching is

three orders of magnitude less likely than photo-conversion, so the activated

fluorophore emits millions of photons before photo-bleaching (see Table 4-1). Figure

4-4 shows three Chinese hamster ovary (CHO) cells growing on a glass slide and

incubated with azido DCDHF-V, which easily inserts into and penetrates the plasma

membrane; fluorescence in the cytosol turns on only after a short flash of low

intensity, diffuse violet light. A fraction of the fluorophores remained stationary at the

activation site, possibly bio-conjugated to relatively static biomolecules (via nitrene

insertion into C−C bonds, as in the case for photo-affinity labeling). The remaining

untethered fraction was free to move throughout the cell. It is also important to note

that single molecules were clearly visible while diffusing in the cell.

Figure 4-5 Demonstration of the photoactivation of azido DCDHF-V in live cells.

A. Three CHO cells incubated with fluorogen azido DCDHF-V are dark before

activation. B. The fluorophore amino DCDHF-V lights up in the cells after activation

with a 10-s flash of diffuse, low-irradiance (0.4 W/cm2) 407-nm light. (False color: red

is the white-light transmission image and green shows the fluorescence images,

excited at 594 nm.) Scalebar: 20 μm. C Single molecules of activated amino DCDHF-

V in a cell under higher magnification. Background was subtracted and the image was

smoothed with a 1-pixel Gaussian. Scalebar: 800 nm. Reprinted with permission from


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Figure 4-6 Single molecule trajectories of DCDHF-V-P-amine fluorophore

diffusing in the membrane of a CHO. (left) The trajectory of a single copy of the

DCDHF-V-P-amine fluorophore diffusing in the membrane of a CHO cell after

photoactivation. Dotted red lines indicate when the fluorophore was dark (i.e.

blinking). (right) A background-subtracted intensity time-trace of the molecule in the

trajectory on the left. Red lines indicate when the fluorophore was dark (i.e., initially

blinking events, then finally bleaching). Single-particle tracking was performed using

ImageJ and the “SpotTracker” plugin,59

with the following parameters: maximum

displacement of 5 pixels, intensity factor of 80%, intensity variation of 0%, movement

constraint of 20%, and center constraint of 0%. Reprinted with permission from


In the following section, I discuss our strategy to install target-specificity to the

azido DCDHF-V-P to enable cellular imaging.

Cellular labeling considerations 4.1.3

Since Coons et al.’s seminal publication in 1941,60

the field of biology has

relied heavily on immunofluorescence for target-specific microscopy. The basis of

immunofluorescence, immunohistochemistry (IHC), is very simple and bridges three

scientific disciplines: immunology, histology, and chemistry. The fundamental

concept behind IHC is the demonstration of the presence of antigens (Ag) within

biological samples by means of specific antibodies (Abs). Once antigen–antibody (Ag-

Ab) binding occurs, a fluorescently labeled secondary antibody is used to recognize

the first antibody (usually termed primary antibody). However, to access cytosolic

targets within the cellular interior, fixation and subsequent membrane

permeabilization of the cell sample are required. The fixation step is accomplished by

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cross-linking or dehydration of cellular proteins by organic solvents (e.g.

paraformaldehyde, methanol), and the permeabilization is achieved by treatment of the

cell with chemical surfactants (e.g. TritonX, Tween). Please see Chapter 2 for further

details. This permeabilization step makes cellular delivery of large molecules like

antibodies possible.

Unfortunately, many biological processes can be drastically halted and

compromised by the fixing and permeabilizing techniques used in most super-

resolution experiments that rely on antibody-basesd targeting of organic fluorophores,

such as STORM.13, 61

This pressing problem motivates the development of additional

techniques and probes that can be used for super-resolution imaging in living cells.

Single-molecule imaging in living cells using exogenous fluorophores faces the dual

hurdles of cell permeability and targeted labeling. There are a few exceptions in the

literature that did not use fixation and immunostaining. For instance, Heilemann et

al.61

imaged mRNA in living cells using oligomers labeled with small organic

fluorophores and Conley et al.45

labeled the external lysines of bacterial cells using a

Cy3–Cy5 heterodimer. Very recently, fluorophores were targeted with trimethoprim

and intrinsic cellular reductants enabled photo-induced blinking.62

These examples

demonstrated some possibilities for live-cell labeling and super-resolution, but there is

still a need45

for photoactivatable organic labels for super-resolution imaging. As

mentioned in 4.1.1., this is because each molecule is localized only once, while with

blinking or photoswitching, each molecule can be localized a variable number of times

complicating possible subsequent interpretations.

In addition to the number of photons, which determines the localization

accuracy, fluorophore labeling density is another important variable that determines

the ultimate resolution. Because super-resolution imaging by switching point sources

is effectively a sampling of the true underlying structure, there are well-known

requirements on the labeling density (or spatial sampling frequency) for a correct

reproduction of the structure from the samples at a given resolution. As mentioned in

4.1.1., the Nyquist–Shannon sampling theorem7, 8, 50, 54, 63

requires that the

fluorophores label the structure of interest at a frequency (number per spatial distance)

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that is at least double the desired resolution. For an ultimate resolution of ~20 nm in

two dimensions, this requirement means that there must at least one label every 10 nm

on average, translating to hundreds of labels in each diffraction-limited area; for 3D

imaging, this requirement increases to thousands or tens of thousands of labels per

diffraction-limited volume. To satisfy this criterion, a high availability of labeling sites

and labeling efficiency are necessary. In other words, we need a labeling strategy in

which we can have control over the number of labeling sites, and can achieve high

enough labeling density without causing deleterious effects on the cellular

morphology.

Our cellular labeling strategy: HaloTag/HaloEnzyme Enzymatic 4.1.4

Targeting

The expression of a protein-of-interest (POI) as a fusion protein with an

additional polypeptide (tag) that aids in the characterization of the protein was first

exploited by Shuman et al. in 1980; in this experiment, β-galactosidase was fused to

the cytoplasmic membrane protein MalF to facilitate its purification. 64

Since then,

site-specific labeling of proteins-of-interests via a peptide Tag has received much

development. (Also see Chapter 1 for more background). 65-72

Here, we present a

target-specific photoactivatable organic fluorophore for use inside living and fixed

cells based on the commercial HaloTag targeting approach.70

The HaloTag targeting scheme is based on the efficient formation of a covalent

bond between a specific ligand, the HaloTag substrate (HaloTag), and a reporter

protein, HaloEnzyme (HaloEnz). Therefore, in installing target-specificity to azido

DCDHF, our method requires 2 components: 1) a fluorophore that has been furnished

with a HaloTag; 2) a genetic fusion of the POI to the HaloEnz. The HaloEnZ forms a

covalent linkage to the HaloTag, thus labeling the POI with the azido fluorophore (i.e.

a protein–HaloEnz–HaloTag–fluorophore covalent unit). The HaloEnZ is a

monomeric protein (MW 33 KDa) that cleaves carbon halogen bonds in aliphatic

halogenated compounds. Upon nucleophilic attack by the chloroalkane to Asp 106 in

the enzyme, an ester bond is formed between the HaloTag ligand and the HaloEnZ

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(Figure 4-7). The HaloEnZ protein contains a critical mutation in the catalytic triad

(His 272 to Phe) so that the ester bond formed between HaloEnZ and HaloTag ligand

cannot be further hydrolyzed. See Figure 4-7 for an illustration of the conjugation

mechanism.

Figure 4-7 Schematic of the specific conjugation mediated by the HaloEnz protein

and its HaloTag ligand. The terminology used here differs from the commercial

terminology to provide a clearer explanation.

Photoactivatable azido DCDHF, specifically azido DCDHF-V-P, as described

in 4.1.2 was furnished with a HaloTag. Figure 4-8, molecule 3, is the azido DCDHF

HaloTag used in our experiments.

Figure 4-8 Photochemical activation of azido DCDHF. Photochemical activation

produces 2 from 1. A mixture of photoproducts is produced,14,15

but the primary amine

with R1=R2=H is the significant product (see Table 4-1 for reaction yield of primary

amine). HaloTag versions of 1 and a separate non-photoactivatable fluorophore are

also shown (3 and 4). Reprinted with permission from Ref.75. Copyright 2011


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To enable covalent linkage to this HaloTag-functionalized fluorophore, the

POI must be genetically fused to the protein HaloEnZ at either its N- or C- terminus.

The resulting fusion protein can then be conjugated to the fluorophore through the

reaction between HaloTag ligands and HaloEnZ (Figure 4-7). Figure 4-9A illustrates

the flow of necessary steps for this experiment. First, a plasmid that encodes a

HaloEnZ-POI fusion must first be constructed; then, the cellular samples must be

transfected with this plasmid to express the HaloEnZ-POI fusion. The fluorogen must

then be washed into the cell, to enable the HaloEnZ-HaloTag reaction. Finally, the

unbound fluorogen must be washed out before photoactivation. Please see 4.2.3. for

details of the experiment.

Figure 4-9 (A) Flow chart of experimental procedures in using HaloTag-DCDHF-

fluorogen. (B) HaloTag-DCDHF fluorogen forms covalent bond with HaloEnzyme.

In this scheme, a plasmid that encodes a HaloEnZ-POI fusion must first be

constructed. Then, the cellular samples must be transfected with this plasmid to

express the HaloEnZ-POI fusion. The fluorogen must then be washed into the cell, to

enable the HaloEnZ-HaloTag reaction (B). Finally, the unbound fluorogen must be

washed out before photoactivation.

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In sum, we present the first target-specific photoactivatable organic

fluorophore for use inside both living and fixed cells, azido DCHDF-V HaloTag,

based on the commercial HaloTag targeting approach. 70

This method requires a

genetic fusion to the HaloEnzyme (HaloEnz), which forms a covalent linkage to the

HaloTag substrate, thus labeling the protein-of-interest (i.e. a protein–HaloEnz–

HaloTag–fluorophore covalent unit). Specifically, after describing experimental

details in Section 4.2, we present in the following section: (i) the basic photophysical

properties of a new targeted photoactivatable probe; (ii) proof-of-principle labeling of

known structures in fixed and living mammalian cells validated by co-staining with

antibodies or co-transfection with fluorescent proteins; (iii) specific super-resolution

imaging of microtubules in a mammalian cell with quantification of resolution

enhancement; (iv) demonstration of targeted labeling in living bacteria with

diffraction-limited imaging; and finally, (v) super-resolution imaging of poorly

understood structures inside living bacteria.

Experimental 4.2

In the following, specific experimental details required to illustrate the

specificity of the HaloTag azido DCDHF-V-P and its utility in super-resolution

imaging in both bacterial and mammalian cells are detailed.

Plasmids for HaloEnz Fusion Construction 4.2.1

We constructed several HaloEnZ fusion proteins to illustrate the facility of our

approach: HaloEnz–-tubulin for mammalian expression, FtsZ-HaloEnz, AmiC-

HaloEnz, and HaloEnz-PopZ for bacterial expression. The construction details for the

necessary plasmids are detailed in the following.

4.2.1.1 Construction of HaloEnz–-tubulin Plasmid for Mammalian Cell

Expression

The gene encoding -tubulin (Clontech) was amplified by PCR using primers

that incorporate NheI and BamHI restriction sites at the 5’ and 3’ ends, respectively,

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digested with NheI and BamHI, and ligated into HaloEnz pHT2 (Promega) to make

the plasmid HaloEnz–-tubulin that enables C-terminal HaloTagging of human -

tubulin for mammalian cell expression.

Construction of FtsZ-HaloEnz and AmiC-HaloEnz Plasmid for 4.2.1.2

Caulobacter crescentus

The gene encoding the HaloTag enzyme (Promega) was amplified by PCR

using primers that incorporate EcoRI and NheI restriction sites at the 5’ and 3’ ends,

respectively, digested with EcoRI and NheI, and ligated into pXYFPC-2 73

that was

similarly digested to remove the gene encoding EYFP and make the plasmid

pXHALOC-2. pXHALOC-2 is an integration vector that enables C-terminal Halo-

tagging of a gene of interest. The gene encoding Caulobacter ftsZ was amplified by

PCR using primers that incorporate NdeI and EcoRI restriction sites at the 5’ and 3’

ends, respectively, digested with NdeI and EcoRI, and ligated into pXHALOC-2 that

was similarly digested to make plasmid pEG515. The gene encoding Caulobacter

amiC was amplified by PCR using primers that incorporate KpnI and SacI restriction

sites at the 5’ and 3’ ends, respectively, digested with KpnI and SacI, and ligated into

pXHALOC-2 that was similarly digested to make plasmid pEG254. pEG515 and

pEG254 were introduced into Caulobacter strain CB15N by electroporation and

integrated at the xylX locus to make strains EG603 and EG152 for xylose-inducible

expression of ftsZ-HaloEnz and amiC-HaloEnz, respectively.

Construction of HaloEnz-PopZ Plasmid for Caulobacter crescentus 4.2.1.3

To make the Caulobacter strain for producing HaloEnz-PopZ, the HaloTag

enzyme (Promega) was inserted as an N-terminal tag between the popZ promoter and

coding sequence. The PpopZ-haloEnz-popZ sequence was cloned into the

pNPTS138 vector (MR Alley, unpublished) and subsequently integrated into

the popZ locus in the CB15N Caulobacter genome. This resulted in a merodiploid

strain in which halo-popZ and untagged popZ are expressed from tandem

PpopZ promoters.

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To construct the PpopZ-haloEnZ- popZ sequence, the popZ promoter region

and halo tag were amplified using primer sets "1319 left flank 5'KpnINheI" / "1319

HALO at N olp Left NdeI" and "1319 HALO at N olp Right NdeI" / "3' HALO EcoRI

nostop", respectively. The PCR products were stitched together by overlap PCR, then

cloned into pNPTS138 using NheI and EcoRI restriction sites. Subsequently, the

popZ coding sequence was amplified using the primer set "1319 5' EcoRI" / "1319 3'

EcoRV", then cloned into the vector using EcoRI and EcoRV restriction sites.

Primer sequences:

1319 leftflank 5'KpnINheI: aaaaggtaccgctagcGACGGTCTCGGCGCGCGCTT

1319 HALO at N olp Left NdeI:

TGGCTCGAGcataTGCGGGGCCGTCGTAAAGAG

1319 HALO at N olp Right NdeI:

ACGGCCCCGCAtatgCTCGAGCCAACCACTGAGGA

3' HALO EcoRI nostop: aaaagaattcaccATGTCCGATCAGTCTCAAGA

1319 5' EcoRI: aaaagaattcaccATGTCCGATCAGTCTCAAGA

1319 3' EcoRV: ttttgatatcGGCGCCGCGTCCCCGAGAGA

Cell Labeling Protocol 4.2.2

We labeled several HaloEnZ fusion proteins to illustrate the efficacy of our

approach in cells: HaloEnz–-tubulin in fixed and live mammalian cells, FtsZ-

HaloEnz, AmiC-HaloEnz, and HaloEnz-PopZ in live bacterial Caulobacter

crescentus. The construction details for the necessary plasmids are detailed in the

following.

HeLa Cell Labeling 4.2.2.1

HaloEnz–-tubulin DNA was transfected into HeLa cells along with HaloTag

pHT2 as the control. Transfected cells were split at 1:8 ratio 16 hours later and plated

on ploy L-lysine coated 12 mm glass cover slips. Cellular morphology was not

perturbed by the transfection. Cells were then treated with 1 M, 5 M and 10 M

Azido DCDHF-V-P fluorophore 3 respectively, at 37 °C for 30 min, followed by 3

washings with DMEM medium at 37 °C (10 min for each wash) and subsequent

fixation with 4% paraformaldehyde. Fixed cells went through 0.1% Triton X-100

permeabilization in PBS with 5% Bovine Serum Albumin. Monoclonal anti--tubulin

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antibody was applied onto permeabilized fixed cells at 1: 2,000 dilution and

Alexa488-conjugated goat-anti-mouse secondary antibody was added to label

expressed human -tubulin. Cells were stained with 4', 6-diamidino-2-phenylindole

(DAPI) to localize nuclei before mounting and imaging.

CHO Cell Labeling 4.2.2.2

1000ng DNA/1mL HaloEnz–-tubulin DNA was transfected into CHO cells

along with -tubulin–eGFP as the control using Lipofectamine 2000 and Opti-MEM l.

Transfected cells were split at 1:9 ratio 24 hours later and plated on poly L-lysine

coated #1 borosilicate glass cover slips. Cellular morphology was not perturbed by the

transfection. Cells were then treated with 1 M DCDHF-V-P fluorophore 4 at 37 °C

for 30 min, followed by 3 washings by DMEM medium at 37 °C (10 min for each

wash).

BS-C-1 Cell Labeling 4.2.2.3

HaloEnz–-tubulin DNA was transfected into BS-C-1 cells on a 24 well plate

at 1000ng DNA/1mL using Lipofectamine 2000 and Opti-MEM. Cellular morphology

was not perturbed by the transfection. Transfected cells were split at 1:12 ratio 12

hours later and plated on chambered #1 borosilicate glass cover slips. 24 Hours later,

cells were fixed with 4% PFA, followed by 3 times PBS wash, permeabilization with

0.15% Triton X-100, and pre-block with 1mg/mL BSA. Cells were then treated with

12 nM HaloTag DCDHF-V-P fluorophore 3, at room temperature for 90 min,

followed by 3 washings with 0.15% Triton X-100 containing PBS.

Caulobacter Crescentus Cell Labeling 4.2.2.4

The protocol for labeling Caulobacter crecentus cells expressing HaloEnz

fusions with 3 or 4 is as follows: Cells were first grown in PYE growth medium

overnight and then diluted to M2G minimal-fluorescence buffer, to which was added

0.03% xylose to induce expression of the HaloEnz fusions of FtsZ and AmiC.

HaloEnz-PopZ is not on an inducible promoter and as such, no xylose is needed for

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expression of the fusion protein. After inducing for 3 h and when the cell suspension

had reached OD 0.3, the cells were centrifuged at 8 krpm for 90 s to pellet and rinsed

with clean M2 buffer. To label with the fluorescent probe, HaloTag DCDHF 3 in

DMSO was added very slowly to a final concentration of 1 nM of DCDHF and 8%

DMSO v/v. After 90 min incubation, cells were pelleted and re-suspended with clean

M2 to wash 3 times, followed by incubation for 30 min in clean M2. Finally, the cells

were then pelleted and re-suspended 6 more times in M2 buffer for final wash. Cells

were then placed on a 1.5% agarose in M2 pad and imaged on an inverted Olympus

IX71 microscope as described below. All steps involving 3 were performed in a dark

room under dim red lights.

Cell Imaging Protocol 4.2.3

We imaged fixed and live mammalian cells, and live bacterial Caulobacter

crescentus. Experimental details can be found in the following.

Diffraction-Limited Imaging of HeLa Cells 4.2.3.1

Phase contrast images of HeLa cell shape and location were recorded with a

Cool Snap HQ charge-coupled device (CCD) camera (Roper Scientific), with focused

illumination from a 12 volt Halogen lamp (Zeiss). Fluorescence images were acquired

with a Zeiss Axiovert 200M microscope under the following conditions: Zeiss 20×

objective, excitation under a Xenon Lamp, green, red and DAPI fluorescence filters.

Samples were photo-activated by 2-hours Halogen lamp illumination or 1 min UV

illumination at 365 nm.

Diffraction-Limited Imaging of CHO Cells 4.2.3.2

The fluorescence imaging of the cells was performed with wide-field epi-

illumination using an inverted microscope (Olympus IX71). Laser illumination at 594

nm provided an intensity of ~1 kW/cm2 at the sample plane. The epifluorescence was

collected with a 100× magnification, 1.4 NA, oil-immersion objective (PlanApo,

Nikon) and an infinity-corrected objective adaptor. A 594 nm dichroic beamsplitter

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and 610 nm long-pass filter were used to filter the emission. The emission was

collected using a 512×512 pixel Andor Ixon EMCCD at the frame rate of

100ms/frame.

Diffraction-Limited and Super-resolution Imaging of Caulobacter 4.2.3.3

crescentus Cells

For bulk DL imaging, we labeled the HaloEnz fusion proteins in living

Caulobacter crescentus cells by incubating with 4 as described above. In the cells

expressing HaloEnz-FtsZ, approximately 25% of the cells exhibited the expected

protein localization; the remaining were either unhealthy, or were in a part of the cell

cycle in which targeted protein does not assemble at the division plane.74

In the cells expressing HaloEnz-PopZ, about 50% exhibited the expected

protein localization; the remaining 50% exhibited no labeling, probe aggregates within

the cells, or unhealthy morphology. Cells that were unlabeled may have been new

daughter cells that reproduced during the washing steps of the labeling. A certain

degree of aggregation is to be expected as a result of the imperfect solubility of the

probe in buffer. Unhealthy looking cells might have been the result of overexpression

of the HaloEnzyme, disrupting the essential functions of the targeted proteins. The

fluorescence imaging of the cells was performed with wide-field epi-illumination

using an inverted microscope (Olympus IX71). Laser illumination at 594 nm provided

an intensity of ~1 kW/cm2 at the sample plane. The epifluorescence was collected with

a 100x magnification, 1.4 NA, oil-immersion objective (PlanApo, Nikon) and an

infinity-corrected objective adaptor. A 594 nm dichroic beamsplitter and 610 nm long-

pass filter were used to filter the emission. The emission was collected using a

512×512 pixel Andor Ixon EMCCD at the frame rate of 100ms/frame.

Super-resolution Imaging of BS-C-1 Cells 4.2.3.4

Samples were prepared as described in 4.2.3.1. Fluorescence imaging of the

cells was performed with wide-field epi-illumination using an inverted microscope

(Olympus IX71). Laser illumination at 594 nm provided an intensity of ~1 kW/cm2 at

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the sample plane. The epifluorescence was collected with a 100× magnification, 1.4

NA, oil-immersion objective (PlanApo, Nikon). A 594 nm dichroic beamsplitter and

610 nm long-pass filter were used to filter the emission. The emission was collected

using an 512×512 pixel Andor Ixon EMCCD at the frame rate of 150ms/frame.

Super-resolution Image Processing 4.2.3.5

From raw image stacks, super-resolution (SR) images were obtained using

previously published image processing techniques as previously described 50, 51

.

Briefly, for each imaging frame, the position of the a single emitter was determined by

fitting the signal above background in a small region of interest containing the single-

molecule spot to a 2-D Gaussian with nonlinear least squares regression analysis

(nlinfit, in MATLAB). Each single-molecule point spread function was fit to

determine the following parameters: background, amplitude, width, center(x) and

center(y). Finally, each single-molecule position was re-plotted using a macro written

in the image processing software program ImageJ as a 2-D Gaussian profile defined

by the measured integrated intensity and a width given by the average statistical error

in localization of the center (95% confidence interval, averaged over all single-

molecule localizations). The resulting mean localization precision was 32 ± 12nm.

Photoactivation Quantum Yield Characterization 4.2.4

Bulk solution absorption and emission spectra of the 2 were acquired on a Cary

6000i UV–vis spectrometer and a Horiba Fluorolog-3 fluorimeter using standard 1-cm

path length, quartz cuvettes. Molar absorption coefficients were measured from

dilutions of solutions with known concentrations.

The overall chemical reaction yields to fluorescent product listed in Table 4-2

were measured from the absorbance values in the photoactivation spectra. Yield was

defined by [amino DCDHF] f / [azido DCDHF] i = (Aamino/εamino)f/(Aazido/εazido)i, where

A is the absorption, ε is the molar absorptivity, i and f refer to initial and final values,

respectively.

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Photo-conversion of compound 1 in ethanol was performed using a 385-nm

diode flashlight (1.1 mW cm–2

). Photo-conversion was measured by monitoring

changes over time in absorbance values of the reactant and photoproduct of interest in

ethanol (such as the absorption spectra in Figure 4-4). The photo-conversion quantum

yield ФP is defined in equation 4.1. Note that ФP is the probability that the starting

material will photo-convert for each photon absorbed; a fraction of those photo-

converted molecules become fluorescent, because the photoreaction chemical yield is

less than unity (see Table 4-2 for overall chemical yield). The higher the value of ФP,

the more the sensitive the fluorogen is to the activating light, so less potentially cell-

damaging blue or UV irradiation is required to activate fluorescence. For further

details and discussion, see Reference.43

Results 4.3

Photophysical characterization of Azido DCDHF HaloTag 4.3.1

As mentioned, compared with the original azido DCDHF-V,40

azido DCDHF-

V HaloTag exhibits similar spectral changes upon optical pumping of the aryl azide,

but a higher photo-conversion quantum yield (ΦP, see Table 4-2).

Table 4-2 Photophysical/photochemical parameters of Azido DCDHF-V and

derivatives

λabs,azide (nm)a

λabs,amine/λfl,amine (nm)b

Yieldc

ΦPd

Azido-DCDHF-V

424

570/613

65%

0.0059

Azido-DCDHF-V-NHS

(Azido DCDHF-V HaloTag) 443 572/627 ~50% 0.095

a Peak absorbance for azido fluorogen.

b Absorbance and fluorescence peak

wavelengths of the amino fluorophore. c Overall chemical reaction yield to the

primary fluorescent amine. d Photo-conversion quantum yield of azido fluorogens to

any product (i.e., the probability of photo-converting after absorbing one photon).

The oxygen on the aryl azide stabilizes the intermediate nitrene, making photo-

conversion more favorable.38, 43, 57

, 38, 40, 43

The photo-conversion of 3 was so sensitive

such that an additional activating blue laser was not necessary; instead, the diffuse

ambient light (e.g. the blue light emitted from a nearby computer monitor in an

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otherwise dark room) was sufficient to activate sparse sets of the fluorogen. The

thermal activation rate and activation by the 594 nm imaging laser were significantly

lower, as measured in complete darkness in a covered sample. This characterization is

elaborated in the following.

Effects of diffuse white light 4.3.1.1

Figure 4-10 Effects of diffuse white light on Azido DCDHF HaloTag. Reprinted

with permission from Ref.75. Copyright 2011 American Chemical Society.

The effects of diffuse white light on 3 were characterized as shown in Figure.

4-10. In dim red lights, a sample of 3 was prepared on a poly-lysine coated

borosilicate cover glass at 1 M concentration in pH7.4 PBS buffer. 500 ms of

illumination and recording were used to sample the fluorescence above background in

the images (dots in the figure). A small amount of fluorescence appeared when the

sample was covered and not exposed to the ambient diffuse white light from the

nearby computer monitor (tan strip) – this degree of activation was presumably due to

the reading light. As the sample was exposed to additional weak diffuse light from a

small handheld single LED flashlight, more molecules were photoactivated to

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fluoresce (blue strip). As the amount of diffuse light was increased, the number of

photoactivated molecules increased, contributing to a higher fluorescence signal (red

strip). Finally, when the sample was covered again (green strip), we saw a decrease in

the fluorescence signal illustrating the attenuation of photo-activation and photo-

bleaching.

Because the fluorogen’s sensitivity to photo-conversion is so high, it was

possible to set the level of the ambient light such that the bleaching rate from 594 nm

pumping was similar to the activation rate. For instance, with room lights off and a

nearby computer monitor on, we maintained a steady-state concentration of isolated

emitters. This reduced the complexity of the experiment by allowing the use of only

one laser.61

Moreover, because no blue or UV laser was necessary for activation,

photo-damage to the imaging sample was greatly reduced. (The drawback to this high

sensitivity is that it increases the difficulty of preventing photoactivation before

imaging. We successfully minimized preactivation by performing all preparations in

complete darkness or under limited exposure to dim red lights only).

Specificity and efficacy in in mammalian culture 4.3.2

We verified the specificity of 3 in mammalian culture. Wild-type HeLa cells

and HeLa cells transfected to express HaloEnz–-tubulin were stained live with 3,

fixed and immunostained with Alexa488-mAb to -tubulin. Fluorescence images after

photoactivation clearly demonstrate that 3 is only retained in the cells that expressed–

HaloEnz--tubulin (Figure 4-11 A–G). Next, live CHO cells were co-transfected with

HaloEnz--tubulin and -tubulin–eGFP, and the labeling by 4 was shown to co-

localize well with the eGFP labeling (Figure 4-11 H–I).

167

Figure 4-11 Evidence that the HaloTag-targeted fluorogen correctly labels specific

proteins and enables SR imaging in mammalian cells. (A) Phase image of fixed WT

HeLa cells. (B) The cells in A imaged in the DCDHF-V-P channel. (C) The cells in A

imaged in the Alexa488 channel. (D) Phase image of fixed HeLa expressing HaloEnz-

-tubulin labeled with 3. (E) The cells in D imaged in the DCDHF-V-P channel. (F)

The cells in D imaged in Alexa488 channel. (G) Overlay of E, F, and additional blue

DAPI channel to show nuclei. (H) Live CHO cells co-transfected to express both

HaloEnz–-tubulin and -tubulin–eGFP labeled with 3 and imaged in DCDHF-V-P

channel. (I) Cells from H imaged in the EGFP channel. (The higher background in H

may be the result of nonspecific binding and imperfect washing of untargeted

fluorophores.) (J) Fixed BS-C-1 cells expressing HaloEnz--tubulin labeled with 3

imaged using conventional diffraction-limited imaging. Indicated microtubule

measures 450±40nm FWHM. (K) Same cell as J with SR imaging. Indicated

microtubule measures 85±15nm FWHM. See 4.2.2 for sample preparation procedures.

Reprinted with permission from Ref.75. Copyright 2011 American Chemical Society.

168

Most importantly, to directly demonstrate the SR imaging enabled by our new

targeted fluorogen, BS-C-1 cells were transfected with HaloEnz--tubulin, fixed,

stained with 3 and washed before SR imaging by PALM. Comparing DL and SR

fluorescence images (Figure 4-11 J, K), the microtubule structure is clearly imaged

with resolution beyond the optical diffraction limit. After corroborating the utility of

the DCDHF fluorophores for labeling known structures in mammalian cells, we then

moved our attention to cells with protein organization that is not fully understood.

Specificity and efficacy in live bacterial samples 4.3.3

Bacteria are tiny (about a thousand bacteria could fit within one HeLa cell),

and the details of protein localization in prokaryotes are poorly understood, yet

essential for function and phenotype.74 DL imaging of labeled proteins in bacteria

only produces diffuse blobs, complicating meaningful interpretation. For these

reasons, living bacterial studies benefit greatly from SR imaging.20, 38 We used

HaloTag-DCDHFs to highlight protein localization patterns in live Caulobacter

crescentus bacteria,50, 51, 76-81 which demonstrates the biologically interesting

ability to divide asymmetrically. Elucidating the mechanisms of asymmetric cell

division and intracellular organization requires understanding how cytoskeletal

proteins localize through the life cycle of the cell.74

We first show that without any expression of HaloEnZ protein fusions,

Caulobacter crescentus bacteria is not labeled by azido DCDHF HaloTag (Figure

4-12). The cells also exhibit normal morphology and divide after labeling indicating

that our labeling procedures have not caused deleterious effects on the cells. To

demonstrate the target-specificity, a polar protein PopZ,77, 78 and mid-plane proteins

FtsZ79 and AmiC were expressed as HaloEnz fusions. PopZ, FtsZ, and AmiC have

distinct roles: PopZ anchors the chromosomal origin at the “swarmer” pole; FtsZ and

AmiC are recruited to the mid-plane and are components of the cell division

machinery.82 DL imaging using HaloTag targeting of the (non-photoactivatable)

fluorophore 4 shows correct PopZ localization at cell poles and FtsZ at the cellular

169

division plane as expected (Figure 4-13), again confirming that this HaloTag labeling

system does not significantly interfere in phenotype.

Figure 4-12 Negative bacterial controls. In this image, the white light transmission

image of the cells is in the black channel and the fluorescence from activated DCDHF

fluorophores is in the red channel. Caulobacter crescentus expressing HaloEnz-PopZ

incubated with 1 (no HaloTag functionality) shows no labeling inside the cells.

Additional control experiments in which cells not expressing any HaloEnz fusions

were incubated with HaloTag Azido DCDHF-V-P 3 also showed no labeling inside

the cells.

Figure 4-13 Diffraction-limited imaging of DCDHF HaloTag inside live C.

crescentus cells expressing fusion proteins to FtsZ and PopZ. These proteins localize

as expected,25

indicating that the HaloTag–DCDHF labeling does not disrupt typical

cellular behavior. Reprinted with permission from Ref.75. Copyright 2011 American

Chemical Society.

170

SR images produced by photoactivation of fluorogen 3 display the expected

localization patterns, but also reveal additional detail unseen in the DL images of

Figure 4-13. For PopZ at the cell pole, the protein forms an asymmetric cap-like

structure with a curvature that hugs the shape of the bacterial membrane (Figure 4-14

A–C). Also, in the case of AmiC, the protein localizes to the cellular mid-plane, as

expected (Figure 4-14 D–E). The SR images of AmiC may reveal a tighter

organization than seen in DL microscopy, but further study is necessary before any

definitive statement can be made. In either case, these SR images provide new detail

not available in DL images.

Figure 4-14 SR imaging of protein fusions inside live C. crescentus cells using

azido-DCDHF HaloTag. (A–C) PopZ forms a polymeric network at the poles of the

cells. Compared to the DL images in Figure 3, these SR images reveal distinct shapes

of the PopZ structure, including the cap-like network in C. (D–E) AmiC is recruited to

the division plane early in the cell cycle. These SR images indicate that AmiC may

form a structure that hugs the membrane. The SR images are extracted from

171

localizations over 75 seconds with a mean localization precision of 32 ± 12 nm.

Reprinted with permission from Ref.75. Copyright 2011 American Chemical Society.

The target-specific DCDHF single-molecule fluorogen presented here

represents the first successful installation of target-specificity to small organic

photoactivatable fluorogens for single-molecule SR imaging. Compared to existing

schemes, the photoactivation of the fluorogen does not require other additives (e.g.

thiols for Cy5,83

or redox chemicals)61

nor activation by UV light, and thus can be

used inside living cells. SR imaging has been directly demonstrated for fixed

mammalian and live bacterial cells; additional effort to improve washout for live

mammalian cells is an important topic for future work. This and future

photoactivatable fluorogens should be helpful tools for SR imaging in the complex

environment within the living cell.

Acknowledgements 4.4

This work was supported in part by Grant No. R01-GM086196 from the

National Institute of General Medical Sciences. We thank Professor S. Pfeiffer for BS-

C-1 cells. We thank Dr. Julie Biteen and Dr. Steven Lee for helpful discussions.

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179

5.FLUORESCENT SAXITOXINS FOR SUPER-

RESOLUTION IMAGING OF VOLTAGE-GATED

SODIUM ION CHANNELS ON LIVE NEURONAL

CELLS

180

Abstract

Neuronal function is shaped by the proper expression, distribution, and

operation of voltage-gated sodium ion channels (NaVs). A desire to better understand

the role of NaVs in signal conduction and the relationship between their disregulation

and specific disease states motivates the development of high precision tools for their

study in dynamic living systems. Nature has evolved a collection of small-molecule

guanidinium-based nerve-blocking agents, the most celebrated of which are (–)-

tetrodotoxin and (+)-saxitoxin, that bind with low nanomolar potency and with

exquisite fidelity to select NaV isoforms. De novo chemical synthesis has enabled the

preparation of two fluorescently labeled forms of (+)-saxitoxin, STX-Cy5 and STX-

DCDHF, which maintain reversible nanomolar affinity for endogeneous NaVs

(NaV1.2, 1.7) expressed in differentiated PC12 cells. Fluorescence colocalization

studies with a pan-NaV antibody confirm that binding of these STX-based dyes is

highly selective to NaV. Single-molecule imaging experiments reveal subcellular

distributions of NaVs in the soma, axons, and dendrites of these cells and offer the first

super-resolution images of NaV locations in filopodia and in neuritic spines beyond the

optical diffraction limit. Collectively, these data establish STX-Cy5 and STX-

DCDHF as selective molecular probes for real-time NaV imaging and offer a dynamic

view of neuronal differentiation with unprecedented visual detail.

This work has been submitted to PNAS,1 and represented a collaborative effort

between the Moerner and DuBois labs. The fluorophores were covalently attached to

the amine-functionalized saxitoxins in the DuBois lab by A. Ondrus following a

reaction first devised by B. Andresen. A. Ondrus and W. Parson verified the potency

of the fluorescent saxitoxins by electrophysiology. Confocal experiments were by

performed by A. Ondrus and H.-L. Lee, from the Moerner Group. Colocalization

experiments and analysis were done by H.-L. Lee. Wide-field imaging experiments,

single-molecule tracking, sub-diffraction analysis and reconstructions were performed

by H.-L. Lee assisted by S. Iwanaga in the Moerner Group.

181

Introduction 5.1

Cell membrane potential and action potential 5.1.1

As the cellular membrane is composed largely of lipid molecules with a

hydrophobic interior, it is intrinsically impermeable to ions such as Na+ and K

+.

Responsible for about 70% of energy consumption of the entire nervous system, the

Na+/K

+ ion pump ATPase transports three Na

+ ions out of the cell while pumping two

K+

ions into the cell per pump cycle, creating Na+

and K+

concentration gradients

across the cell membrane. Given that the plasma membrane is very thin (measuring

approximately 8 nanometer in thickness on average), a very small transmembrane

voltage is all that is needed to generate a large electric field. In this light, the Na+/K

+-

ion pump ATPase is an integral component that regulates the cell membrane potential.

2-4

Stemming directly from the equilibration of electric potential and chemical

potential across a plasma membrane, a properly maintained cellular potential is an

important characteristic of living cells. The function of the cellular membrane

potential can be understood as two-fold. First, a stable membrane potential allows a

cell to provide power to operate a myriad of transmembrane proteins integral to cell

function. Transmembrane potentials exist for all mammalian cells and range in value

from -40 to -90 mV. In non-excitable cells (e.g. fibroblasts), and in excitable cells

(e.g. neurons, myocytes, electrocytes) in their baseline states, the membrane potential

is held at steadfast level, also known as the resting potential. For excitable cells such

as neurons, the typical values of resting potential range from approximately –70 to –80

millivolts across the cell membrane relative to 0 volts outside the cell.3, 5-7

Secondly, in electrically excitable cells such as neurons and cardiomyocytes,

changes in membrane potential are used for transmitting signals between different

parts of a cell and even between cells. More relevant to the following discussion,

opening or closing of specific ion channels at a localized single point in the plasma

membrane can produce a local change in the membrane potential, which causes

electric current to flow rapidly to other points in the plasma membrane, causing

182

further changes in the membrane potential. This traveling transient reversal of

membrane potential is termed the action potential. This process will be discussed in

detail in the following paragraphs.

Opening and closing of transmembrane Na+/K+ ion channels can induce a

departure from the resting potential, called a depolarization. Depolarization is used

commonly to describe the rise in cellular interior voltage, typically from –70 mV to –

65 mV. On the other hand, if the cellular interior voltage becomes even more

negative, for example changing from –70 mV to –80 mV, we call this phenomenon

hyperpolarization. Synaptic inputs to a neuron may cause the membrane to depolarize

or hyperpolarize. In neurons, a sufficiently large depolarization can evoke a short-

lasting all-or-nothing action potential, in which the membrane potential very rapidly

undergoes a large change, often briefly reversing its sign.3, 4, 7-9 The precise

orchestration of action potential depends on the actions of special types of voltage-

gated ion channels.

Figure 5-1 Action potential as a plot of membrane potential vs. time.3

The action potential, as illustrated in Figure 5-1, can be understood in the

following sequence. First, at resting potential, a small subpopulation of the K+

channels are open, permitting small amounts of K+ to enter the cell. Second, as a

183

neuron receives a depolarization current, Na+ ion channels (NaV), open in response,

letting Na+ ions rush into the cellular interior and increasing the cellular membrane

potential, making the interior more positive. Third, if enough NaV open such that the

membrane potential increases past the threshold (~ -65mV), more sodium ion channels

open, taking the membrane potential through the rising phase of the action potential.

At the same time, more potassium ion channels open to allow for the exit of potassium

ions from the cellular interior. In other words, in the rising phase of the action

potential, the NaV channels are open, and the potassium channels are also open,

resulting in the overall entrance of Na+ ions into the cells and K+ ions exiting the cells.

Next, at the peak of the action potential, the sodium channels close, while potassium

channels remain open and K+ ions continue to leave the cell. The efflux of potassium

ions decreases the membrane potential or hyperpolarizes the cell, leading the cell into

the falling phase of the action potential. Finally, as the membrane potential

hyperpolarizes, potassium ion channels start to close, allowing the membrane potential

to recover, via Na+/K+ ion pump ATPase transport. 2, 3, 10, 11

The proper response (opening) of sodium channels to depolarization dominates

the first phase of an action potential, overwhelming any currents associated with other

ion leakage channels and causing further depolarization of the membrane. In this

“rising phase” of an action potential, the membrane potential rapidly approaches the

sodium equilibrium potential as sodium ion channels represent most of the

membrane’s ionic conductance. In addition, this “all or nothing” quality of the action

potential implicates that once a certain threshold potential is achieved, an action

potential is “fired”. The various characteristics of action potential depend not on the

initial stimuli, but rather, on the organization and concentration of different ion

channels in the neuron. The action potential is propagated along the membranes of

excitable cells via the opening-closing of voltage-gated Na+ and K+ channels and,

along with synaptic neurotransmission, forms the basis of all neuronal signaling. In the

following, the discussion focuses on sodium ion channels.

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Sodium ion channels 5.1.2

As mentioned in 5.1.1, voltage-gated sodium channels (NaVs) serve as an

essential component of the molecular machinery that controls initiation and

propagation of action potentials in electrically excitable cells.3, 4, 12 NaVs are large

integral membrane complexes and are expressed as a ~260 kDa pore-forming -

subunit with one or two ancillary 33–36 kDa -proteins. Sequence analysis and

homology modeling, extensive mutagenesis data, and cryoelectron microscopy13

reveal that the ion conducting -subunit assumes a pseudo-fourfold symmetry in

which reentrant loops from each of four domains create the outer vestibule and

selectivity filter.14-17 The channel gating properties and ion transport kinetics are

refined through extensive post-translational processing of the -subunit and by

association with the auxiliary -proteins. Intrinsic differences in the ten known

mammalian sodium channel -subtypes and four -auxiliaries, their expression levels,

tissue distribution, and subcellular patterning help shape a vast ensemble of neural

processes.18 Not surprisingly, misregulation of NaV function is associated with

profound pathologies such as neurodegenerative disease, epilepsy, and chronic pain.19

A desire to understand the precise role of NaV isoforms in signal conduction and their

causative role in neuropathic disease motivates the development of molecular tools to

probe dynamic processes associated with this protein class.

In nerve cells, NaVs accumulate at the axon initial segment and nodes of

Ranvier through an intricate network of relations with -subunits, cellular adhesion

molecules, and cytoskeletal adaptor proteins.10, 11, 20 The distribution of NaVs to specific

cellular locales is achieved through a combination of protein sorting, targeted

transport, and endocytosis.21, 22 The precision and responsiveness of these NaV

regulatory pathways are tightly controlled to shape the specific electrophysiological

properties of the cell in response to firing activity, neuronal development, myelination,

and nerve injury. Recent studies present compelling evidence that functional NaVs

assemble in sub-diffraction-sized neuritic structures, intimating a multifaceted set of

roles for these proteins beyond axonal depolarization.23 Such data is beginning to

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reveal how compartmentalization of NaVs in these highly fluxional neuronal

substructures can fine-tune responses to the network of inputs that are necessary for

synaptic transmission. Deciphering the mechanisms that impart spatiotemporal control

of NaV expression is thus tightly entwined with understanding how these proteins

orchestrate the diverse processes that underlie neurological function.24-26

Our present understanding of NaV function at the systems level is shaped

through a combination of cyto- and immunohistochemical studies on fixed cells27, 28

and tissue8, and the ability to record real-time changes in ionic currents using the tools

of electrophysiology. Fluorescent sodium ion sensors find applications for live cell

imaging, but lack sensitivity to detect submillimolar changes in local concentration

gradients and give no information on NaV activity below the action potential

threshold.29 While GFP-labeling of individual NaV isoforms is viable30-35,

electrophysiological measurements reveal that attachment of the fluorescent protein to

the NaV -subunit greatly diminishes plasma membrane expression levels.36, 37 Studies

with NaV fusion proteins may be further compromised by improper protein folding

and/or possible alterations in channel gating kinetics.38, 39 Related experiments with

fluorescently labeled proteins that contain specific NaV binding sequences reveal, at

most, only a subset of factors involved in the trafficking of intact NaVs.40 Imaging tags

based on naturally occurring NaV peptide toxins are available, but their use for

tracking NaV dynamics is disadvantaged by the complex and/or poorly defined binding

kinetics of these probes.41-43 Notably, no existing probes demonstrate applicability in

live cell imaging of endogeneous NaVs at the single-molecule level. Accordingly, new

technologies capable of detecting native, functional NaV proteins in living cells at the

highest degree of resolution are desired.

Saxitoxin 5.1.3

Nature provides a collection of naturally occurring, topologically unique

guanidinium-derived poisons that bind with low nanomolar affinity to NaVs (see

Figure 5-2 for computational model).44 These compounds, arguably the most fabled of

which are tetrodotoxin (TTX) and saxitoxin (STX), bind with nanomolar potency to

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six of the ten known mammalian sodium channel isoforms (NaV1.1–1.9, NaX).

Mutagenesis data, electrophysiology studies, and computational modeling give

evidence for reversible, state-independent binding to the outer mouth of the channel

pore (Figure 5-2). Owing to their exquisite selectivity and favorable binding kinetics,

these molecules are recognized as indispensable tools for electrophysiology. For our

purposes, these toxins also serve as blueprints for the design of NaV-selective imaging

agents. Investigations in the DuBois laboratory have demonstrated that a number of

structural modifications to STX are permissive without abrogating channel-binding

function. Access to derivative forms of the natural product are made possible through

de novo chemical synthesis.45-49

Figure 5-2 Computational model of STX bound in the sodium channel pore, side

view. Domain I (red); domain II(green); domain III(yellow); domain (blue). Selectivity

filter residues are shown as spheres; outer vestibule residues are shown as lines; STX

is shown in ball-and-stick model. For clarity, some domain IV resides have been

removed.50

In this work, we developed NaV-selective, fluorescent small molecules,

patterned after STX, which enable real-time imaging of NaVs at the single-molecule

level. The design and preparation of these unique probes are described together with

electrophysiological characterization and fluorescence microscopy experiments using

an established cellular model for neuronal differentiation. Super-resolution

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reconstruction of sparse single-molecule positions 51, 52 makes possible the localization

of NaVs to within ~40 nm in small cellular features that include neuritic spines and

growth cones of filopodia, a level of structural detail well below the diffraction limit

of conventional fluorescence imaging (~250 nm). Such findings, together with single-

molecule tracking of individual NaVs, underscore the versatility of STX-based

fluorogenic probes for sodium channel study.

Experimental 5.2

This section describes the synthetic design, electrophysiology, confocal

microscopy, wide-field microscopy, and data analysis for this study.

Synthetic design of fluorescent STX 5.2.1

Investigations by Dubois et al. have demonstrated that a number of structural

modifications to STX are permissive without abrogating channel-binding function.

Access to derivative forms of the natural product have been made possible through de

novo chemical synthesis.45-49 Figure 5-2 and Figure 5-3 illustrate the binding of (+)-

saxitoxin to the sodium ion channel pore via Pymol rendering of computational

models. The site of STX modification is pointing away from the channel pore, and

was shown previously to not interfere with the binding of saxitoxins to NaVs. An NHS

ester was furnished onto saxitoxin on the N21 site, and then reacted with the amine

functionalized fluorophores Cy5 and DCDHF (Figure 5-4). Cy5 and DCDHF were

chosen for their excellent emission and cell imaging properties. For a more in-depth

discussion of desirable properties for cell imaging, see Chapter 1.

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Figure 5-3 Natural and synthetic saxitoxins for NaV studies. Computational model

of STX ligated in the outer mouth of the NaV pore. The site of STX modification

(N21) is indicated by an arrow.1

Figure 5-4 Natural and synthetic saxitoxins for NaV imaging. Synthesis of

fluorescent STX derivatives STX-Cy5 and STX-DCDHF from STX-NH3+ using a

selective chemical ligation method.1

Cell culture 5.2.2

We chose to use two systems to illustrate the specificity and applicability of

the fluorescent saxitoxins: Chinese hamster ovary cells (CHO) and

pheochromocytoma 12 (PC12) cells. CHO cells express no endogenous NaVs and are

therefore good negative controls; PC12s are well established to be a good neuronal

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cell model, expressing endogenous NaVs that can be easily up-regulated by NGF-

induced differentiation.

CHO Cell Culture 5.2.2.1

Chinese Hamster Ovary (CHO) cells were grown in RPMI 1640 phenol red-

free media (Gibco BRL, Grand Island, NY) supplemented with 10% fetal calf serum

(HyClone, Logan, UT), 10mM HEPES. (4-(2-hydroxyethyl)-1-

piperazineethanesulfonic acid), 1mM sodium pyruvate, 20µM 2-mercaptoethanol (2-

hydroxy-1-ethanethiol) and 0.1mM nonessential amino acids, 100units/ml penicillin,

100µg/ml streptamycin, 10µg/ml gentamicin, 0.5mg/ml geneticin (Gibco, BRL, Grand

Island, NY), and kept in 5% carbon dioxide incubators at 37 ºC. Cells were grown on

chambered coverglass (Nalge Nunc International, Naperville, IL). To facilitate

adhesion of cells, the coverglass was coated with 50ug/ml fibronectin (human plasma,

CalBiochem, San Diego, CA) in phosphate buffered saline, PBS, pH 7.4 (Gibco BRL,

Grand Island, NY) for 1hr at room temperature prior to deposition of cells.

PC 12 Cell Culture 5.2.2.2

PC12 (pheochromocytoma 12) cells were grown in DMEM phenol red-free

media (Gibco BRL, Grand Island, NY) supplemented with 10% horse serum (Gibco

BRL, Grand Island, NY), 5% fetal calf serum (HyClone, Logan, UT), 100 units/mL

pen-strep (Gibco, BRL, Grand Island, NY), and kept in 5% carbon dioxide incubators

at 37 °C. For fluorescence imaging experiments, PC12 cells were grown on

chambered coverglass (Nalge Nunc International, Naperville, IL) or glass bottom

dishes (MatTek Corp, Ashland, MA). To facilitate adhesion of cells, the chambered

coverglass was coated with 50 g/mL collagen (human plasma, CalBiochem, San

Diego, CA) in phosphate buffered saline (PBS), pH 7.4 (Gibco BRL, Grand Island,

NY) for 1 h at room temperature prior to deposition of cells. Cells were induced to

differentiate with 100 ng/µL solutions of nerve-growth factor NGF (BD Biosciences,

Bedford, MA) prior to imaging. Throughout the course of single-molecule imaging

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experiments cells were kept at 37 °C and 5 % CO2 using a stage heater and CO2

chamber (Narashige, Japan) on the microscope.

Electrophysiology experiments 5.2.3

To measure the potency of the fluorescent saxitoxins to achieve NaV block,

electrophysiology experiments were conducted on PC 12 Cells. Sodium currents were

measured using the voltage-clamp technique in the whole-cell configuration.

Borosilicate glass micropipettes (Sutter Instruments, Novato, CA) were fire-polished

to a tip diameter yielding a resistance of 1.2–2.0 MΩ in the working solutions. The

pipette was filled with (in mM): CsCl 140, KCl 1, EGTA 1, HEPES 10, and the pH

was adjusted to 7.2 with solid CsOH. The external solution had the following

composition (in mM): NaCl 140, CaCl2 1, HEPES 10, and the pH was adjusted to 7.4

with solid CsOH. Stock solutions of STX-Cy5 and STX-DCDHF were prepared by

serial dilution with external solution and stored at 4 °C in the dark. Current

measurements were recorded under continuous perfusion, controlled manually by

syringe addition.

Signals were amplified with an Axopatch-200B amplifier (Axon Instruments,

Union City, CA), as previously described by Moran.53 The output of the patch-clamp

amplifier was filtered with a built-in low-pass, four-pole Bessel filter having a cutoff

frequency of 10 kHz and sampled at 100 kHz. The membrane was kept at a holding

potential of –100 mV. Pulse stimulation and data acquisition used 16 bit D-A and A-

D converters (Axon Instruments Digidata 1322A) controlled with the PClamp

software (Axon Instruments). Leak currents were subtracted using a standard P/4

protocol of the same polarity. Access resistance was always < 4 MΩ and the cell

capacitance was between 4 and 20 pF, as measured by the compensating circuit of the

amplifier. All measurements were done at room temperature (20–22 °C). Recordings

were made at least 2 min after establishing the whole-cell and voltage-clamp

configuration to allow for stabilization of the voltage-dependent properties of the

channels. Currents were elicited by 10 ms step depolarizations from a holding

potential of –100 to 0 mV. Data were normalized to control currents, plotted against

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toxin concentration and analyzed using custom software developed in the Igor Pro

environment (Wavemetrics). Data were fitted to Langmuir isotherms to elicit IC

values and expressed as mean ± SE.

Confocal microscopy on CHO and PC12 cells 5.2.4

To demonstrate specificity and illustrate the applicability of fluorescent

saxitoxins to biologically relevant experiments, we conducted confocal fluorescence

microscopy on both CHO and PC 12 cells. Confocal images were acquired on a Zeiss

LSM 510 Confocal Laser Scanning Module mounted on a Zeiss Axiovert 200M

inverted microscope in the Stanford Cell Science Imaging Facility. A LCI Plan-

Neofluar 25x/0.8 lmm Korr DIC objective was used for Figure 5-16 a–e and Figure 5-

16 h–m and an A-Plan-Apochromat 63x/1.4 Oil DIC objective was used for Figure 5-

16 f, Figure 5-16 g, and Figure 5-15. Cy5 and DCDHF were excited at 633 nm with a

5mW HeNe laser at 10–15% output and emission was detected through a 650 nm

long-pass filter; secondary antibody-Alexa488 was excited at 488 nm with a 25 mW

Argon laser at 10% output and emission was detected through a 500-550 nm IR

bandpass filter.

For quantification measurements, average laser power was measured using a

FieldMaster GS power/energy analyzer (Coherent, Auburn, CA) and images were

collected at constant laser transmission and acquisition settings. Fluorescence images

were collected as a set of ten z-stacks traversing the cells and projected as a

composite. Quantification was performed using Volocity software; regions of interest

corresponding to the perimeter of the cells were manually selected to obtain voxel

counts. All intensities were corrected for background fluorescence. Data analysis and

plotting were performed using Igor Pro software.

Wash off Experiments 5.2.4.1

Cells were plated on glass-bottom dishes and incubated at 37 ºC with 5% CO2

in phenol red-free DMEM supplemented with 10% CCS and 1:100 v:v pen-strep in the

presence 100 ng/L solutions of NGF for 4–7 days. Immediately prior to imaging,

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media was replaced with phenol red-free DMEM containing 1:100 v/v pen-strep, 100

ng/L NGF, and 10 nM STX-Cy5 or 15 nM STX-DCDHF. Cells were incubated for

15 min under ambient conditions, and pre-wash off images were acquired over a 15

min period. Media was then aspirated and fluorophore-free media of the same

composition was added. Media was replaced an additional two times and post-wash

off images were acquired over a 15 min period. This procedure was repeated using

five different glass bottom plates for each STX-fluorophore. Three areas on each plate

containing 90–120 cells were imaged both pre- and post-wash off (Figure 5-16 a, b).

Absolute fluorescence intensities were quantified as described above (5.2.4) and

normalized to the average intensity of images obtained in the presence of STX-Cy5 or

STX-DCDHF (Figure 5-17). Sample areas were selected at random before and after

wash-off; each image represents a unique population of cells.

± NGF Experiments 5.2.4.2

Cells from the same passage were plated on separate glass bottom dishes and

incubated for 4-7 days at 37 ºC and 5% CO2 in phenol red-free DMEM supplemented

with 10% FCS and 1:100 v/v pen-strep in the presence or absence of 100 ng/L NGF.

Media were replaced prior to imaging with phenol red-free DMEM containing 1:100

v/v pen-strep with or without 100 ng/L solutions of NGF and 10 nM STX-Cy5 or 15

nM STX-DCDHF. Cells were incubated for 15 min under ambient conditions and

fluorescence images were acquired. This procedure was repeated using five different

sets of +/- NGF glass bottom plates for each STX-fluorophore. Three areas on each

plate containing 90–120 cells were imaged (Figure 5-16 d, e). Absolute fluorescence

intensities were quantified as described above (5.2.4) and normalized to the average

intensity of images of NGF-differentiated cells obtained in the presence of STX-Cy5

or STX-DCDHF (Figure 5-18).

Colocalization Experiments 5.2.4.3

To illustrate that fluorescent saxitoxins specificity and its applicability of

utility as probe for NaV distribution, we doubly labeled differentiated PC 12 cells via

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immunofluorescence in addition to fluorescent saxitoxins. We then assessed how the

fluorescent saxitoxins and antibodies to NaVs colocalized via a Mander’s Coefficient

analysis.

Sample Preparation 5.2.4.3.1

After 2 days of NGF induction, coverglass-plated PC12 cells were first washed

with cold PBS buffer pH 7.4 and then fixed for 15 min in cold 3.7% paraformaldehyde

in PBS pH 7.4. Cells were then rinsed 3 times by displacing the paraformaldehyde

solution with fresh PBS buffer. Fixed cells were then permeabilized/pre-blocked with

a block solution made with 0.15% Triton-X 100 (Omnipur, EMD) and 1.5% BSA

containing PBS buffer for 1 h at room temperature. Permeabilized cells were then

treated with 1 g/mL primary antibody (polyclonal rat anti-pan NaV, Spring

Bioscience, Pleasanton, CA) in block solution (0.15% Triton-X and 10% FCS in PBS)

for 8 h at room temperature. Cells were immersed in fresh PBS buffer followed with 3

rinses with fresh PBS buffer before being treated by the block solution for 1 h.

Following this second pre-block step, the cells were treated with secondary antibody

conjugate to Alexa488 (Invitrogen) for 6 h at 4 °C. Cells were then washed 5 times

with PBS and treated with 10 nM STX-Cy5 or 15 nM STX-DCDHF in PBS for 15

min before imaging. Confocal images were acquired sequentially in multi-track mode

and detected through a HFT UV/488/543/633 dichroic beam splitter with a 650 nm

long-pass filter for STX-Cy5 and STX-DCDHF and a HFT 488 dichroic beam splitter

with a 500–550-nm IR bandpass filter for Alexa488 (Figure 5-16 h–m).

Mander’s Colocalization Coefficient 5.2.4.3.2

Confocal images were acquired on a Zeiss LSM 510 Confocal Laser Scanning

Module mounted on a Zeiss Axiovert 200M inverted microscope. For quantification

measurements average laser power was measured using a FieldMaster GS

power/energy analyzer (Coherent, Auburn, CA) and images were collected at constant

laser transmission and acquisition settings. 3 ROIs were chosen for both STX-

DCDHF and STX-Cy5 stained PC12 cells. Fluorescence images in 2 colors (STX-

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Fluor, and Alexa488) were collected as a set of ten z-stacks traversing the cells and

every substack was used in this quantification. Quantification was performed using an

Image J plugin (JACoP v2.0). In every substack, the Mander’s54 was calculated using

the following equation. Averages of Mander’s colocalization coefficient over the ten

stacks of the 3 different ROIs are reported for each fluorescent STX.

∑

∑

Equation 5-1

Briefly, Mander’s Coefficient M is defined by summing intensities of co-

localizing pixels from one channel and dividing it by its integrated intensity. A pixel

from channel STX-fluor (PSTX)is considered as co-localized if it has a non-zero

intensity counterpart in the antibody channel. As a result, Mander’s coefficient gives

an estimate of the amount of co-localizing signal from a STX channel over the

antibody channel, without making any assumption on the stoichiometry it may adopt.

The value of 1.0 for Mander’s colocalization coefficient indicates perfect

colocalization.54

Confocal and Wide-field imaging of CHO cells transfected to express NaV 5.2.4.4

rNaV1.4 plasmid information 5.2.4.4.1

The nucleotide sequence of the rNaV1.4 gene used in these studies (sequenced

by Sequetech Corporation, Mountain View, CA) was aligned with the published gene

sequences of the two channel isoforms, NaV1.7 (GenBank accession no. AAB50403)

and NaV1.2 (GenBank accession no. NM_012647), induced in NGF differentiated

PC12 cells using an academic FASTA sequence comparison software program

(University of Virginia; http://fasta.bioch.virginia.edu/fasta_www2/fasta_list2.shtml).

This analysis shows nucleotide identities of 70.7% for rNaV1.4 and NaV1.7 (PC12)

and 70.0% for rNaV1.4 and NaV1.2 (PC12), and confirms that the proteins are nearly

identical in the pore region surrounding the key residues responsible for STX binding

(D400, E755, K1237, and A1529, NaV1.4 numbering) (Figure 5-5).

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Figure 5-5 Sequence comparison of the pore region in rNaV1.4, NaV1.7 (PC12), and

NaV1.2 (PC12). Amino acid sequences in the STX binding site are identical.1

Confocal microscopy experiments 5.2.4.4.2

Wild-type CHO cells were plated on plastic 6-well plates in 2 mL media and

grown to 40–50% confluency. Cells were transfected with an expression vector

containing the full-length cDNA coding for rat NaV1.4 (rNaV1.4) sodium channel α-

subunit using the method of calcium phosphate precipitation; cotransfection with

eGFP was used as a marker of transfection efficiency. Transfected cells were allowed

to grow for 48 h, plated and grown on glass-bottom dishes for 2 h until adherent, and

incubated in imaging media containing 40 nM STX-Cy5 or 60 nM STX-DCDHF for

15 min prior to data collection. Samples of untransfected CHO cells were prepared in

the same fashion. Confocal images were acquired sequentially in multi-track mode

and detected through a HFT UV/488/543/633 dichroic beam splitter with a 650 nm

long-pass filter for STX-Cy5 and STX-DCDHF and a HFT 488 dichroic beam splitter

with a 500–550 nm IR bandpass filter for eGFP.

Wide-field microscopy experiments 5.2.4.4.3

Wild-type CHO cells were plated onto 400µL fibronectin-coated borosilicate

coverglass and grown to 85% confluency. 500 ng NaV 1.4 (or NaV1.7) DNAs and 100 ng

GFP DNAs (fluorescence proteins) were diluted into 50µL OptiMEM and mixed

gently by tapping on the side of the eppendorf tube. Lipofectamine TM complexes

were prepared by diluting 1 µL of Lipofectamine TM into 50 µl OptiMEM and

incubating for 5 min at room temperature. The DNA mix dilution was then combined

with the diluted Lipofectamine reagent; 30 min of incubation time at room

temperature was used to allow DNA-liposome complexes to form. The growth

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medium on the cells were replaced with 300 µl DMEM with 10% FCS. The DNA-

liposome containing medium was then added drop-wise onto the cells which were then

incubated for 8 h at 37˚C in a CO2 incubator. Following incubation, the transfection

mixture was replaced with complete growth medium as described in Section 5.2.2.2.

Ampicillin was also used as a selection antibiotic. Continued cell growth was allowed

for 24 hours, and cells were then imaged using the epi-fluorescence set-up as

described in Section 5.2.5.1 after being labeled using the same protocol as described in

Section 5.2.5.2. In addition to the 633 nm and 594 nm lasers used for exciting

fluorescent STXs, a 488 nm Argon ion laser was also used in the excitation path to aid

in finding successfully transfected CHO cells. Operationally, only CHO cells

exhibiting green fluorescence were deemed to have been successfully transfected with

NaV DNA.

Single-molecule microscopy 5.2.5

The fluorescent saxitoxins exhibit favorable emitter properties and can be used

in single molecule and sub-diffraction localization of NaV distributions. In the

following, we detail experiments illustrating these points.

Imaging setup 5.2.5.1

Both transmission images of the cells and epi-fluorescence images of single

molecules were acquired using an inverted microscope (IX71, Olympus, Center

Valley, PA). Brightfield illumination from a condenser allowed for visualization of

the edges of the cells. Fluorescence imaging of the cells was performed with wide-

field epi-illumination in an area of ~42μm × 42 μm. Because the mechanical drift of

the stage (Semprex) was negligible during the recording time, a fiduciary correction

was not necessary. Laser illumination at 633 nm afforded an intensity of ~200W/cm2

at the sample plane for Cy5 imaging. The 594 nm laser illumination provided

~40W/cm2 for the DCDHF fluorophore. Epi-fluorescence images were collected with

a 60× magnification, 1.45 NA, oil-immersion objective (PlanApo, Olympus, Center

Valley, PA) and imaged through a 640 nm long-pass filter and a 635 nm dichroic

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mirror (Omega Optical Inc., Brattleboro, VT) for images acquired with 633nm

excitation, and 595RDC and 615LP for the images acquired the 594nm excitation. An

EMCCD-camera (IXonDV887, Andor, South Windsor, CT) was used for data

collection. The pumping beam intensity was adjusted to give an acceptable signal-to-

noise ratio with neutral density filters.

Sample Preparation 5.2.5.2

Differentiated PC12 cells were plated 1-2days on collagen-coated borosilicate

coverglass and differentiated by 100ng/mL NGF as described in the Cell Culture

section. Cells were briefly washed with PBS pH 7.4 (Gibco) before immersion into

either STX-DCDHF (13 nM) or STX-Cy5 (3nM or 10nM) in phenol red-free DMEM.

The samples were then placed in a Tokai Hit thermo incubator maintained at 37C

with 5% CO2, which is fixed onto the Semprex precision stage via a screwed-in

custom-machined adaptor.

Wide-field image and data analysis 5.2.6

Maxplot Stack 5.2.6.1

Figure 5-6 Schematic for maxplot stacking of a set of images. At the end of maxplot

stacking, the resulting image would have each of its pixels exhibit that pixel’s highest

intensity through the duration of the original set of images.1

From raw image stacks, maxplot images were obtained using ImageJ function

Z Project with maximum intensity. Briefly, this function takes the highest intensity a

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pixel exhibits through time as the intensity of that pixel. See Figure 5-6 for a

schematic.

Single-Particle Tracking 5.2.6.2

Differentiated PC12 cells are labeled with 12.8µM STX-DCDHF in phenol-red

DMEM. All movies were recorded at 50 ms integration time with continuous epi-

illumination. The successive (x,y) positions of the single molecules on the plane of the

cell surface were recorded as a function of time. In order to investigate the behavior of

single NaV in a consistent fashion, only the fluorescent spots that appeared on the cell

after the start of illumination and disappeared before the end of recording were

considered. The visibility time (τR) is defined as the time from the initial appearance of

a single-molecule spot on the membrane (from the solution) to the time the single

molecule disappears from the imaging focal plane in a single step.

Trajectories of single conjugates were extracted by using a Single Particle

Tracking (SPT) program from ETH-Zurich (while now available as an ImageJ plugin,

http://www.mosaic.ethz.ch/Downloads/ParticleTracker, a previous Java version was

used). In this approach, particle positions were iteratively refined by using the

intensity centroid for sub-pixel interpolation. Trajectories were computationally

extracted from the recorded movie sequences using the feature-point tracking

algorithm described in Ref.55. All trajectories were visually inspected to insure that the

tracking program was operating properly. The mean-squared displacement from each

individual single-molecule trajectory (truncated to 10 time steps, typically at 0.050 s

per time step) was used to extract an observed short-time diffusion coefficient D. As

usual, each single-molecule trajectory produced an apparent diffusion coefficient, D,

which can be regarded as an estimate of the true diffusion coefficient of the particles.

The distribution of apparent diffusion coefficients for the individual trajectories was

generated to test for heterogeneity in the motion. To perform this test, an estimate of

error is needed. Under the hypothesis that all molecules are moving as unconstrained

Brownian diffusers, the distribution of estimates of D follows a well-known shape 56

http://www.mosaic.ethz.ch/Downloads/ParticleTracker

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( )

( ) (

) ( ) (

)

Equation 5-2

where D is the diffusion coefficient estimate extracted from an individual trajectory,

D0 is the underlying Brownian diffusion coefficient, and N is the number of time steps

in each trajectory. This expression was used to generate the smooth curves in the text

by fixing N=10 and using the arithmetic mean of observed D values for D0.

Super-resolution imaging and reconstructions 5.2.6.3

STX-Cy5 on NaV for Target-Specific PAINT (Points Accumulation for 5.2.6.3.1

Imaging in Nanoscale Topography)

Figure 5-7 Fluorescence intensities over time of STX-Cy5 on NaVs in the membrane

of NGF differentiated PC12 cells and surface immobilized Cy5-NHS. (A)

Immobilized Cy5 signal decays exponentially over time while binding of new STX-

Cy5 to NaVs affords sustained fluorescence at the cell membrane. (B) Cy5

immobilized on a polyelectrolyte multilayer. (C) STX-Cy5 from solution binding to

NaVs in the cell membrane.1

Pointillist super-resolution imaging schemes rely on efficient turn on/off events

both for maintenance of sparse concentrations of emitters in each image and for time

economy in imaging; this efficiency is especially important to be able to image at the

rate of the specific biological process in live samples. In Figure 5-22(l) in 5.3.9, we

show that our imaging rate is on par with the highly dynamic neuritic structures on

PC12 cells. The efficient appearance of emitters (i.e. “on”) is enabled by the

continuous binding of new STX-Cy5 from solution to the NaV in the cellular

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membrane. The precise manner for the disappearance (i.e. “off”) of emitters from the

image is likely a combination of photobleaching and blinking. The mechanism of the

“turn-off” events will be further substantiated in future experiments. Figure 5-7

illustrates that at the cellular membrane, through time, new binding events of

fluorescent STX from the solution to the NaV affords sustained fluorescence signal at

the membrane. On the other hand, for a system where there is no new fluorescent STX

in solution, in this case, Cy5-maleimide immobilized on a polyelectrolyte multilayer

via a lysine residue, we see a characteristic overall exponential decay in fluorescent

signal.

Given the >10nM concentration of fluorescent STX in solution, in a single

diffraction-limited spot, and given focal depth of 500nm, we would expect a signal-to-

background on the order of or less than 10 to 1. Given cellular auto-fluorescence and

the usual losses in emission collection, the signal-to-background values in our

experiments (1.8±0.4 for STX-Cy5, and 4.2±0.5 for STX-DCDHF), are reasonable.

Super-resolution Localization and Reconstruction 5.2.6.3.2

Figure 5-8 (a) Statistical localization precision for STX-Cy5 on PC12. (b) Steady

number of localizations through imaging time.1

From raw image stacks, super-resolution images were obtained using

previously published image processing techniques as described.57 Briefly, for each

imaging frame, the position of the a single emitter was determined by fitting the signal

above background in a small region of interest containing the single-molecule spot to a

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2-D Gaussian with nonlinear least squares regression analysis (nlinfit, in MATLAB).

Each single-molecule point spread function was fit to determine the following

parameters: background, amplitude, width, center(x) and center(y). Fits with a

localization precision greater than 180 nm were discarded. The resulting statistical

error of localization at 96% confidence interval (2× standard deviation) was found to

have a peak mode at 60 ± 3 nm (Figure 5-8). To reconstruct a super-resolution image,

each single-molecule position was re-plotted using a custom macro written in ImageJ

as a two-dimensional Gaussian having a constant amplitude and σ (standard deviation

or width) equal to the average standard deviation error of the center of the fit, i.e. 30

nm (from 60nm÷2, where 60nm is the peak mode of the statistical localization

precision at 96% confidence internal. See Figure 5-8). This is a conservative value to

use as compared to the theoretical mean statistical localization precision of our data.

The theoretical mean statistical localization precision is described by the following

equation58

√

⁄

Equation 5-3

where s is the standard deviation of the Gaussian fit to the PSF, a is the pixel size, N is

the number of photons , and b is the background noise. In these data we obtained an s

of 340nm, N of 4640 ± 1800 photons/frame, a pixel size of 107 nm, and b of

21±1photons/pixel. Using these parameters, the theoretical mean statistical

localization precision is ~25±3nm, smaller than the width used in our reconstructions,

30nm. Since every molecule is plotted identically, the brightness in the plot is

reasonably proportional to density of STX binding events. Lastly, given the 50ms

exposure time and 5300 localizations/s, as afforded by the new binding events of

STX-Cy5 (Figure 5-8b), 5s was sufficient to reconstruct axonal structure (Figure 5-22

a).

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Results and discussion 5.3

Studies by Dubois et al. on the chemical synthesis of (+)-STX have enabled the

furnishing of an efficient and flexible synthetic route to not only the natural toxin, but

also to structural variants that are inaccessible by other means. Despite the absence of

a protein crystal structure for any member of this family of ion channels, mutagenesis

data and structure-activity relationship studies with TTX, STX, and a small collection

of naturally occurring congeners, have informed their efforts to modify STX through

rational design. This work has demonstrated that the STX receptor site can

accommodate changes to the N21 position of STX (Figure 5-3) with minimal to

modest loss in affinity between ligand and protein.48 The availability of STX-NH3+

(Figure 5-4) through chemical synthesis offers a convenient starting material from

which to append other functional groups through mild and selective conjugation

strategies. In this way, we have been able to fashion fluorescent STX conjugates.

For our studies, cationic Cy559 and a neutral DCDHF fluorophore (specifically

DCDHF-V-P found in reference60) were selected as bright, photostable, long

wavelength fluorescent dye molecules that are particularly suited for experiments with

live cells61 and for application of single-molecule imaging techniques. The availability

of activated N-hydroxysuccinimide (NHS) ester derivatives Cy5-NHS and DCDHF-

NHS enabled “post-synthetic” coupling reactions with STX-NH3+ to furnish STX-

fluorophore conjugates, STX-Cy5 and STX-DCDHF (Figure 5-4). Purification by

reverse-phase HPLC afforded the pure trifluoroacetate salts of these compounds as

water-soluble blue and violet powders, respectively, which can be stored for several

months at –20 °C when shielded from light. To the best of our knowledge, STX-Cy5

and STX-DCDHF represent the first examples of fluorescently labeled paralytic

shellfish toxins.62 In the following, I detail the key findings of our experiments.

Photo-physical properties of STX-Cy5 and STX-DCDHF 5.3.1

Live-cell fluorescence imaging requires bright, stable, long-wavelength-

emitting probes that are soluble and stable in aqueous cell media.63 For more details,

please see Chapter 1. Both Cy5 and DCDHF exhibit high total numbers of emitted

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photons and large fluorescence quantum yields in environments appropriate for cell

viability,59, 60 characteristics that minimize the requisite power of the excitation source

and potential for cellular photo-damage. Short data acquisition times also help to limit

cumulative exposure during experiments and enable rapid data collection while

monitoring dynamic cellular processes. Direct and oxidative injury to the cells is

further attenuated as a result of the red-shifted absorbance and emission characteristics

of these probes (STX-Cy5, abs/em = 655/665 nm; STX-DCDHF, abs/em = 580/640 nm

in H2O, Figure 5-9). At the excitation wavelengths employed for these probes (ex =

594, 633 nm) auto-fluorescence from endogenous fluorophores (e.g., NADPH, flavin

cofactors, aromatic amino acids) is negligible.64 The properties of these labels specific

to single-molecule imaging will be described below.

Figure 5-9 Absorbance/emission spectra of STX-Cy5 and STX-DCDHF.

Absorbance (dotted lines) and emission (solid lines) of STX-Cy5 (blue) and STX-

DCDHF (violet) in PBS solution at pH 7.4. STX-Cy5, λabs/em = 655/665 nm; STX-

DCDHF, λabs/em = 580/640 nm.1

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Figure 5-10 Fluorescence enhancement of membrane-bound STX-DCDHF. 12.8

μM STX-DCDHF was used. 3 day differentiated PC12 cells were detached from

culturing flasks, pelleted, and re-suspended in PBS to make a cell suspension solution.

The resulting cell suspension had ~5E6 cells/mL. The figure shows the relative

fluorescence enhancement (~10×) in cell suspension (red line) compared to PBS

(violet line) as a result of the viscosity/polarity sensitivity of the DCDHF fluorophore.1

An additional favorable property of the STX-DCDHF is its environmental

sensitivity (please see Chapter 1 for fluorophore considerations, and Chapter 4 for

details about the DCDHF family of fluorophores). STX-DCDHF brightens in more

viscous conditions such as the cellular membrane, enabling higher signal-to-

background in imaging. In fact in a bulk-level fluorescent emission experiment, we

find that fluorescence emission of STX-DCDHF was found to be 10 times larger in

cell suspension compared to in PBS (Figure 5-10). In cellular imaging experiments,

this property afforded favorable signal-to- background ratios for signal molecules on

cell surfaces even at somewhat high STX-DCDHF concentrations in solution.

Fluorescent saxitoxins bind specifically to NaV and not to the plasma 5.3.2

membrane

Examined via confocal microscopy, the majority of CHO cells successfully

transfected to express NaV were labeled by fluorescent saxitoxins. This is evidenced

by Figure 5-11; we see that only CHO cells that expressed cytosolic eGFP (sign for

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successful transfection and is indicative of NaV expression) also showed membrane

labeling by the two different fluorescent saxitoxins (STX-Cy5 and STX-DCDHF), and

vice versa. Small populations of cells showing exclusively eGFP expression or STX-

fluorophore binding were also visible.

Figure 5-11 Confocal images of specific rNaV1.4 labeling by STX-fluorophores in

CHO cells. Cells were transfected with rNaV1.4 and eGFP. Media for CHO cells

contained 40 nM STX-Cy5 or 60 nM STX-DCDHF. (A) Transmission image of

CHO cells transfected with eGFP and rNaV1.4. (B) eGFP fluorescence image of A.

(C) STX-Cy5 fluorescence image of A. (D) Merge of B and C. (E–H) Corresponding

images of cells treated with STX-DCDHF.1

In a wide-field fluorescence microscopy experiment, we found very similar

results. Operationally, only CHO cells exhibiting green fluorescence were deemed to

have been successfully transfected with NaV DNA (Figure 5-12 a-c, g,h). Fluorescent

STX signals were only found on successfully transfected CHO cells, and none were

observed from CHO cells that are not transfected (Figure 5-12 d-f, i-k).

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Figure 5-12 Single-molecule wide-field images of specific rNaV1.4 labeling by STX-

fluorophores in CHO cells. Media contained 15 nM STX-Cy5 or 100 nM STX-

DCDHF. (A) Transmission image of a representative CHO transfected with both

eGFP and rNaV1.4. (B) eGFP fluorescence image of A. (C) STX-Cy5 fluorescence

image of A showing several single-molecule spots. (D) Transmission image of a

wild-type CHO cell. (E) STX-Cy5 fluorescence image at the cover glass supporting

the cell in D. (F) STX-Cy5 fluorescence of the cell in D showing no labeling at the

apical membrane. (G–K) Corresponding images of cells treated with STX-DCDHF

(eGFP image not shown). For J and K, diffuse fluorescence arises from STX-DCDHF

in solution and adhered to the fibronectin coated cover glass.1

NaV blocking by STX, STX-Cy5 and STX-DCDHF in NGF-differentiated 5.3.3

PC12

As mentioned, pheochromocytoma 12 (PC12) cells are neoplastic rat adrenal

medulla cells that find extensive use as a model for neuronal differentiation.65

Exposure of this cell type to nerve-growth factor (NGF) potentiates differentiation to a

neuronal phenotype, engendering characteristics that include neurite extension,

suspended proliferation, and gated ion current. Whole-cell voltage clamp recordings

show that treatment of PC12 with NGF for 3 to 14 days induces surface expression of

functional NaVs. Electrophysiological and molecular hybridization studies indicate

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that membrane excitability results from upregulation of NaV1.266 and NaV1.7.67 Due to

the predominant expression of these STX-sensitive isoforms and extensive

characterization of the biochemical pathways potentiated by NGF,68 we have used this

functional neuronal cell model to interrogate the potency and performance of STX-

based fluorescent probes.

Figure 5-13 Sodium ion channel current blocking by natural and synthetic

saxitoxins. Dose-response curves and IC50 values for STX (), STX-Cy5 (), and

STX-DCDHF () on NGF differentiated PC12 cells (mean ± standard deviation, n =

3).1

Prior efforts to measure the affinity of STX towards endogenous NaVs in NGF-

treated PC12 cells employed 3H-STX to determine an apparent Kd in the low

nanomolar range (3.0 ± 0.5 nM).69 To evaluate the ability of STX and derivatives

STX-Cy5 and STX-DCDHF to block sodium ion currents, electrophysiological

measurements were performed against this same cell type following 4–7 day treatment

with NGF. Whole-cell current recordings were made as solutions of increasing

concentrations of STX, STX-Cy5, and STX-DCDHF were applied; the resulting

normalized current densities were fit to a Langmuir isotherm (Figure 5-13).

Consistent with the nanomolar Kd value reported previously for STX, whole cell

voltage clamp recordings give an IC50 for STX of 1.9 ± 0.2 nM.50 Significantly, the

IC50 values of 39 ± 9 and 95 ± 21 nM determined for STX-Cy5 and STX-DCDHF,

respectively, are within 1.5 orders of magnitude of the parent toxin. These results are

commensurate with an STX binding model that orients N21 towards the exposed

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extracellular space.4863 As with the parent toxin, block by both STX conjugates is fully

reversible and peak Na+ ion current is restored upon perfusion of the patched cell with

buffer solution.

Approximate on times for NaV block by STX, STX-Cy5, and STX-DCDHF

were measured by applying solutions at saturating concentrations to NGF

differentiated PC12 cells until complete Na+ current block was observed. Off times

for current recovery from block by each toxin were measured by perfusing with

external solution until the original current was restored. Manual toxin application and

wash-off were performed at a constant rate of ~1 mL/min through a fast exchange

diamond bath perfusion chamber (Warner instruments, Hamden, CT). Rate constants

for Na+ block (kon) and recovery (koff) for each toxin were derived from the Langmuir

model for STX binding and calculated according to the equations: 70

(

)

[ ]⁄

Equation 5-4

where τon and τoff are the measured times to complete block and current recovery,

respectively, and [toxin] is the saturating concentration, taken as twice the measured

IC50 value, for each toxin (Figure 5-14).

Figure 5-14 On/off times and rate constants for Na

+ current block by STX toxins.

(A) Measured τon and τoff values for Na + current block on application of saturating

toxin solutions. (B) Tabulated kon and koff values.1

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Reversible, extracellular, specific NaV labeling by STX-Cy5 and STX-5.3.4

DCDHF in PC12 cells

Figure 5-15 Extracellular labeling of NGF differentiated PC12 cells by STX-

DCDHF. Media contained 15 nM STX-DCDHF. Representative z-stack images from

near the top to the bottom of the cells show that labeling is restricted to the outer

membrane.1

Figure 5-16 Live cell confocal imaging of NaVs. NaVs were labeled with 10 nM

STX-Cy5 or 15 nM STX-DCDHF. (A) NGF differentiated PC12 cells in media

containing STX-DCDHF. (B) Mitigated fluorescence in the same sample following

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media replacement due to the reversibility of STX-DCDHF binding. (C)

Immunostaining image of NaV1.7 in a fixed NGF differentiated PC12 cell, taken from

ref. 68. (D) Undifferentiated PC12 cells from the same passage in media containing

STX-Cy5, showing faint uniform labeling at the cell periphery. (E) NGF

differentiated PC12 cells in media containing STX-Cy5. (F) Live cell image of an

NGF differentiated PC12 cell in media containing STX-DCDHF, showing a

fluorescence pattern similar to E. (G) Transmission image of F. (H–M) NGF

differentiated PC12 cells fixed and exposed to a rat antipan NaV antibody/Alexa488

secondary antibody then treated with PBS solution containing STX-Cy5 or STX-

DCDHF. (H, K) Antibody-Alexa488 fluorescence. (I) STX-DCDHF fluorescence in

H. (L) STX-Cy5 fluorescence in K. (J) Merged image of H and I. (M) Merged image

of K and L.1

NGF-differentiated PC12 cells were treated with 10 nM STX-Cy5 or 15 nM

STX-DCDHF and examined using confocal fluorescence microscopy (Figure 5-15.

Figure 5-16). Reconstitution of z-stack images acquired at times ranging between 15

min and 4 h of incubation in absence of fluorescence in the cytoplasm and in

organelles affirms that STX-Cy5 and STX-DCDHF neither passively diffuse across

the cell membrane nor activate endosomal internalization (Figure 5-15). To confirm

the reversibility of probe binding, the intensity of fluorescence in the presence of 10

nM STX-Cy5 or 15 nM STX-DCDHF was compared to the intensity of residual

fluorescence after media replacement. Consistent with a reversible ligand-protein

binding interaction, the fluorescent signal was effectively absent following

replacement of the cell medium with PBS solution (Figure 5-16 A–B, Figure 5-17).

Figure 5-17 Quantification of probe wash-off and PC12 ± NGF fluorescence.

Media contained 10 nM STX-Cy5 or 15 nM STX-DCDHF. Relative fluorescence of

NGF differentiated cells before and after 3× media replacement.1

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The specificity of probe binding to NaVs was affirmed by colocalization with a

rat anti-pan NaV antibody (Figure 5-16 H-M). Fluorescence images of NGF-

differentiated PC12 cells obtained in the presence of PBS solutions of 10 mM STX-

Cy5 or 15 mM STX-DCDHF after fixation, membrane permeabilization, and

sequential staining with anti-pan NaV antibody and an Alexa488 secondary antibody

yielded Mander’s overlap coefficients of 0.94 0.02 and 0.93 0.08( 5.2.4.3.2),

respectively (Figure 5-16 H–M). The strong overlap of STX-Cy5 or STX-DCDHF

fluorescence signal on top of signal from fluorescent antibodies offers compelling

support that our modified STXs exclusively target NaV proteins.

NaV expression and distribution in NGF-differentiated PC12 5.3.5

The cellular distribution of NaVs in live cells has been explored indirectly

through voltage-clamp measurements, which show an enhancement of Na+ current

density in excised neurites relative to the cell body of intact cells.71 Immunocytology

experiments to mark the location of NaVs in NGF-differentiated PC12 cells appear in a

single report.25 These data are obtained with a specific antibody raised against NGF-

induced NaV1.7 protein and reveal enhanced channel populations in neurite growth

cones and in the membrane of the cell soma (reprinted in Figure 5-16 C). Consistent

with this data, confocal images recorded with STX-Cy5 and STX-DCDHF against

NGF-treated PC12 cells highlight specific labeling of NaVs at neuritic processes and in

the somatic membrane (Figure 5-16 F). Regions of enhanced fluorescence intensity

are focused at the growth cones and in the periphery of the cell body. Interestingly, in

both fixed and live cells, membrane labeling (i.e., NaV distribution) is non-uniform.

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Figure 5-18 Quantification of probe wash off and PC12 ± NGF fluorescence.

Media contained 10 nM STX-Cy5 or 15 nM STX-DCDHF. Relative fluorescence of

undifferentiated and NGF differentiated PC12 cells.1

To query the relation between overall surface NaV expression and NGF

treatment, we compared the integrated fluorescence intensity of STX-Cy5 or STX-

DCDHF in samples of identical PC12 cells incubated with and without NGF.

Undifferentiated cells exposed to 10 or 15 nM STX-Cy5 or STX-DCDHF gave

average fluorescent intensities of 25% relative to NGF-treated cells (Figure 5-18),

further evidence of the influence of NGF to upregulate surface expression of NaV

(Figure 5-16 D–E). The detection of a small population of NaVs in undifferentiated

cells, which show no measurable Na+ ion current in electrophysiology recordings,

suggests that a basal level of non-functional NaVs is maintained in the cytoplasmic

membrane at all times, primed for activation by NGF-promoted pathways.72 The

ability to detect non-functional NaVs in live cells is compelling and could have far-

reaching consequences with regards to our understanding of how posttranslational

protein modification is used to regulate neuronal cellular plasticity.

Single-molecule and super-resolution imaging 5.3.6

DCDHF and Cy5 dyes were selected for their utility imaging experiments.60, 73,

74 Images of membrane-localized NaVs are obtained by adding STX-Cy5 or STX-

DCDHF to imaging media at concentrations well below the IC50 values of either

probe (12.8nM for STX-DCDHF and 10nM for STX-Cy5). In the single-molecule

measurements, by carefully selecting the exposure times used, even with an unbound

fraction of fluorescent STX still in solution, single-molecule signals of NaVs as labeled

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by fluorescent STXs are clearly visible above the background emission from the

solution (S/B 1.8 ± 0.4 for STX-Cy5, and 4.2±0.5 for STX-DCDHF). We are able to

image single molecules of NaV-bound STX-Cy5 by carefully selecting the intensity

and exposure time, such that free STX-Cy5 would only contribute to background. On

the other hand, the favorable signal-to-background of DCDHF stems from its ability to

brighten when interacting with the more constrained NaV local environment compared

to the aqueous media outside the cell60 (Figure 5-10). The ability to maintain a fixed

concentration of STX-Cy5 or STX-DCDHF in the medium enables new binding

events to occur during imaging. Additionally, experiments performed over the course

of days in media containing STX-Cy5 or STX-DCDHF show no appreciable effect on

cell morphology or viability, demonstrating the utility of these probes for long-term

imaging applications.

On the cellular membrane, STX-Cy5 exhibits a short visibility time of

100±10ms (2 exposure frames, Figure 5-20 I) on average. This short visibility time

combined with binding of new STX-Cy5 from solution to NaV contributes to steady

on/off single-molecule signal required for super-resolution imaging. This process can

be regarded as a target-specific variation of the PAINT method of super-resolution

imaging.75 Further, STX-Cy5 and STX-DCDHF require no use of exogeneous thiol

reductants or photoactivation. The ability to fine-tune the photophysical properties of

STX-fluorophores through judicious choice of dye molecule and selective coupling

with STX–NH3+ is testament to the versatility of our preparative scheme for

assembling these imaging agents.

Single-molecule tracking of labeled NaVs in NGF-differentiated PC12 5.3.7

Single-molecule tracks of fluorescently labeled proteins provide the potential

for quantitative analysis of the mechanisms by which such proteins partition to distinct

subcellular regions.76 Figure 5-19 B show single epi-fluorescence imaging frames

where single-molecule spots are clearly visible on the cellular soma and an axon. Over

the cellular soma, both singles and clusters of NaVs are observed. Several single

molecules from Figure 5-19 B are tracked with time to yield illustrative tracks of the

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motion shown in Figure 5-19 D, which exhibit characteristics reminiscent of confined

diffusion, Brownian motion, and directed transport. In principle, such data can

differentiate among physical processes associated with NaV behavior, including

membrane diffusion, cytoskeleton-associated transport, and endocytotic turnover.77

Tracking experiments may also distinguish whether sites of NaV accumulation in live

cells comprise proteins that reside permanently in a distinct state or represent dynamic

transitions of a single population – information unavailable through ensemble

imaging.73, 78 Critical to tracking analysis is the availability of fluorescent markers that

preserve the motility of the protein of interest, thus favoring fluorescent labeling

methods that target endogenous proteins over fluorescent fusion constructs.

Figure 5-19 Single-particle tracking of NaVs in the soma and neurites of an NGF

differentiated PC12 cell. NaVs were labeled with 13 nM STX-DCDHF. (A)

Transmission image. (B) Single 50 ms frame showing individually resolved NaVs in

the soma and neurites. (C) Distribution function for a population of NaVs undergoing

Brownian diffusion (solid red line, Davg = 0.036 ± 0.027 um2/s, n = 370) superposed

on a histogram of measured diffusion coefficients of NaVs in the soma membrane of

the cell in A. The observed deviations evidence the heterogeneous motion of NaVs in

this region. (D) Representative single-molecule tracks on the soma of the cell in A.

(E) Track of a single NaV in a neurite comprising 10 time steps at 50 ms per step (blue

line) superposed on a single frame image containing the tracked molecule (green

arrow). These trajectories yield instantaneous velocities of 680–970 ± 10 m/s (n =5).1

While exhaustive analysis of the modes of motion of NaV is beyond the scope

of this work, ~ 4000 single NaV, found in the somatic plasma membrane of NGF-

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differentiated PC12 cells treated with 12.8 nM STX-DCDHF, were tracked using low

imaging intensity to reduce photobleaching for the purposes of a preliminary

characterization. Within the somatic membrane, STX-DCDHF labeled NaVs are

observed to undergo a variety of trajectories before disappearance and exhibit an

average visibility time of 250±7ms. Of these NaV trajectories, 370 trajectories with 10

steps or more were selected to compute short-time diffusion coefficients from mean

square displacements (Figure 5-19 E). We compared the measured D distribution with

the hypothesis that all measured values arise from underlying random Brownian

motion characterized by a single diffusion coefficient, Do, which yields a well-known

probability distribution56 plotted by the smooth red curve (Figure 5-19 C), with the

average diffusion coefficient, Davg= 0.036±0.027µm2/s of the 370 molecules as Do.

Departures from this distribution indicate heterogeneous motion, as is observed here

(Figure 5-19 C). Reported values of NaV diffusion coefficient in subsomatic

environments such as axon initial segment (AIS) and dendrites4240 have been in a

similar range. The heterogeneity in NaV motion as shown in the distribution may

reflect differences in association with cytoskeletal elements, regulatory proteins,

and/or glycolipids in the respective membrane regions. Futhermore, fluorescent STX

also enables direct visualization of NaV in PC12 neurites, such as shown in Figure 5-

19 E. The trajectory of the indicated single NaV is plotted at 100ms temporal

resolution (closed circles), which translates to a velocity of 880±10nm/s (75mm/day)

through the length of the axon. This value is consistent with previous velocities of

axonal transport of vesicular structures, synaptic vesicles, neuropeptides, transmitters

and associated enzymes of between 8mm-400mm/day.79 These preliminary results

establish the utility of STX-fluorophores for querying the differential motility of

endogenous NaVs.

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Single-molecule imaging of NaV distributions on filopodia 5.3.8

Figure 5-20 Single-molecule imaging of NaVs in filopodia of an NGF differentiated

PC12 cell. NaVs were labeled with 3 nM STX-Cy5. (A) Diffraction-limited image

averaged over 50 s. (B) Transmission image of A. (C) Single 50 ms frame with

labeled NaVs resolved as single molecules. (D) A 50 s fluorescence maxplot of the

cell in A at t = 0 highlighting the base of an extending filopodium (blue box). (E) A

50 s fluorescence maxplot of the cell in A at t = 820 s showing elongation of the

highlighted filopodium. (F) Magnified image of the boxed area in D. (G) Magnified

image of the boxed area in E. (H) Length of the filopodium highlighted in D–G

plotted as a function of time, showing rapid extensions and retractions. (I) Visibility

time of STX-Cy5 on the membrane of the cell in A (average Tvis = 100 ± 10 ms).1

Epifluorescence images also reveal clear presence of NaV channels in filopodia

(Figure 5-20 C) in differentiated PC12 cells exposed to 3nM STX-Cy5. Contrasting

with STX-DCDHF, the visibility time of STX-Cy5 is smaller, approximately

100±10ms (Figure 5-20 I), allowing for rapid stochastic sampling of single NaV

locations. By maxplotting over 50s, a method by which the maximum intensity value

of an individual pixel over 50s is taken to be the value of that pixel, the locations of

217

individual NaVs over time are summed into a single image (Figure 5-20 D-E, Figure 5-

21 A). In this manner, by using the NaV distribution to outline cellular filopodia,

growth cone motility can be observed at 10s time resolution. Through maxplotting

analysis over 24 minutes, we are able to see cellular filopodia extend and retract

(Figure 5-20 D-G, Figure 5-21 A). Filopodia length is defined as the distance between

the terminus of soma and the tip of filopodia. For frames summed over 10s, the soma

and filopodia termini positions are measured by hand. A triplicate of the positions are

measured and error bars in position come from 2× the standard deviation in those

position measurements.

Filopodia stretch rate is determined by the absolute difference in filopodia

lengths through time divided by the time elapsed (10s). This “stretch” measurement

includes both extension and retraction. The maximum “stretch” rate observed over the

length of 990 seconds is 360nm/s, with an average stretch rate of 80±90nm/s. These

values are consistent with previous reports of 60-400nm/s found in DRG, and

Hippocampal filopodia projections.80 Given that non-continuous and rapid extension

and contraction in this range are likely due to sliding of actin filaments,81 our data

implicate association of NaVs with the cytoskeleton in filopodia. See Figure 5-21 b for

more information.

Figure 5-21 Maxplot images of NaVs labeled with STX-Cy5 and stretch rates in

filopodia of an NGF differentiated PC12 cell. (a) Time lapse sequence of maxplot

images. Each image is a maxplot average of data stacks acquired over 50 s. (b)

Stretch rates of filopodia in (a) over 990 s showing a maximum stretch rate of 360

nm/s and an average rate of 80 ± 90 nm/s.1

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Super-resolution imaging of labeled NaVs in filopodia and neuritic spines 5.3.9

The small dimensions of neuritic spines in NGF-differentiated PC12 cells

render them intractable for detailed study by conventional diffraction-limited (DL)

confocal or epifluorescence microscopy (Figure 5-22 c,e,k). Super-resolution (SR)

imaging of individual proteins in neuritic substructures thus has the potential to reveal

intricate details of these cellular features, information that may give deeper insight into

molecular mechanisms of channel behavior. By harnessing the short visibility time of

STX-Cy5, we record multiple single-molecule positions which yield the rapid

pointillist imaging required for super-resolution reconstruction with a mean statistical

localization precision of 30 nm (Figure 5-8 A).

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Figure 5-22 Super-resolution imaging of NaVs in neuritic spines of an NGF

differentiated PC12 cell. NaVs were labeled with 10 nM STX-Cy5. (a) Super-

resolution image acquired over 5 s at a 50 ms frame rate. (b) Diffraction-limited

image of A. (c) Diffraction-limited image of the blue boxed area in A. (d) Super-

resolution image of the blue boxed area in A. (e) Diffraction-limited image of the

green boxed area in A. (f) Super-resolution image of the green boxed area in A. (g)

Measured full width at half maximum (FWHM) values for the neuritic spine indicated

by the blue arrow in C and D from diffraction-limited (400 nm, blue line) and super-

resolution (81 nm, red line) images. (h) Super-resolution image of A 25 s later,

acquired over 25 s at a 50 ms frame rate. (i) Measured FWHM value for the super-

resolved neuritic spine indicated by the blue arrow in H and J. (j) Super-resolution

SR

220

image of the yellow boxed area in h. (k) Diffraction-limited image of the yellow

boxed area in h.1

We find that 5s imaging time is needed to accumulate enough localizations for

a well filled-out axonal structure in a ~ 450µm2 region of interest containing the

imaged axon (Figure 5-22 a). Since each single-molecule localization event is

reconstructed equally, our reconstructions also preserve the density of STX-NaV

binding events in different axonal regions. In the SR reconstruction, punctate

populations of NaVs are observed at nodes along the neurite shaft and at the origins of

numerous projections extending from the axon body. Neuritic spines, approximately

85 nm in width (Figure 5-22 a, d, h, j), are perceptible as discrete structures containing

NaVs distributed both individually and in small clusters. Averaging 5300±200

localizations per second, a quasi-real-time SR movie with STX-Cy5 on differentiated

PC12 cells reveals that these neuritic outgrowths exhibit highly fluxional lateral

movements, extensions, and retractions on the timescale of hundreds of milliseconds

(shown as a sequence of images in Figure 5-22 l), on par with neuritic spine motility in

neurons.82 These images, captured in quasi-real-time by sliding boxcar summations of

positions at an unprecedented level of resolution, are uniquely enabled by STX-based

fluorescent dyes. To the best of our knowledge, these data represent the first targeted

super-resolution images of wild-type living animal cells using reversibly binding,

small molecule probes.

Conclusion 5.4

Voltage-gated sodium ion channels constitute the fundamental units of ion

conduction in action potential generation and propagation in electrically excitable

cells. An intricate array of regulatory networks and dynamic partitioning mechanisms

control the concentrations and spatial distributions of NaV subtypes and shape the

specific conductive properties of a cell. Misregulation of NaV trafficking and/or

processing leads to aberrant action potential firing patterns and is associated with

profound neurological disease. Despite the importance of NaVs in neuronal function,

methods for live cell imaging of these proteins are conspicuously few. Saxitoxin

221

offers a unique molecular template for the design of new pharmacological and

imaging agents to interrogate dynamic channel function. Chemical synthesis has

made possible access to fluorescently labeled forms of STX that maintain high

efficacy and reversible binding to the channel. We have described the

electrophysiological characterization of these novel toxin conjugates and have

demonstrated their utility for live-cell imaging of NaVs in NGF-differentiated PC12

cells in confocal, single-molecule, and super-resolution microscopy experiments. Our

study demonstrates that STX-Cy5 and STX-DCDHF can detect functionally silent

NaVs in undifferentiated PC12 cells, a finding that may enable future investigations to

understand protein regulation of NaV activity. The unique and favorable properties of

these probes have been underscored in super-resolution imaging experiments, which

provide unparalleled spatial and temporal resolution of NaV motility in functionally

distinct regions of the excitable cell membrane including direct visualization of NaVs

in live neuritic spines.

Acknowledgement 5.5

I thank Prof. Robert J. Twieg and Jarrod C. Williams for the gift of the

DCDHF-NHS molecule. I am grateful to Professor Merritt Maduke for allowing use

of her electrophysiology equipment. This work was supported in part by Grant No.

R01-GM086196 (W.E.M.) and Grant No. R01-NS45684 (J.D.) from the National

Institute of General Medical Sciences and the National Institute of Neurological

Disorders and Stroke, respectively, and by a grant from the Tobacco-Related Disease

Research Program (J.D.).

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