the impact of ros scavenging on nmda and ampa receptor … · 2013-11-22 · i would like to thank...
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The impact of ROS scavenging on NMDA and AMPA receptor whole cell currents in pyramidal neurons of the
anoxia tolerant western painted turtle
by
David Dukoff
A thesis submitted in conformity with the requirements for the degree of Master of Science
Cell & Systems Biology University of Toronto
© Copyright by David Dukoff (2013)
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The impact of ROS scavenging on NMDA and AMPA receptor
whole cell currents in pyramidal neurons of the anoxia tolerant
western painted turtle
David Dukoff
Master of Science
Cell and Systems Biology
University of Toronto
2013
ABSTRACT
Extended periods of oxygen deprivation cause brain death in mammals but the western painted
turtle overwinters in anoxic mud for months without damage. Neural protection is achieved
through decreases in the whole cell currents of N-methyl-D-aspartate and alpha-amino-3-
hydroxy-5-methyl-4-isoxazolepropionic acid receptors (NMDAR and AMPAR) that are
dependent on a mild increase in intracellular calcium from the mitochondria. The goal of this
research was to determine if natural anoxic decreases in reactive oxidative species (ROS) serve
as the signal to bring about these changes. Reductions in cellular ROS levels were demonstrated
to have no effect on AMPAR currents or intracellular calcium and produced massive increases in
NMDAR currents, indicating that ROS depression does not directly mediate anoxic alterations.
Interestingly, mammalian neural tissue also experiences a similar increase in NMDAR whole cell
current in response to reducing agents suggesting a possible conserved mechanism for normoxic
receptor control.
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ACKNOWLEDGMENTS
“Complete and finished.” That is what I have been told that the writing and submission of
a master’s thesis feels like. Within a hundred or so pages students summarize two years of
laboratory work, complete the requirements of their degree and move to the next phase of their
careers. However, as I approach the end of this project, this feeling of closure has yet to sink in.
During my first year living at the University of Toronto’s Graduate Residence, I was told that the
reason so much of the building seemed raw and unfinished was due to the architect’s belief that
“a graduate student’s work is never done”. Although I came to believe that this explanation was
an excuse for ill-placed support struts and walls so thin you could hear your roommates shift
their blankets, the phrase stuck a chord. Science is a potent addiction and no matter where my
life goes after my degree I don’t think I will ever stop thinking like a researcher or cease
studying the world around me. What is more, the people that have helped and supported me
during my time here are similarly addicting. This section is for all of them.
I would like to begin by thanking the professors and administrative staff whose efforts
have made my project and degree possible. I would like to thank Dr. Les Buck for everything he
has to done to support me during my time in the lab and make research fun. Aside from being a
constant presence to help me with nearly every aspect of my project, he has also been there to
trade jokes and offer advice on all manner of non-science related issues. He has kept me on track
but never been afraid to let me make my own mistakes (of which there have been many) so that I
can learn how to recover and be a better scientist. I would also like to thank my committee
members for getting me this far. The advice provided by Dr. Martin Wojtowicz and Dr. David
Lovejoy has helped shape the directions and aims of my experiments. More importantly, their
questions and comments have kept me versed in information outside the scope of my project and
reminded me of the value of the big picture. Finally, I would thank the administrative staff who
make this department work and who put up with the strange creatures that are graduate students.
A special thanks to Ian Buglass, Tamar Mamourian, Peggy Salmon, James Dix, and Trung Luu.
I would next like to thank my fellow students in our incredible department for the help
and support they have shown me over the last two years. I would like to first thank David Hogg
who, through unstinting assistance, opinionated discussions (often involving words I can’t
include in a thesis) and stories about his family, has kept me going and reminded me that there is
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more to life than what is under the microscope. Next I would like to thank Peter Hawrysh for
being a seemingly infinite resource of technical advice, scientific explanations, dirty stories,
nerdy references, awesome video games, and cold beer (not usually during work hours). I would
like to thank Aaron Chowdhury for his electronic expertise, great sense of humor, inquisitive
questions, and devious pranks that have kept me sharp, motivated, and protective of my laptop. I
would like to thank Hilary Bond for her stories and comments which have helped shape my
decisions about who I want to be. I would like to thank Lydia Farnell for giving me someone to
trade country stories and country music with. I would also like to thank all of my fellow
colleagues in the department whose friendships have kept me sane. A special thanks to Jessica
Pressey, Louise de Lannoy, Zoltan Torontali, Dave Wasserman, Sabina Romanescu, Felix
Gunawan and all the members of CSBGU. I would not have made it without all of you.
I would like to thank my friends and family outside the lab for always being there. My
parents have provided unconditional love and support for every choice I have made. Through
visits to take me out to lunch and time spent editing my writing, they have helped me in any way
possible. I hope I have made them proud. I would like to thank my little brother Andrew for
sending me music on lonely nights at the lab. I would like to thank David Darling and my
godchild Kaylie for Sudbury vacations that recharged my batteries. Finally, I would like to thank
my Toronto/Kitchener friends for creating occasions to get out of the lab and into the sunlight.
Special thanks to Brittany, Serge, Robert and Vanessa for their efforts in this regard.
The last acknowledgement I would like to make is to science and nature (not to be
confused with the professional journals). I went into biology for the same reason I spent my
childhood taking things apart in my Dad’s workshop: I wanted to understand how things work.
There are no greater puzzles or more complex mechanisms than those found in the natural world
and this may be why I find myself, like so many others, completely obsessed with studying it.
The famous biologist Lewis Thomas once wrote about scientific research, “I don't know of any
other human occupation, even including what I have seen of art, in which the people engaged in
it are so caught up, so totally pre-occupied, so driven beyond their strength and resources…
There is nothing to touch the spectacle.” Thank you science for being so interesting.
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TABLE OF CONTENTS
ABSTRACT………………………………………………………………………………………ii
ACKNOWLEDGEMENTS……………………………………………….……………………iii
TABLE OF CONTENTS…………………..……………………………………………………v
LIST OF FIGURES……………………………………………………………………………..ix
ABBREVIATIONS……………………………………………………………………………...xi
CHAPTER 1: INTRODUCTION………..……………………………………………………..1
1.1 The importance of oxygen and low oxygen tolerance………………………………...1
1.2 Ice ages, glacial periods and their contributions to modern day biodiversity…………2
1.3 Life after glaciation and adaptations to low oxygen…………………………………..3
1.3.1 Chrysemys picta belli………………………………………………………..4
1.3.2 Spermophilus parryii parryii………………………………...……………...6
1.3.3 Carassius carassius…………………………………………..……………..8
1.4 Requirements for anoxia/hypoxia tolerance…………………………………………..9
1.5 The importance of neuronal down regulation………………………………………..10
1.6 Models for potential human application……………………………………………..11
1.7 Turtle synaptic organization………………………………………………………….12
1.8 Anoxia mediated neuroprotective mechanisms……………………………………...13
1.8.1 Spike arrest…………………………………………………………………13
1.8.2 Channel arrest……………………………………………………………...14
1.9 Mechanisms controlling “channel arrest”……………………………………………16
1.9.1 Increases in adenosine……………………………………………………...16
1.9.2 Activation of ATP-sensitive K+ (KATP) channels………………………..17
1.9.3 Increases in intracellular calcium…………………………………………..19
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1.9.4 Enhancements of calmodulin and phosphatase activity……………………19
1.10 Reactive oxidative species (ROS)…………………………………………………..20
1.11 The role of ROS in anoxia-intolerant vertebrates…………………………………..22
1.11.1 Redox changes affect NMDAR but not AMPAR activity………………..22
1.11.2 Alterations in ROS affect NMDAR activity……………………………...23
1.11.3 NMDAR redox regulation is blocked by kinase inhibition………………24
1.11.4 mKATP mediated ROS increases affect ischemic preconditioning……...26
1.12 The antioxidant defence system…………………………………………………….27
1.13 ROS regulation of GABA in anoxic turtle cortex…………………………………..28
1.14 Rationale for study………………………………………………………………….29
CHAPTER 2: MATERIALS AND METHODS……………………………………………...30
2.1 Animals………………………………………………………………………………31
2.2 Dissection and whole-cell patch-clamp recording protocol………………………….30
2.3 Neuron identification………………………………………………………………...34
2.4 Evoked NMDAR and AMPAR current amplitude recordings………………………34
2.5 Fluorescence investigation of cytosolic Ca2+ changes……………………………….36
2.6 Whole cell spontaneous neuronal activity recordings………………………………..38
2.7 Pharmacology and drug application………………………………………………….39
2.8 Measurement of oxygen tension……………………………………………………..40
2.9 Chemicals…………………………………………………………………………….41
2.10 Statistical analysis…………………………………………………………………..41
CHAPTER 3: RESULTS………...…………………………………………………………….42
3.1 Pre-current recording evaluations……………………………………………………42
3.1.1 Anoxic conditions are produced within the bathing chamber……………...42
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3.1.2 All recordings were conducted within pyramidal cells…………………….43
3.2 Whole-cell NMDAR current experiments…………………………………………...44
3.2.1 Normoxic NMDAR currents are stable throughout the time course………44
3.2.2 Normoxic NMDAR current amplitudes are reduced during anoxia……….45
3.2.3 NMDAR current amplitudes are increased by ROS scavenging…………..47
3.2.4 NMDAR current amplitudes are decreased by H2O2 addition……………..49
3.2.5 Blocking mitochondrial ROS production increases NMDAR currents……49
3.2.6 Anoxia attenuated NMDAR currents are not reversed by ROS addition….51
3.3 Whole-cell AMPAR current experiments……………………………………………53
3.3.1 Normoxic AMPAR currents are stable throughout the time course……….53
3.2.2 Normoxic AMPAR current amplitudes are reduced during anoxia………..53
3.3.3 AMPAR current amplitudes are unaffected by ROS scavenging………….55
3.3.4 AMPAR current amplitudes are unaffected by H2O2 addition…………….55
3.4 Oregon green intracellular calcium assessments…………………………………….57
3.4.1 Excitation scans confirmed dye loading in turtle brain sheets……………..57
3.4.2 Normoxic calcium concentrations are stable throughout the time course…57
3.4.3 Intracellular calcium concentrations are increased by anoxia……………..58
3.4.4 Intracellular calcium concentrations are unaffected by ROS scavenging….58
3.4.5 Intracellular calcium concentrations are unaffected by H2O2 addition…….59
3.5 Action potential frequency assessments in spontaneously active pyramidal cells…..62
3.5.1 Action potential frequency is reduced by ROS scavenging………………..62
CHAPTER 4: DISCUSSION…………………………………………...……………………...64
4.1 Control normoxic and anoxic NMDA and AMPA currents…………………………64
4.2 Pharmacological manipulation of ROS………………………………………………64
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4.2.1 The NMDAR modulatory redox site in turtle cortical slices………………64
4.2.2 Mitochondrial produced ROS in NMDAR redox modulation……………..67
4.2.3 A potential mechanism for ROS mediated NMDAR control……………...69
4.2.4 AMPAR redox insensitivity………………………………………………..74
4.2.5 Anoxia-mediated block of NMDAR redox control………………………..75
4.3 A role for anoxia-mediated calcium increases in blocking redox control…………...76
4.4 Roles for redox sensitive GABA release…………………………………………….79
4.5 Potential mechanism of NMDAR/AMPAR control under anoxic conditions……….80
4.6 Concluding remarks and directions for future research……………………………...83
REFERENCES………………………………………………………………………………….85
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LIST OF FIGURES
CHAPTER 1: INTRODUCTION
Figure 1.1: The ranges of the four currently recognized subspecies of Chrysemys picta……...…6
Figure 1.2: NMDAR and AMPAR structures……………………………………………………16
Figure 1.3: Schematic illustration of anoxia-mediated changes contributing to channel arrest…20
Figure 1.4: The mechanisms of ROS production and cysteine oxidation………………………..25
CHAPTER 2: MATERIALS AND METHODS
Figure 2.1: Diagrammatic representation of cortical sheet excision and experimental setup…...33
Figure 2.1: NMDAR and AMPAR whole-cell patch clamp recording timeline………………...35
Figure 2.2: Oregon Green fluorescent recording timeline……………………………………….38
Figure 2.3: Whole cell spontaneous activity recording timeline………………………………...39
CHAPTER 3: RESULTS
Figure 3.1: Change in oxygen tension during normoxic to anoxic transition……………………43
Figure 3.2: Spike frequency adaptation in pyramidal and stellate neurons……………………...44
Figure 3.3: Effect of anoxia on whole cell NMDAR currents…………...………….…………...46
Figure 3.4: Effect of ROS scavenging on whole cell NMDAR currents………………………...48
Figure 3.5: Effect of H2O2 manipulation on whole cell NMDAR currents……………………...50
Figure 3.6: Effect of H2O2 addition on anoxia-attenuated whole cell NMDAR currents………52
Figure 3.7: Effect of anoxia on whole cell AMPAR currents……………………………………54
Figure 3.8: Effect of ROS manipulation on whole cell AMPAR currents………………………56
Figure 3.9: Oregon Green excitation scans………………………………………………………57
Figure 3.10: Effect of oxygen and ROS manipulation on Oregon Green fluorescence……...….60
Figure 3.11: Sample Oregon Green fluorescence traces…………………………………………61
Figure 3.12: Changes in spontaneous activity in response to ROS scavenging…………………63
CHAPTER 4: DISCUSSION
x
Figure 4.1: Diagrammatic representation of the proposed mechanism for ROS mediated
NMDAR control under normoxic conditions……………………………………………………73
Figure 4.1: Diagrammatic representation of the proposed mechanism for anoxia mediated
NMDAR/AMPAR downregulation……………………………………………………………...82
xi
ABBREVIATIONS
C degrees Celsius
Ψm mitochondrial membrane potential
[Ca2+]i intracellular calcium concentration
8-PT 8-pentyltheophyilline
5-HD 5-hydroxydecanoic acid
12h:12h twelve hours of light and twelve hours of dark light cycle
aCSF artificial cerebrospinal fluid
AFU arbitrary fluorescence unit
AMP adenosine monophosphate
AMPA α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid
AMPAR α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor
AP action potential
APf action potential frequency
ATP adenosine triphosphate
BAPTA 1,2-bis(o-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid
BDM butanedione-monoxime
Ca2+ calcium ion
CaCl2 calcium chloride
CaM calmodulin
cGMP cyclic guanosine monophosphate
Cl- chloride ion
CO2 carbon dioxide
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CM-H2DCFDA 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester
C.p. Chrysemys picta
Cys cysteine
DTNB 5,5'-dithiobis-(2-nitrobenzoic acid)
DTT dithiothreitol
EPSP excitatory postsynaptic potential
EEG electroencephalogram
ER endoplasmic reticulum
Fig Figure
Fmax maximal fluorescence signal
Fmin background fluorescence
GΩ gigaohm
GABA gamma-Aminobutyric acid
GPx glutathione peroxidase
GSH glutathione
GSSG glutathione disulfide
H+ proton
H2O2 hydrogen peroxide
HCO3- bicarbonate
IMM inner mitochondrial membrane
I/V current/voltage
IV intravenous
K+ potassium ion
KATP channel ATP-sensitive potassium channel
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KCl potassium chloride
Kd dissociation constant
kDA kilo dalton
kHz kilohertz
L litre
MΩ megaohm
MCU mitochondrial calcium uniporter
Mg2+ magnesium ion
MgCl2 magnesium chloride
min minutes
mKATP channel mitochondrial ATP-sensitive potassium channel
mM millimolar
mmHg millimeters of mercury
Mn2+ manganese ion
MnSOD manganese superoxide dismutase
mOsM milliosmole
MPG N-2-mercaptopropionylglycine
mPTP mitochondrial permeability transition pore
ms millisecond
mV millivolt
n sample size
N2 nitrogen gas
Na+ sodium ion
NaCl sodium chloride
xiv
NAC n-acetylcysteine
NaGTP sodium guanosine triphosphate
NaATP sodium adenosine triphosphate
NaH2PO4 sodium dihydrogen phosphate
NaHCO3 sodium bicarbonate
Na-HEPES sodium 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
nm nanometre
nM nanomolar
NMDA N-Methyl-D-Aspartate
NMDAR N-Methyl-D-Aspartate receptor
nNOS nitric oxide synthase
NO nitric oxide
O2 molecular oxygen
O2 superoxide anion
ONOO peroxynitrite
pKATP channel plasmalemmal ATP-sensitive potassium channel
PP protein phosphatase
PP2B calcineurin
PSD95 postsynaptic density protein-95
PSC postsynaptic current
Q10 temperature coefficient
ROS reactive oxygen species
[ROS]i intracellular ROS concentration
s seconds
xv
S. Spermophilus
SA spike arrest
S.E.M. standard error of the mean
sGC soluble guanylate cyclase
SOD superoxide dismutase
S.p. Spermophilus parryii
t time
TTX tetrodotoxin
μM micromolar
WTR water transport route
X/XO xanthine/xanthine oxidase
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CHAPTER 1: INTRODUCTION
1.1 The importance of oxygen and low oxygen tolerance
Humans and most other vertebrates require a constant supply of oxygen to survive. Brief
hypoxic exposure causes hyperventilation and tachycardia that temporarily compensate for
decreased systemic oxygen but sustained anoxia for more than a few minutes results in
neurological damage and eventually death [1, 2]. Aerobic life primarily depends on oxidative
phosphorylation for the production of the cellular energy substrate - adenosine triphosphate
(ATP), and if oxygen transport to tissues becomes compromised energy demands cannot be met
and cellular death occurs [3, 4]. Despite these limitations, some vertebrates have developed the
ability to survive under hypoxic or anoxic conditions for extended periods of time.
Understanding the mechanisms through which tolerance is accomplished has applications within
fields as broad as stroke prevention to space travel [5, 6].
Research has been conducted for decades on hypoxia and anoxia tolerant species to try
and better understand their physiological and molecular adaptations. Running in parallel with
this work has been research to try and understand the conditions that produced these adaptations.
Related species to those with anoxia/hypoxia tolerance rarely possess the same capabilities,
indicating a historical pressure that selected for the trait and brought about a speciation event.
Understanding why some species have adapted to survive long term anoxia may help us better
understand how they do it.
Within the northern latitudes a number of hypoxia tolerant mammals and anoxia tolerant
reptiles and fish are found in environments that demonstrate seasonal temperature decreases and
the development of ice cover. In order to survive these seasonal changes species have adapted
various hibernation and overwintering strategies that involve tolerating extended periods of
anoxia/hypoxia. It has been proposed that these survival strategies were selected for during
periods of globally decreased temperature and increased ice cover. The most recent such event
was the last glacial period.
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1.2 Ice ages, glacial periods and their contributions to modern day biodiversity
When extensive ice sheets are present in the northern and southern hemispheres the
period is designated as an ice age. Our planet has passed through four major ice ages since its
formation and is currently experiencing the fifth that began 2.6 million years ago marking the
beginning of the Quaternary Period [7]. Environmental conditions during ice ages alternate over
tens of thousands of years between two opposing conditions. “Interglacial” periods are
characterized by warmer global temperatures and glacial recession. “Glacial” periods are marked
by colder global temperatures and ice sheet advances. Movement between these two periods is
brought on by large scale changes in the amount of thermal energy present in the Earth’s
atmosphere and oceans caused by alterations in the energy generated by the sun, changes in the
distance of the Earth’s orbit, and adjustments in atmospheric composition. We are currently
experiencing an interglacial period that began 11,400 years ago [7, 8].
The last glacial period began 120,000 years ago and at that time the regional temperatures
and climates were nearly identical to those seen today. Glaciers expanded across the globe
reaching a maximum between 26,500 and 20,000 years ago when glaciers and permafrost
covered more than a quarter of the Earth’s surface [7, 8]. Approximately 28,000 years ago the
surface temperatures of the oceans began to warm leading to slow rises in ocean levels and by
12,000 years ago most regions returned to climates close to what they had been seen previously.
This marked the transition from glacial to interglacial [7, 8].
Worldwide biodiversity was not significantly affected by the initial interglacial-glacial
transition or the sustained 100,000 yearlong glacial period itself. As ice sheets expanded most
species were able to migrate to areas not covered by ice, commonly referred to as refugia. These
areas were found sparsely throughout glaciated regions but primarily existed closer to the
equator [9]. Changing climate and the addition of new plant and animal life caused these regions
to begin to mimic the environments that had previously been found further north and south and
as a result, migrating animals did not face extensive pressures to adapt or change [7]. Most
species were forced into a third of the territory they occupied pre-glaciation, but mild population
decreases and the establishment of new territorial limits prevented vast losses of biodiversity [7].
Increased glaciation decreased sea levels by 350 feet generating land bridges like from Alaska to
3
Siberia that allowed for migrations between refugia, further preserving diversity [7, 10].
However as the glacial cycle came to an end, a major global extinction took place.
This extinction event took place between 16,000 and 10,000 years ago, and unlike
previous major global extinction events took place only on land. Nearly all large terrestrial
mammals over 44 kg were wiped out [7, 10]. Similar extinctions did not occur during exit from
previous glacial periods and the cause remains contested. The most supported hypotheses
involve rapid environmental changes that caused the loss of much of the world’s mega flora,
overhunting by early humans as they migrated into new regions following ice sheets recession,
or a combination of the two [7, 10]. Whatever the cause, all of the continents experienced losses
in large terrestrial animal life: South America (83%), North America (72%), Australia (88%),
Eurasia (35%), and Africa (21%) [10]. The mass extinction left a number of environmental
niches empty and provided surviving species with an opportunity to adapt and expand.
1.3 Life after glaciation and adaptations to low oxygen
About 10,000 years ago plant and animal life began to expand outwards from refugia as
the ice sheets receded [7, 9]. Some of these species/subspecies were new, produced from the
isolation of members of the same species in different refugia throughout the glacial cycle where
genetic drift and varying selective pressures caused speciation events [11, 12]. Expansion of
species from refugia following the last glacial cycle altered the biota from what existed pre-
glaciation to a greater degree than seen during exit from previous glacial cycles [13, 14]. The
new territory opened up by the mass extinction event was quickly colonized by smaller faster
breeding species leading to changes in plant and animal life. Adaptations were developed to
survive under these novel conditions increasing differences between new species/subspecies and
establishing new territorial ranges. These adaptations were generally minor and most
species/subspecies continued to live in similar habitats, experiencing only small changes in
physical features, ranges or behaviours [15, 16]. However, following glacial retreat some species
in North America and Europe expanded into regions that still experienced seasonal ice cover,
reduced temperatures and decreasing available food fostering the development of distinct
adaptations. Species with flight capabilities survived by developing adaptations to seasonally
migrate south. Other small and less mobile species adapted to outlast the winters. Three
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examples of such species are Chrysemys picta belli, Spermophilus parryii parryii, and Carassius
carassius.
1.3.1 Chrysemys picta belli
The North American turtle Chrysemys picta (C.p.) is the most widely distributed turtle
species in North America with a range extending from the Maritime Provinces west to British
Columbia and south to Oregon, Louisiana, and Georgia [17]. Fossils have been found for the
species indicating that they have been widely distributed for at least two, and possibly more than
five million years [17-19]. Previous to the last glacial period C.p. existed as two forms divided
along the Atlantic coastal and west of the Appalachian Mountains [20]. The expansion of glacial
ice and the loss of vegetation forced both forms southward and isolated the turtles into three
distinct populations that became distinct species with C.p. picta in the southeast Atlantic coast,
C.p. dorsalis in the lower Mississippi and C.p. belli in the southwest [21-23]. During glacial
recession all three of the C.p. species migrated out of refugia north and through interbreeding
between C.p. belli and C.p. dorsalis it is believed a hybridized fourth intermediated taxon C.p.
marginata came about [20]. Species eventually settled into their modern day locales with C.p.
picta in the Atlantic Coastal region, C.p. belli on the Great Plains, C.p. marginata around the
Great Lakes, and C. p. dorsalis within the southeastern United States [Figure 1.1] [20]. Recent
genetic investigation has proposed that interbreeding between species has begun to break down
the classic differentiations between groups and they may be more accurately regarded as a
northern subspecies (C. p. picta, C. p. belli, C.p. marginata) and a southern subspecies (C. p.
dorsalis) [24]. However within the scientific literature the classic separations still remain.
During migrations out of the refugia the northern turtle subspecies slowly adapted to
survive conditions of seasonal ice cover, decreased temperatures and reduced food availability
allowing for the expansion of their ranges [20]. They developed physiological and cellular
adaptations to survive the winter submerged in insulating mud at the bottoms of frozen fresh
water bodies in an unresponsive metabolically reduced state [25, 26]. Although the primary
function of these adaptations was probably freezing avoidance and energy conservation it also
produced capabilities for extreme anoxia tolerance as the overwintering environment required
turtles to survive for months without oxygen. Under laboratory conditions C.p. belli, picta, and
5
marginata at 3°C have all been demonstrated to survive over 120 days of anoxia, compared to
the southern subspecies (C.p. dorsalis) that does not overwinter and perishes after only 50 days
under identical experimental conditions [27-29]. Anoxia tolerance is greatest within members of
the subspecies found further north and the Western painted turtle (C.p. belli) that inhabits the
most northern ranges has been demonstrated to withstand the longest periods of anoxia [27, 30,
31].
The overwintering capabilities of C.p. belli are thought to have arisen from enhancements
of pre-existing diving abilities common to most turtle species [32, 33]. In comparative analysis
between C.p. dorsalis and C.p. belli it was concluded that increased tolerance is the result of
better capabilities to lower metabolic rate and limit lactate accumulation [27]. During
overwintering energy is conserved via decreases in brain activity, heart contraction, protein
synthesis, urea production and the maintenance of a comatose like state [34-38]. This leads to a
70-80% reduction in ATP turnover [34]. The remaining energy demands are met via glycolytic
fermentation supported by enhanced glycogen stores within the liver (15% of the organs mass)
[39]. The two units of lactate and free hydrogen ions produced for every unit of glucose
anaerobically consumed are buffered by a high plasma bicarbonate concentrations and ion
exchange systems between the shell and bones (comprising 37.5% of their total mass) [40].
Induction of the overwintering state occurs in response to low oxygen and does not appear to be
controlled seasonally like within mammal hibernators [41].
6
Figure 1.1: The ranges of the four currently recognized Chrysemys picta subspecies.
Figure adapted from online source [42].
1.3.2 Spermophilus parryii parryii
The arctic ground squirrel (Spermophilus parryii parryii) is the largest of the American
ground squirrel genus and the most northern dwelling of the squirrel family. They are found
throughout the Canadian Arctic, subsist socially in large burrows, and during the coldest 7 to 8
months of the year they hibernate in their dens at temperatures sometimes below 0C [43, 44].
S. parryii originally arose in Asia from a common ancestor to the central Siberian S.
undulatus approximately 100 000 years [45, 46]. The speciation event took place during the last
7
glaciation within a large refugium consisting of portions of eastern Siberia, Alaska, and the
western Yukon Territories connected by the Bering Land Bridge [8, 11, 47-51]. The newly
speciated S. parryii migrated to North America and further subdivided into a large arctic (S. p.
parryii) and subarctic subspecies (S. p. plesius). Following glacial recession the two subspecies
dispersed north into the regions they currently occupy today with S.p. parryii found north of the
Brooks Range in Alaska and arctic Canada and S.p. plesius found south in interior Alaska and
subarctic Canada [45, 46].
Like the western painted turtle the arctic ground squirrel survives the winter through
physiological and cellular changes that conserve energy and allow them to tolerate low
temperatures. During hibernation resting body temperature is reduced to between 2-10 °C, heart
rate decreases (25 fold), neurological protein synthesis is arrested (0.04% of active levels) and
blood flow to various brain regions is reduced (80-90%). These changes cumulatively result in
glucose utilization rates that are 1-2% of resting levels [3, 52-55]. The resting respiration rate is
also decreased (25 fold) and O2 consumption is reduced (90-96%) allowing the squirrels to
survive prolonged durations in burrows that can become hypoxic during the winter season [3,
56]. Metabolic needs are met through the breakdown of fat and lean muscle reserves. Increased
levels of steroids/androgens and lipases three weeks prior to hibernation allows the squirrels to
increase their mass by 42% (87% fat and 3% protein) ensuring an adequate fuel supply [57, 58].
Also like the western painted turtle, the adaptations utilized by the arctic ground squirrel
to survive the winter season are believed to have been produced through modifications of
existing capabilities common to ground squirrels as a whole. Several other grounds squirrel
species hibernate although for shorter periods at not as low temperatures [59]. Most ground
squirrel species also experience oxygen fluctuations within their burrows (6-20%) at different
points of the year [60-62].
Ground squirrels do not remain quiescent throughout the hibernation period and at
regular periods briefly warm and reverse depressions in brain activity and heart rate without
awakening. These arousal periods are part of the torpor arousal cycle and although energetically
costly, are thought to be necessary for the maintenance of neural tissue and to undergo sleep
cycles that cannot be accomplished during hibernation [63]. They also allow squirrels to filter
8
out and excrete metabolic waste products that accumulate during the inactive state. The torpor
arousal cycle and entrance/exit from the hibernation state are based on a circannual cycle and
persists under constant photoperiods with food provided ad libitum [43, 64-66].
1.3.3 Carassius carassius
The crucian carp (Carassius carassius) is widely distributed throughout northern Europe
with a range spanning from England to Russia [67]. The species lives within lakes and ponds
that for several months every winter become anoxic as the surface freezes over preventing
aquatic photosynthesis and atmospheric oxygen diffusion [68]. During this period only the
crucian carp survives, remaining active in water at 2°C, without food, for up to 5.5 months [69].
Previous to the last glacial period, the crucian carp was widely dispersed across Eurasia.
During glaciation, the species survived within bodies of water in southern and eastern refugia
and re-dispersed to central and north European waters following ice sheet recession [70].
Because the carp are aquatic organisms it is unclear if their current range or adaptations to
seasonal ice cover are the result of the post glacial extinction event or are products of previous
glacial periods.
The physiological adaptations that allow the carp to survive under the ice involve
decreases in metabolic rates and body temperature, similar to those seen in the species discussed
above. Body heat production is reduced by 37%, whole organism protein synthesis is down
down-regulated 50-95%, and heart rates is lowered from 88 to 10 beats per minute [71-73].
Although crucian carp remain active throughout the anoxic bout, they do experience a major
reduction in activity within optical centres of the brain (90% decrease) that brings about
blindness without deficits to survival [73, 74]. The cumulative down regulation of these
processes decrease metabolic energy requirements to levels that can be met through anaerobic
glycolysis. To accomplish this, the crucian carp also possess a large glycogen reserve
accounting for 30% of the liver’s wet mass and 4.5% of the fish’s weight that allows them to
survive anaerobically and without feeding through the winter [75]. In order to deal with lactate
accumulation, the carp convert lactate to ethanol and release it through their gills into the
surrounding environment [72]. Similar to the western painted turtle the crucian carp brings about
9
cellular and physiological changes in response to decreases in available oxygen and not as a
result of circannual rhythms [76].
Similar to the other species discussed above the crucian carp’s adaptations for winter
survival are believed to be the result of adjustments to pre-existing capabilities. The goldfish
(Carassius auratus) is a cousin of the crucian carp that is found farther south in regions
throughout Eastern Asia and sheltered within the same refugia during the last glacial period [70,
77]. In order to tolerate decreases in oxygen brought on by brief ice cover or algae blooms
goldfish have adapted to tolerate several weeks of hypoxia or hours of anoxia [78].
1.4 Requirements for anoxia/hypoxia tolerance
It has been stated that in order for prolonged anoxia/hypoxia tolerance to be
accomplished adaptations must exist that allow organisms to conserve energetic substrates and
prevent self-pollution by end-product accumulation [79]. All of the species discussed above
possess strategies to accomplish these goals, many of which are common between species.
Reduction of metabolic activities is the primary method through which both of these
goals are met, termed “metabolic arrest”. Down regulations of ATP consuming processes
decrease rates of substrate utilization and by-product production. During overwintering all of the
species discussed above experience depressions in heart rate/blood flow, protein synthesis and
brain activity. Decreasing environmental temperatures also allow the species to reduce resting
body temperature and heat production. The temperature effects aid in the reduction of metabolic
activities but are not solely responsible. Most organisms exhibit 2-3 fold depressions per 10 °C
reduction (Q10= 2-3) but the western painted turtle (Q10= 8.4 at 3-10 °C), the arctic ground
squirrel (Q10=14.1 at 4-20 °C), and the crucian carp (Q10=5.1 at 10–15°C) experience greater
decreases indicting other mechanisms at work [37, 80, 81].
Reduced metabolisms are supported for prolonged periods by increased internal fuel
stores. Both the crucian carp and the western painted turtles possess large livers with enhanced
glycogen concentrations in order to support extended anaerobic metabolism [39, 75]. Squirrels
increase fat stores and lean muscle prior to hibernation and aerobically metabolize all of it during
10
the winter season [57]. Waste products are still produced, although slowly and in small quantities
that can accumulate over the anoxic period. The strategies to deal with this accumulation are
diverse and less universal between the species discussed involving increases in buffering
capabilities, exchange processes between shell/bone, arousal periods to filter blood/excrete
wastes and the conversion of lactate into ethanol for expulsion into the environment [40, 72, 81,
82].
1.5 The importance of neuronal down regulation
The mammalian brain accounts for around 2% of total body mass but is responsible for
20% of the total body oxygen consumption making it the most energy demanding tissue [83, 84].
It is similarly responsible for a lower but significant percentage of the aerobic capacity within
reptiles and fish [3, 37, 85]. All of the species discussed above undergo decreases in neuronal
function during their overwintering periods believed to be vital for energy conservation and
anoxia/hypoxia tolerance. In the absence of these neuroprotective changes, oxygen deprivation
within the brain triggers a lethal cascade.
Approximately 50-60% of total brain ATP is used by cellular Na+/K+ ATPase in order to
maintain and re-establish cellular ion gradients [83, 84]. The gradients are perturbed through the
activities of post-synaptic currents, action potentials and neurotransmitter uptake [86]. The high
proportion of energy devoted to simply maintaining homeostasis makes neurons extremely
vulnerable to excitability during times of ATP depletion like hypoxia/anoxia. The loss of the
membrane ion gradients results in a net outward movement of K+ ions preventing cells from
repolarizing after firing. Na+ and Ca2+ channels become over activated and the whole cell is
depolarized. The resulting Ca2+ influx into the cell triggers vesicular release of excitatory
neurotransmitters such as glutamate into the synaptic cleft activating glutamatergic receptors on
associated cells [87, 88]. Neurotransmitter excision is compromised by ATP depletion and
glutamate lingers in the synapse causing receptor over activation and excessive Ca2+ influx.
Ca2+ plays important roles in second messenger pathways and metabolic regulation but in
high quantities is toxic to the cell. Under normoxic conditions extracellular Ca2+ levels are
maintained 10,000 times greater than intracellular levels through the activities of plasma
11
membrane transporters. Inside the cell calcium is stored in the mitochondria and endoplasmic
reticulum or sequestered by calcium binding proteins [92-94]. These processes are ATP
dependent and compromised by anoxia/hypoxia [95]. Excessive glutamatergic activation and
compromised Ca2+ management capabilities result in excessive intracellular concentrations of
free Ca2+ that cause cell swelling and trigger enzymatic cascades that eventually bring about cell
death [85][96]. This is termed excitotoxic cell death (ECD).
1.6 Models for potential human application
Stroke represents the third leading cause of death in North America killing 14,000
Canadians each year and hospitalizing a further 46,000 that may suffer neurological impairments
for the rest of their lives [89-91]. This represents a cost of $3.6 billion a year in physician
services, hospital fees, lost wages, and decreased productivity [92]. About 80% of all strokes are
caused by the interruption of blood flow to the brain due to a blood clot. The affected brain
regions are deprived of glucose and oxygen (ischemia), triggering the neurotoxic set of events
discussed above.
Tissue death caused by an obstruction of blood supply is termed infarction and cell death
in the tissues localized to the blockage is an infarct. Until blood flow is returned the infarct
grows in size affecting a greater proportion of the surrounding tissue. Within mammals, the size
of this infarct can be reduced and cell death prevented through “ischemic preconditioning” that
involves subjecting whole organisms to brief periods of ischemia prior to a prolonged lethal
ischemic insults in order to reduce infarct sizes. Ischemic preconditioning was first
demonstrated in canine myocardial tissue and has since been identified to offer protective roles
in a multitude of mammalian models within organ systems including brain, heart, liver, small
intestine, skeletal muscle, kidney and lung [93, 94] . In neurological evaluations in rats and
gerbils the development of global ischemia through temporary surgical blockage of the carotid
artery for 2-3 minutes reduced infarct sizes in response to lethal ischemia (5-10 minutes) by 53-
98%. Neuroprotection from lethal ischemia required 24-72 hours to develop after the initial
ischemic exposure and was maintained for 1-7 days [95-98].
12
The time span required for ischemic preconditioned neuroprotection to be achieved and
the short amount of time it is maintained for limit practical applications within the field of
medicine. As well, repeatedly exposing patients to brief ischemic periods is dangerous and
impractical. Attempts have been made to pharmacologically mimic the protective response but
have met with limited success in human application [94, 99]. Finally, ischemic preconditioning
has been demonstrated to reduce the size of infarcts but not prevent them all together resulting in
minor to moderate brain damage.
The hypoxia/anoxia tolerant organisms discussed above do not suffer neurological
impairments during their hibernation periods. Studying the mechanisms through which their
tolerances are achieved may offer an alternative route to dealing with conditions brought on by
blood flow reduction or may aide in our understanding and successful application of the
mechanisms controlling ischemic preconditioning. Preconditioning reveals that even within
mammals there is capacity to increase the hypoxia/anoxia tolerance of a tissue and the
underlying mechanisms may be similar to those used by anoxia-tolerant species. Of the three
species discussed above, the western painted turtle is the ideal model for study. As a terrestrial
vertebrate they are a better comparative model for potential human application than the crucian
carp. The neurological mechanisms permitting anoxia tolerance are triggered rapidly in response
to oxygen deprivation and not controlled circanually like the arctic ground squirrel. Lastly the
western painted turtle brain is tolerant to ischemia as well as anoxia [100].
1.7 Turtle synaptic organization
The anatomy and synaptic organization of the turtle brain has been investigated within
various turtle species and some of the structures have been compared against those found within
other organisms [101-104]. The turtle dorsal cortex shares several important anatomical
similarities with the mammalian neocortex including afferent and efferent connections to various
regions of the brain and electrically innervated organs [104-106]. However, although the
extrinsic connectivity of the turtle dorsal cortex resembles neocortex, the intracortical structure
of the turtle is greatly simplified compared to that of the mammal. Instead of a six-layered
isocortex, the turtle has only three cortical layers (molecular, cell and subcellular) that are
composed of two general classes of neurons: pyramidal cells that form the main output of the
13
cortex, and stellate interneurons [104, 107, 108]. The turtle brain has similarities to the
mammalian brain allowing for a degree of comparability in the context of anoxic/ischemic
investigation. However, it should be stressed that the resemblance between organs is limited.
Electrophysiological analyses have been performed within the dorsal cortex of the turtle
Pseudemys scripta elegans in order to better characterize the pyramidal and stellate cell types.
The pyramidal cells have their somata in the cell layer, extend several densely spined apical
dendrites through the molecular layer to the pia, and have spiny basilar dendrites directed
through the subcellular layer toward the ependymal border. Physiologically, pyramidal cells have
relatively prolonged action potentials that show marked frequency adaptation during a sustained
suprathreshold current pulse. In contrast, stellate cells have their somata in the molecular and
subcellular zones, and their dendrites are horizontally oriented and spine-free. Stellate cells have
relatively brief action potentials and show little or no spike frequency adaptation [101].
1.8 Anoxia mediated neuroprotective mechanisms
As discussed above, entrance into the overwintering state by the western painted turtle is
associated with major decreases in metabolic processes. Although the effects of decreasing
temperature aid in reducing metabolism (Q10 effects) oxygen deprivation is the primary trigger
for metabolic down regulation. Turtles submerged in tanks at 20 C experience a four-fold
depression in the frequency of heart contractions, and after 8 hours exhibit an 80 % reduction in
metabolic rate [35]. Similarly, submergence at 25 C decreases protein synthesis and urea
production by 92% and 72% respectively after 12 hours [36]. Anoxic down regulation of
metabolic activity leads to reductions in neuronal activity and the induction of a comatose-like
state as a result of spike and channel arrest.
1.8.1 Spike arrest
Gamma-aminobutyric acid (GABA) is the primary inhibitory neurotransmitter within the
mature central nervous system of vertebrates and mediates wide spread electrical depression in
most neurons [109-111]. Activation of postsynaptic GABAA receptors increases chloride (Cl-)
conductance through the channel’s pore region. Activation of postsynaptic GABAB receptors
increases K+ conductance via G-protein coupling to K+ channels. The reversal potentials for Cl-
14
and K+ are near neuronal resting potential and opening of GABA receptors generates an
inhibitory shunting current near the GABA reversal potential (EGABA) that impedes action
potential (AP) generation [112]. Within the brain of the western painted turtle, extracellular
GABA concentrations are elevated to 90 times normoxic levels after 4 hours of anoxia. As well
GABAA-receptor density increases by 22% and 29% after 12 and 24 hours of anoxia,
respectively [113, 114]. This produces decrease in whole brain activity demonstrated by
reductions in action potential frequency (APf), field potentials and electroencephalogram (EEG)
activity in the anoxic turtle [115-117]. Depressed electrical activity is termed “spike arrest” and
serves to conserve ATP throughout the anoxic period [118].
1.8.2 Channel arrest
Activation of post-synaptic N-methyl-D-aspartate receptors (NMDAR) is the primary
route for extracellular Ca2+ entry. NMDARs are hetero-tetramers formed by two GluN1 and two
GluN2 subunits (previously denoted as NR1 and NR2) of which there are four subtypes in
vertebrates (GluN2A-D) [Figure 1.2]. In mammals GluN2A and GluN2B are expressed
throughout the forebrain including the hippocampus, GluN2C is restricted to the cerebellum and
GluN2D is found predominantly in the midbrain [119]. Binding of glutamate to the GluN1 and
the co-agonist glycine to the GluN2 is necessary to trigger Na+/Ca2+ influx and K+ outflow [120].
NMDAR activation is inhibited at resting membrane potentials by a magnesium ion (Mg2+)
block within the pore region. The blockade is maintained in a voltage-dependent manner and
removed by cellular depolarization following postsynaptic α-amino-3-hydroxy-5-methyl-4-
isoxazolepropionic acid receptor (AMPAR) activation. AMPARs are hetero-tetramers composed
of different combinations of the four GluA (previously denoted as GluR) subtypes (GlurA1-4)
each possessing a binding site for glutamate [Figure 1.2]. AMPARs are permeable to Na+, K+
and also Ca2+ in configurations without GluR2. AMPARs are opened when two of the glutamate
binding sites become occupied producing a depolarizing current, the removal of the Mg2+ block
from NMDARs, and Ca2+ influx [121]. Activation of these receptors and the resulting Ca2+
influx plays important roles in learning, memory, synaptic plasticity and cognitive function
[122].
15
As previously discussed, glutamate concentrations are rapidly elevated following oxygen
deprivation within anoxia intolerant nervous systems. The resulting excessive Ca2+ influx is
produced primarily by NMDAR and AMPAR over activation. In the anoxic brain of the western
painted turtle, excessive glutamate release is partially prevented through GABA mediated
decreases in cell firing that keep glutamate levels relatively constant throughout (∼1 μM) [113,
123]. Adaptations are also present that decrease the degree to which NMDAR and AMPAR are
activated by glutamate during anoxia. Following 40 minutes of anoxia NMDAR and AMPAR
currents are decreased by 45-65% and 50-60%, respectively [124-128]. Rapid changes are
believed to be the result of reversible modifications made to the receptor subunits that affect
open time [129]. Over long term anoxia (3-21 days) NMDARs are removed from the cellular
membrane demonstrated through a 40% reduction in GluN1 subunit abundance and a depression
of Ca2+ permeability after 6 weeks [5]. It is unclear if a similar mechanism exists for AMPARs.
Rapid and long term changes in NMDARs and AMPARs are collectively termed “channel
arrest” and are utilized to conserve energy and prevent ECD [79].
16
Figure 1.2: NMDAR and AMPAR structures.
A, schematic diagram of the NMDA and AMPAR receptor complexes. B, schematic diagram of
the NMDAR and AMPAR subunit arrangements. Figures adapted from online sources [130, 131]
1.9 Mechanisms controlling “channel arrest”
Initial decreases in NMDAR and AMPAR whole cell currents are triggered rapidly in
response to oxygen deprivation. The mechanisms controlling these changes are believed to be
signalled by changes associated with anoxia. The effects of anoxia mediated changes in
adenosine, ATP-sensitive K+ (KATP) channels and intracellular calcium have all been
investigated.
1.9.1 Increases in adenosine
A temporary but significant decrease (20%) in ATP concentrations occurs in the brain of
the western painted during transition into anoxia. This decrease lasts for approximately an hour
before metabolic processes are reduced sufficiently for energetic costs to be met by anaerobic
17
glycolysis [34]. When ATP breakdown exceeds ATP production, AMP begins to accumulate in
the cell and 5’-nucleotidase activity increases. 5’-nucleotidases are a family of enzymes found in
the vertebrate CNS and inhibited by ATP. When activated they break down AMP into adenosine
[3, 132].
Adenosine is a “retaliatory metabolite” produced in response to oxidative stress in a
number of vertebrates [133]. Extracellular adenosine increases trigger protective changes
through interactions with adenosine receptors in different tissues (A1, A2A, A2B, and A3) [134].
Changes involved in neuroprotection include: suppression of pre-synaptic excitatory
neurotransmitter release (ex: glutmate), stimulation of anaerobic glycolysis, increased cerebral
blood flow and reduction of neuronal excitability via inhibition of Ca2+ channels/activation of G
protein-coupled inwardly-rectifying K+ channels [3, 134-138]. Neuronal extracellular adenosine
levels after 100 minutes of anoxia are 12 fold of normoxic concentrations [139]. This increase is
temporary and as metabolic processes are depressed in the turtle as discussed above, energy
needs are met by anaerobic glycolysis and normal ATP concentrations are re-established. After
300 minutes adenosine concentrations return to baseline levels [139].
Increased extracellular adenosine levels in the western painted turtle brain increase
resistance to K+ efflux in order to preserve cellular ion gradients. This is mediated by binding of
the A1 receptor [140]. Adenosine and A1 receptor activation has also has also been implicated as
a mediator of NMDAR channel arrest. Within neural tissue from the western painted turtle
normoxic adenosine addition reduces NMDA-mediated Ca2+ entry by 62 %. This decrease is
abolished by perfusion of the A1 antagonist 8-pentyltheophyilline (8-PT) [141]. However during
anoxia, A1 receptor antagonism does not block decreases in NMDAR open time or NMDA-
mediated Ca2+ entry [142]. This suggests that adenosine is capable of reducing NMDAR activity
but is not a necessary component of anoxia-mediated NMDAR channel arrest.
1.9.2 Activation of ATP-sensitive K+ (KATP) channels
Like adenosine production, KATP channel activity is controlled by ATP concentrations.
These channels, found in the plasma membrane and internal organelles like the mitochondria, are
gated by high concentrations of ATP and activated when levels are low like during oxidative
18
stress. These channels are found within the brain of vertebrates, particularly in the hippocampus
[97]. KATP channels are comprised of pore-forming Kir subunits and ATP-binding sulfonyl urea
subunits (SUR) [143]. The configuration and combination of the subunits varies based on the
type of membrane they are found within. The plasmalemmal KATP (pKATP) channel is composed
of two Kir6.1 and Kir6.2 bound to two SUR1 and SUR2 [144]. The mitochondrial KATP (mKATP)
is also thought to be a heteromultimer consisting of mitoKir and mitoSUR and containing neither
Kir6.1 or Kir6.2 [143, 145].
The freshwater turtle (Trachemys scripta) is another anoxia tolerant turtle species able to
survive at least 48 h of anoxia at 25°C [146]. Within this species increased pKATP channel
activity is necessary during anoxia for the maintenance of cellular ion gradients. K+ efflux from
neurons is reduced by 50% within one hour of anoxia, but abolished with administration of the
KATP channel blocker glibenclamide [147]. Increases in mKATP channel activity work in concert
with adenosine to block glutamate release within the first 3 hours of anoxia. Addition of the KATP
channel blocker BDM or the adenosine receptor antagonist theophylline produced increase in
glutamate release of approximately 70% above anoxic levels. At 5 hours, adenosine inhibition
produces an approximate 100% increase in glutamate release but KATP channel blocking has no
effect [136].
In the western painted turtle, mitochondrial specific KATP (mKATP) channels are involved
in anoxia-mediated down regulation of NMDAR and AMPAR currents. Addition of the mKATP
channel activator diazoxide under normoxic conditions reduces both whole-cell AMPAR and
NMDAR currents to levels comparable to anoxia. Inversely mKATP blockade with 5-
hydroxydecanoate (5-HD) abolishes anoxia-mediated reduction of NMDAR and AMPAR
currents [125, 126]. Activation of mKATP channels is also associated with an uncoupling of turtle
mitochondria by 10–20% and reduced Ca2+ uptake into the mitochondria [126] [148]. The
resulting rise in intracellular Ca2+ levels can bring about downstream effects in NMDAR and
AMPAR activity.
19
1.9.3 Increases in intracellular calcium
Following the onset of anoxia in turtle neural tissue, intracellular calcium ([Ca2+]i )
concentrations increase [126, 128, 149]. The [Ca2+]i increase is necessary for the development of
anoxia mediated AMPAR/NMDAR channel arrest. The inclusion of the Ca2+ chelator 1,2-bis(o-
aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA) in the pipette solutions of whole-cell
patch clamp recording electrodes abolishes anoxia-mediated reductions in AMPAR/NMDAR
currents [125, 129]. Increases in [Ca2+]i occur in the absence of extracellular calcium, indicating
that Ca2+ is released from an intracellular source [126, 128, 149]. Within neurons Ca2+ is stored
in the mitochondria and the endoplasmic reticulum (ER). Inhibition of ER specific calcium
release via ryanodine does not abolish anoxia mediated NMDAR current reductions, indicating
mitochondrial specific Ca2+ release is responsible for anoxia mediated changes [126]. As
discussed above activation of mKATP channels is associated with mitochondrial uncoupling and
reduced Ca2+ uptake. Activation of mKATP channels with diazoxide or levcromakalim produces
increases in [Ca2+]i and decreases in NMDAR/AMPAR currents of similar magnitude to those
seen during anoxia. The addition of the mKATP antagonists 5-HD or glibenclamide prevented
both of these changes [125, 126]. Increases in [Ca2+]i trigger the activation of a variety of
intracellular regulating proteins that can serve as the intermediate step in signalling
NMDAR/AMPAR current down regulation in response to mitochondrial Ca2+ release.
1.9.4 Enhancements of calmodulin and phosphatase activity
Calmodulin is a calcium-binding messenger protein expressed in all eukaryotic cells.
Calmodulin can modulate NMDAR directly by binding to the COOH terminus of the NMDAR
GluN1 subunit and inactivating the channel in a Ca2+-dependent manner [150]. Within the
western painted turtle inhibition of calmodulin with calmidazolium attenuates anoxia mediated
decreases of NMDAR currents within neural tissue [129]. Inhibition has yet to be evaluated on
anoxia mediated AMPAR current reductions.
NMDAR activity can also be reversibly modulated by intracellular kinases and protein
phosphatases (PPs), some of which are controlled by intracellular calcium concentrations.
Dephosphorylation via PP1, PP2A and the Ca2+/calmodulin-dependent phosphatase PP2B
20
(calcineurin) all attenuate NMDAR activity [151-154]. Within the anoxic turtle brain, addition of
the PP1 inhibitor okadaic acid or the PP1/PP2A inhibitor calyculin A prevent anoxia-mediated
attenuation of NMDAR currents [129]. The inhibition of calcinuerin via cypermethrin similarly
attenuates NMDAR currents over the short term (20 minutes) but not the long term suggesting
that calcinuerin might only play a role in early attenuation [129]. The effect of phosphatases on
anoxia mediated AMPAR current reductions have yet to be tested.
Figure 1.3: Schematic illustration of anoxia-mediated changes contributing to channel arrest.
1.10 Reactive oxidative species (ROS)
Other cellular changes aside from decreases in intracellular ATP concentrations occur
during anoxia that may contribute to “channel arrest” and/or “spike arrest”. One such change is
reductions in reactive oxidative species (ROS). All reactive molecules and free radicals produced
as unwanted by-products from the incomplete reduction of diatomic oxygen (O2) during
oxidative metabolism are termed as ROS [155]. All aerobic organisms produce ROS and have
developed cellular anti-oxidant defence systems that prevent concentrations from reaching
harmful levels. Under normoxic conditions, intracellular ROS levels are maintained at relatively
21
low concentrations but never completely removed. During oxygen deprivation in anoxia tolerant
organisms, oxidative phosphorylation halts and ROS levels decline to immeasurable levels [155].
Within the brain of the western painted turtle CM-H2DCFDA fluorescence evaluations have
demonstrated that ROS concentrations decrease by 25% within 10 minutes of anoxia [156].
ROS are primarily produced within the mitochondria between complexes and I and III
where 2-3% of all O2 consumed is left partially reduced as the highly reactive superoxide anion
(O2) [Figure 1.3] [157, 158]. This molecule is short lived and in most cases binds with water
either spontaneously or through induction by mitochondrial superoxide dismutase (SOD) to
produce hydrogen peroxide (H2O2) [159-161]. H2O2 is one of the longest lived cellular ROS but
is eventually converted to water and O2 by catalase or glutathione peroxidase (GPx) [93, 162].
Levels of H2O2 are directly linked to metabolic activities and increases or decreases can serve to
signal cellular functions such as apoptosis, gene expression, and the activation of cell signaling
cascades via reversible oxidation of cysteine (Cys) thiol residues on signalling proteins and
membrane ion channels [163-167]. H2O2 is cell permeable and capable of moving through the
lipid bilayer by free diffusion or through aquaporins (AQP)/water channels and oxidizing
extracellular as well as intracellular Cys residues. In this manner, H2O2 acts as an inter and intra
cellular messenger [168-171]. The oxidized Cys residues can be reduced by elements of the
antioxidant defence system or can react with other sulfide compounds to produce disulfide
bridges [Figure 1.3] [165, 168-171].
Although there remains controversy over whether CNS neurons contain aquaporins
neurons do exhibit low water permeability under resting conditions and are capable of moving
water through alternative means such as membrane cotransporters (ex: KCC1) [172]. H2O2 is
very water soluble and should be capable of following the movement of water through available
channels similar to movement through APQs [168, 173, 174]. For the purposes of this thesis, the
routes for facilitated water transport (aquaporins, cotransporters, or other) will be grouped and
designated as water transport routes (WTRs) for the sake of simplicity.
Another source of cellular ROS is nitric oxide (NO) produced by neuronal NO synthase
(nNOS). nNOS is tethered to the NMDAR by the postsynaptic density protein-95 (PSD95) [175,
176]. nNOS produces NO from arginine and O2 and is triggered by the activated Ca2+-binding
22
protein calmodulin (CaM) [Figure 1.3] [177-179]. Tethering of nNOS to PSD-95 and Ca2+
dependent activation couple the production of NO to NMDAR activation. Like H2O2 NO is short
lived, capable of moving through the lipid bilayer by free diffusion or through WTRs, and can
reversibly oxidize cysteine residues through a processes termed S-nitrosylation. NO can also
react with O2 and form peroxynitrite (ONOO) that is another effective oxidizing agent. It can
also activate soluble guanylate cyclase (sGC) and produce cyclic guanosine monophosphate
(cGMP) capable of triggering downstream changes that include further ROS generation [168,
179-182]. [Figure 1.3].
1.11 The role of ROS in anoxia-intolerant vertebrates
Under normoxic conditions, the role of ROS in cellular signalling has been well
documented in a number of anoxia-intolerant organisms. Changes in ROS increase and decrease
the activities of a number of ion channels and intracellular proteins [183, 184]. Changes in the
NMDAR redox state and intracellular ROS levels affect NMDAR activity. Like anoxia-mediated
regulation, these changes appear to be affected by the phosphorylation state of the NMDAR.
1.11.1 Redox changes affect NMDAR but not AMPAR activity
The structure and function of several membrane bound receptors containing Cys residues
are sensitive to redox conditions [185]. Specific Cys residues in the NMDAR form redox
modulatory sites whose oxidation/reduction state affects the activity of the receptor. Redox
modulatory sites have been demonstrated to exist within GluN1, GluN2A and GluN2B subunits
[186-188]. Experiments using site-directed mutagenesis in rat and frog oocytes have identified
residues within the GluN1 (Cys744, Cys798, Cys 79 and Cys 308) and GluN2A (Cys 87, Cys
320 and Cys 399) subunits whose alteration affects redox modulation but the site in the GluN2B
has yet to be localized. All of the sites identified have been localized to extracellular regions of
the receptor subunits within their respective species. However, there is some controversy as to
whether intracellular or transmembrane redox sites also exist but have yet to be identified [185,
187-189].
23
The effect of redox modulation on NMDA receptors has been observed in a wide variety
of tissue preparations including intact chick retina, frog oocytes, rat retinal ganglion,
hippocampal, cortical and striatal slices and post-mortem human cortex [188, 190-198].
Evaluations of NMDAR activity within these preparations using methods including intracellular,
extracellular, and patch-clamp recording have demonstrated that addition of oxidizing agents
(DTNB and GSSG) decrease receptor activity while reducing agents (DTT) increase receptor
activity. The increases produced through reduction are greater than decreases through oxidation
(6-11 times), suggesting that redox sites are maintained in a more oxidized state [190] [196,
199].
Unlike the NMDAR, no modulatory redox sites have been found in the structures of the
AMPAR GluR subunits. Within many of the same tissue preparations used to demonstrate the
effects of NMDAR redox modulation, AMPA addition has been shown to be unaffected by
oxidizing or reducing agents [190, 196-198].
1.11.2 Alterations in ROS affect NMDAR activity
Like reducing/oxidizing agents, changes in ROS have been shown to alter NMDAR
activity. Within rat cortical neurons oxygen free radicals generated by xanthine and xanthine
oxidase (X/XO) are capable of reversing increases in NMDAR activity brought on by reducing
agents and decreasing activity similar to oxidizing agents [191, 196]. The inhibition of
mitochondrial ROS production has also been demonstrated to effect NMDAR activity. Rotenone
is a complex I inhibitor and decreases mitochondrial ROS production in mitochondria from rat
brain, heart, liver, dog brain and human cell cultures [157, 200-202]. This is accomplished
through inhibition of electron transfer from iron-sulfur centers in complex I to ubiquinone
[Figure 1.3]. In whole cell patches of dopamine neurons from slices of rat midbrain the addition
of rotenone increases the average amplitude of evoked NMDAR currents by 162% without
affecting AMPAR activity [203].
The effect of NO manipulation has also been evaluated in cultured rat cortical neurons
and frog oocytes through Ca2+ imaging and whole cell patch clamp recordings. Like oxidizing
24
agents and ROS generators, the application of NO generators produced marked decreases in
NMDAR activity [189, 204, 205].
1.11.3 NMDAR redox regulation is blocked by kinase inhibition
As discussed above, increased phosphatase activity is associated with decreases in
NMDAR activity during anoxia in the turtle brain [151-154]. There is some evidence to suggest
that phosphorylation of the NMDAR is necessary for oxidation/reduction of the redox
modulatory site. Electrophysiological recordings in hippocampal rat slices demonstrated
increases in NMDAR-mediated synaptic responses with DTT application. Increases were
blocked by inhibition of Ca2+/calmodulin-dependent protein kinase II but not by inhibition of the
protein phosphatases PP1 or calcineurin [196]. Similarly in horizontal slices from rat ventral
midbrain rotenone augmentation of N-methyl-D-aspartate (NMDA)-evoked currents were
inhibited by the inclusion of a tyrosine-kinase in intracellular pipettes [206]. These findings may
indicate that the NMDAR redox site is only accessible when the NMDAR is phosphorylated.
25
Figure 1.4: The mechanisms of ROS production and cysteine oxidation.
A, depiction of the mitochondrial electron transport chain showing the sites of reactive oxygen
species production and rotenone inhibition. B, depiction of NMDAR coupled nNOS activation
and nitric oxide synthesis. C, depiction of the effect of H2O2 and NO oxidation on cysteine
residues. Figures adapted from (A) Pamenter 2007 and (B) Nakamura 2007 [156, 179].
26
1.11.4 mKATP mediated ROS increases affect ischemic preconditioning
Changes in ROS have recently been linked to the development of protection produced
through ischemic preconditioning. Like anoxia mediated NMDAR/AMPAR downregulation in
the brain of the western painted turtle, activation of the mKATP channel has been identified to be
involved in ischemic preconditioning. Within cardiac tissue from rats and rabbits the addition of
the selective mKATP antagonist 5-HD prevents the development of protective effects from
ischemic preconditioning [207-209]. In neural tissue from rats and pigs the addition of the
mKATP agonist diazoxide protects the brain against ischemic and anoxic stresses similar to
ischemic preconditioning [210-213].
Activation of mKATP channels in the western painted turtle is believed to trigger a mild
increase in intracellular calcium necessary for the development of downstream changes. In
ischemic preconditioning however the activation of mKATP channels and downstream changes
are associated with increased production of mitochondrial ROS. In rat hippocampal slices,
isolated rabbit hearts, and human atrial-derived cell lines the addition of the ROS scavenger N-2-
mercaptopropionylglycine (MPG) reversed diazoxide mediated protection similar to 5-HD
addition [214-216]. Fluorescent investigation in human atrial derived cell line, rabbit myocardial
tissue, and rat retinal tissue has demonstrated that mKATP channel activation increases oxidation
of the detective probes indicative of increases in cellular ROS levels [216-218]. This seems to
indicate that mKATP channel opening increases ROS generation that then triggers the
development of the preconditioned state.
There is also some evidence to suggest that mKATP channels are more favourably
activated in the presence of mitochondrial ROS or NO. Within rat myocytes, reconstituted
bovine ventricular myocardium mitochondria, embryonic chick ventricular myocytes, rabbit
ventricular myocytes, rat dorsal root ganglion neurons, and rat cardiac submitochondrial particles
the addition of NO donators, O2•- generators, ROS scavengers and exogenous H2O2 have all
been demonstrated to affect mKATP activity [219-224]. Increased ROS caused mKATP channels to
become less inhibited by ATP and more readily activated. This increase has been attributed to
27
increased oxidation of cysteine residues in the SUR subunit, similar to what occurs in the
NMDAR redox site [223]. In this way, mKATP activation during ischemic precondition may
cause channel oxidation and promote further activity in a positive feedback loop [221].
1.12 The antioxidant defence system
Antioxidants are substances that when present at low concentrations, compared to that of
an oxidizable substrate, significantly delays or inhibits oxidation [225]. The antioxidant defence
system protects the cell from unwanted reactions by maintaining levels of ROS generated
through processes like nNOS activation and mitochondrial oxidative phosphorylation at low
concentrations [226].
The three intracellular antioxidants most relevant to nNOS and mitochondrial ROS
generation are superoxide dismutase (SOD), catalase and glutathione (GSH)/glutathione
peroxidase (GPx). SOD is found within the mitochondria of all eukaryotes and differentiated
from SOD found in chloroplasts and prokaryotic organisms by the type of metal cofactor
(MnSOD) [227]. SOD catalyses the dismutation of O2 , produced from oxidative
phosphorylation, into H2O2 at a reaction rate that is 10,000-fold faster rate than spontaneous
dismutation (O2 + O2
+ 2H+ => 2H2O2 + O2) [228]. SODs are nuclear encoded and there is
some evidence to suggest that synthesis is sensitive to environmental stressors and can be altered
in response to changes in ROS production [227]. Intracellular concentrations of H2O2 are
managed through a wide range of enzymes, including catalase and GPx. Catalase forms a
intermediate complex with H2O2 and generates water and O2 (2H2O2 => O2 + 2H2O). GPx
catalyzes the reaction of GSH with H2O2 to produce oxidized glutathione (GSSG) the helps to
maintain a redox balance in the cellular compartments (2GSH + H2O2 => GSSG + 2H2O). A
central nucleophilic cysteine residue is responsible for the high reductive potential of GSH,
allowing it to scavenge cytotoxic H2O2 and reacts non-enzymatically with other ROS like
singlet oxygen, superoxide radicals, hydroxyl radicals, and NO [226, 227]. The cumulative
actions of these and other antioxidants maintain the intracellular environment within a reduced
state [168].
28
As discussed above both H2O2 and NO are capable of leaving the cell through free
diffusion or WTRs and interacting in the extracellular environment. The resulting extracellular
ROS concentrations are managed by antioxidants produced within the cell and transported to the
extracellular environment. GSH, catalase and GPx act in the extracellular environment but
generally exist at lower concentrations than found within the cell [229]. The major extracellular
thiol is free L-Cys that exists in both a reduced (Cys) and oxidized (Cyss) form, although in the
plasma the oxidized form is predominant. Influx and efflux of reduced Cys between the
intracellular environment may contribute to extracellular redox status and help to maintain a
Cys/Cyss balance [168]. Due to lower antioxidant concentrations and the movement of ROS out
of the cell, the extracellular environment exists within a predominantly oxidized state [168].
Antioxidants in both the extracellular and intracellular environments can act to reverse
ROS reactions, such as by reducing thiol residues in proteins oxidized through ROS interactions.
Aside from scavenging roles, GSH and free L-Cys have been implicated to act as the primary
intracellular and extracellular protein reductants, respectively [230]. Intracellular and
extracellular levels are maintained relatively constant in most cells and act to establish a baseline
balance with ROS products between oxidation and reduction states of cellular proteins [168,
230].. During anoxia in the western painted turtle when ROS concentrations fall the balance can
be disturbed leading to a shift towards protein reduction in both the intracellular and extracellular
environment.
1.13 ROS regulation of GABA in anoxic turtle cortex
As previously discussed one of the mechanisms through which turtles reduce ATP
consumption within the brain during the over wintering period is through increases in GABA
receptor postsynaptic currents (PSCs) that serve to decrease whole brain activity via spike arrest
[113, 114]. Recent investigation within our lab has demonstrated that in cortical slices from the
western painted turtle under normoxic conditions, ROS scavenging increased GABAergic PSC
amplitudes similar to anoxic exposure. Application of the mitochondrial complex IV inhibitor
cyanide decreased intracellular ROS to the same extent as anoxia and also resulted in a similar
increase in GABAergic PSCs. Finally GABAergic PSC increases from ROS scavenging and
29
anoxia were both reversed by exogenous application of H2O2 [231]. These findings demonstrate
that anoxic decreases in mitochondrial ROS production signal increases in GABA release.
1.14 Rationale for study
Decreases in reactive oxidative species following entrance into anoxia within the western
painted turtle have been documented [156] [231]. Pharmacologically scavenging ROS or
inhibiting mitochondrial ROS synthesis under normoxic conditions increases GABAergic PSCs
similar to anoxia [231]. Theses finding demonstrate that ROS decreases play a role in signalling
anoxia-mediated changes. Aside from increases in GABA release, during anoxia neuroprotective
decreases in NMDARs and AMPARs currents also occur and are associated with mild increases
in intracellular calcium. The primary aim of this study was to determine whether
pharmacological reduction of cellular ROS under normoxic conditions produced decreases in
whole-cell evoked NMDAR/AMPAR currents similar to those seen during transition from
normoxia to anoxia. The secondary aim was to evaluate whether the mild increase in Ca2+
associated with receptor down regulation is also affected by changes in ROS.
Within anoxia-intolerant vertebrates decreases in ROS and reduction of the NMDAR
redox site are associated with increased NMDAR activity. Because the western painted turtle
experiences decreases in ROS during entrance into anoxia, you would expect to see a similar
increase in activity if a similar mechanism were present. Instead NMDAR activity is
downregulated suggesting that either the structure of the NMDARs in the turtle brain are
different from those seen in other documented vertebrates and respond to ROS changes in an
opposing manner or an anoxia mediated mechanism is present that overrides and reverses normal
redox modulation. The mechanism will also need to down regulate AMPAR activity, that is
documented to be unaffected by ROS modulation.
Based on the material discussed above on ion channel control during normoxia and
anoxia, there are a couple of viable routes through which anoxia-mediated channel arrest could
be accomplished. Redox control could be overridden through increases in the activity of cellular
phosphatases. As discussed above, there is evidence that NMDARs may need to be
phosphorylated in order for redox modulation to take place. Increases in phosphatase activity
30
suggested to occur during anoxia in the western painted turtle may be sufficient to override and
reverse redox modulation. The increases in phosphatase activity may be also triggered by ROS
changes. As discussed above the activity of the mKATP channel can be affected by its
oxidation/reduction state and in anoxia-intolerant organisms is preferentially activated in the
presence of ROS. The mKATP subunit that exists in the turtle brain may be modified such that it
reacts to decreases in ROS/ increased reduction by becoming more easily activated and
triggering anoxia-mediated increases in intracellular calcium and phosphatase activity. Based
upon this reasoning I propose the following:
Hypothesis: Decrease in reactive oxidative species during anoxia triggers a down regulation of
NMDAR and AMPAR whole cell currents via an increase in the activity of mKATP and a
subsequent increase in intracellular calcium.
Aims:
1. Pharmacological reduction of cellular ROS will reduce normoxic NMDAR/AMPAR currents
and addition of H2O2 under anoxia conditions will increase NMDAR/AMPAR currents.
Using whole-cell patch-clamp techniques NMDAR/AMPAR whole cell currents will be
measured following pharmacological manipulation of ROS under normoxic and anoxic
conditions.
2. Decreases in ROS will trigger increases in cytosolic calcium levels comparable to those
measured during anoxia.
The fluorescent Ca2+ indicator dye, Oregon Green, will be used to measure changes in cytosolic
calcium levels following anoxic exposure and/or pharmacological manipulation of ROS.
31
CHAPTER 2: MATERIALS AND METHODS
2.1 Animals
This study was approved by the University of Toronto Animal Care committee and
conforms to the relevant guidelines for the care and handling of experimental animals as outlined
in the Guide to the care and Use of Experimental animals as determined by the Canadian Council
on Animal Care.
Adult male and female western painted turtles were purchased from Niles Biological Inc.
(Sacramento, CA, USA). Animals were housed in an indoor aquarium equipped with a flow-
through de-chlorinated freshwater system, a heating lamp, and a basking platform. The water
temperature was maintained at 20°C. The turtles were given continuous access to food, and were
kept on a 12h:12h light-dark photoperiod.
2.2 Dissection and whole-cell patch-clamp recording protocol
Turtles were selected at random and decapitated. Whole brains were rapidly excised from
the cranium of western painted turtles and cortical sheets were isolated and placed in artificial
cerebral spinal fluid (aCSF; mm: 107 NaCl, 2.6 KCl, 1.2 CaCl2, 1 MgCl2, 2 NaH2PO4, 26.5
NaHCO3, 10 glucose, 5 imidazole, pH 7.4; osmolality 280–290 mosmol l−1). Individual cortical
sheets (3-4) were cut medially from the visual cortex of each cerebral hemisphere [Figure 2.1].
32
Cortical sheets were placed in an RC-26 chamber with a P1 platform (Warner
Instruments, CT, USA). During recordings of NMDAR activity, cortical brain tissues were
bathed in standard aCSF. During recordings of AMPAR activity, cortical tissues were bathed in a
modified high Mg2+ aCSF solution [4 MgCl2] in order to block NMDAR activity, and isolate
AMPAR currents specifically.
aCSF solutions were gravity perfused into the chamber via 1-L glass bottles attached to
intravenous (IV) drippers. A three way valve allowed for transitions to be made between initial
and experimental bottles during recordings. Conditions of normoxia were developed by bubbling
with 95% O2/5% CO2. Investigations performed under anoxic conditions were accomplished by
switching to a secondary bottle of aCSF bubbled with 95% N2/5% CO2. The perfusion tubing
was double jacketed and gassed with 95% N2/5% CO2 to maintain anoxic conditions. As well, a
plastic rim with anoxia tubing connected was placed around the saline bath to gently gas 95%
N2/5% CO2 during anoxic conditions. All experiments were conducted at room temperature (22
°C).
Whole-cell recordings were performed using 5-8 MΩ micropipettes produced from
borosilicate glass capillary tubes with a P-97 micropipette puller model (Sutter Instruments).
Micropipettes were fire-polished using a MF-830 Microforge (Narashige Group). Pipette
solution contained the following (in mM): 8 NaCl, 0.0001 CaCl2, 10 Na-HEPES, 130 potassium
gluconate, 1 MgCl2, 0.3 NaGTP, and 2 NaATP (adjusted to pH 7.4 and osmolarity 290-300
mOsM). Cell-attached 1-20 GΩ seals were obtained using the blind-patch technique of Blanton
et al. (1989). To break into the cell, the recording electrode potential was voltage-clamped to -80
mV and a sharp pulse of negative pressure was applied. Once the whole-cell configuration was
33
established, cells were given at least 2 minutes to acclimate before series resistance was
measured, which normally ranged from 20-100MΩ. Patches were discarded if series resistance
varied by >25% over the course of an experiment. Data was collected at 5-10 kHz using an
Axopatch-1D amplifier, a CV-4 head stage, and a Digidata 1200 interface (Axon Instruments)
and stored on computer using Clampex 10 software (Axon Instruments). A fast-step perfusion
system (VC-6 perfusion valve controller and SF-77B fast-step perfusion system; Warner
Instruments) was used to deliver pharmacological agents to the tissue.
Figure 2.1: Diagrammatic representation of cortical sheet excision and experimental setup.
Figure adapted from an online source [232].
34
2.3 Neuron identification
All investigations were carried out within pyramidal cells. Neurons were identified as
pyramidal or stellate based on the expressed electrophysiological properties. Previous to all
whole cell patch clamp experiments cells were current clamped and a current was injected for
450 ms to cause a 30 mV depolarizing change in membrane potential and action potential firing.
Characterization was accomplished by examining the adaptation pattern and amplitudes of
action-potential firing and comparing these to the standards previously developed for
differentiating between stellate and pyramidal neurons [101].
2.4 Evoked NMDAR and AMPAR current amplitude recordings
After patches stabilized neurons were voltage-clamped to a holding potential of -80 mV
for the entirety of the recording period. All recordings were carried out within the same time
frame in order to ensure comparability. Recordings were made at 0 min, 10 min, 30 min, 50 min,
70 min, and 90 min [Figure 2.2]. Access resistance was measured at the beginning and end of
each recording and if resistance fluctuated by >20% from the initial value patches were
discarded. Initial recordings at 0 and 10 were made under normoxic conditions to establish a
baseline for comparison and confirm currents were stable. The initial amplitude recording at t= 0
min was set to 100% and all subsequent recordings were normalized to that control value for
comparison purposes. Patches were discarded if t=10 min varied by more than 20% from t= 0
min. Experimental conditions were induced immediately after the establishment of a stable
baseline and currents were evaluated after 20 and 40 minutes. Normoxic conditions were then
reintroduced in order to reverse experimental changes and evaluated after 20 and 40 minutes.
35
The peak current values were taken from the recordings and used for the purposes of comparison
and statistical analysis.
Tetrodotoxin (TTX) was administered via drip perfusion 5 minutes previous to all
recordings in order to prevent the opening of voltage gated sodium channels and ensure currents
elicited were only from NMDAR/AMPAR activation. The only exception was during H2O2
experiments where 5 minute TTX addition was carried out 10 minutes prior to all recordings.
This allowed for H2O2 to be drip perfused 5 minutes prior to recordings to optimize its effects.
The effects of TTX have been shown to extend beyond 15 minutes ensuring that
NMDAR/AMPAR currents were isolated throughout the extended recording period. After TTX
administration cortical sheets were drip perfused with NMDA or AMPA for 3-10 seconds to
induce a current response. The same application time was then used for every subsequent
recording performed on the same neuron within a single experiment.
Figure 2.2: NMDAR and AMPAR whole-cell patch clamp recording timeline.
NMDAR and AMPAR current amplitude measurements were all taken at specified time points
(top line) during varying treatments (bottom).
36
2.5 Fluorescence investigation of cytosolic Ca2+ changes
To assess changes in cytosolic Ca2+ cortical slices were loaded with the Ca2+-sensitive
dye Oregon-Green 488 BAPTA-1 AM (Invitrogen). Oregon-Green 488 BAPTA-1 was chosen
due to its high Ca2+ affinity (Kd ≈170 nM). Tissue sheets were incubated individually in an
opaque vial containing 2 mL of aCSF with 40 uM Oregon-Green (from a 1 mM Oregon-Green
stock solution in dimethylsulfoxide and 20% pluronic acid). Cortical slices were loaded within
the vials for 3 hours at 3-5 °C and subsequently washed in unmodified aCSF for 15 minutes.
Following dye loading slices were placed in a flow-through recording chamber as previously
described for whole-cell patch clamp experiments. During anoxic experiments a plastic cuff was
placed around the objective in order to gently gas 95% N2/5% CO2 across the bath.
Oregon-Green was excited at 488 nm using a DeltaRam X high-speed random access
monochromator and a LPS220B light source (Photon Technology International) at a bandwidth
of 4.5 nm. Fluorescence measurements were acquired at 1 second intervals using an Olympus
BX51W1 microscope and U-CMAD3 camera. Fluorescence emission measurements at 488 nm
were acquired at 10 second intervals under 400x magnification using an Olympus BX51W1
microscope (Olympus Canada Inc.) and Rolera-MGi Digital EMCCD camera (QImaging).
Camera gain and exposure time values were 4.0 and 3 s, respectively. Previous to all recordings
excitation scans were performed from 450 to 550 nm in order to confirm peak excitation at 488
nm and successful cellular loading of Oregon Green.
Fluorescent recordings were all carried out within the same time frame to ensure
comparability. The time frame was reduced compared to that utilized within whole cell patch
37
clamping experiments to compensate for the rapid rate of photo bleaching and ensure that the
fluorescence signal was not exhausted. Cells were recorded under normoxic conditions for 10
minutes to obtain a stable baseline, followed by a treatment period for a further 10 minutes, and
concluded with normoxic reperfusion for a final 10 minutes [Figure 2.3]. All fluorescent
experiments were conducted with bulk perfusion as drip perfusion proved unreliable and affected
image stability. Solutions were gas bubbled in the same manner as solutions used for whole cell
patching. After the reperfusion period, aCSF flow was halted and tissues were incubated with the
Ca2+ ionophore ionomycin (2 μmol) for 5 minutes to increase extracellular Ca2+ uptake and
obtain the maximal fluorescence signal (Fmax). This was followed by addition of MnCl2 (2 mM)
to preferentially bind the Oregon green dye and quench fluorescence so that the minimal
fluorescence signal could be obtained (Fmin). The protocol and concentrations used were based
on previous investigations [149, 233].
A minimum of 10 neurons were selected for analysis in each experimental recording
period. In order to ensure good dye uptake and cellular health neurons exhibiting low to
moderate basal calcium fluorescence were chosen. The change in fluorescence was determined
from a line of slope drawn over the treatment area and compared against a parallel line of slope
across the pre and post treatment area (resting fluorescence signal) [see Figure 3.7 for
illustration]. The difference between these lines at the midpoint of the experimental period (F)
was taken and divided by the established range of fluorescence (Fmax-Fmin) to establish the
change in Ca2+ fluorescence. Expressed as the function: Ca2+ Change= (F/(Fmax-Fmin)) x 100.
The percent change value for all neurons selected within a trace was averaged and used as a
single n-value for final statistical analysis.
38
Figure 2.3: Oregon Green fluorescent recording timeline.
Fluorescent recordings were all carried out within the timeframe with the treatment applications
(bottom) all at the same time points (top).
2.6 Whole cell spontaneous neuronal activity recordings
Spontaneously firing pyramidal neurons were utilized to assess changes in neuronal
activity. Whole cell patches were obtained using the same procedure as described above.
Neurons displaying spontaneous activity were confirmed to be pyramidal cells in the same
manner as used in evoked NMDAR/AMPAR recordings. Following successful identification
neurons were recorded over a 40 minute time frame. Cells were recorded under normoxic
conditions for 5 minutes to establish a measurement of baseline activity, followed by a treatment
period for 20 minutes, and concluded with normoxic reperfusion for a final 15 minutes [Figure
2.4]. All experimental solutions were administered via bulk perfusion. AP frequency (APf) was
evaluated for 2 minutes after the cell was allowed to stabilized for 1 minute (t=1-3 min). This
value was set to 1 and all subsequent values were normalized to the value for comparison
purposes. APf values for pre-treatment/normoxia (t=3-5 min), treatment (t=23-25 min), and
washout/recovery periods (t=38-40 min) were evaluated within the last 2 minutes of each
treatment period.
39
Figure 2.4: Whole cell spontaneous activity recording timeline.
Spontaneous activity recordings were all carried out within the timeframe with the treatment
applications (red) all at the same time points (black). APf was evaluated for 2 min within the
various treatment periods (blue).
2.7 Pharmacology and drug application
During NMDAR/AMPAR current measurements drugs were applied either by drip perfusion
alone or in conjunction with bulk perfusion. Drip perfusion was utilized for agents whose mode
of action required little exposure time or whose extended presence could prove toxic to the cell.
1 uM TTX, 300 uM NMDA and 50 uM AMPA were concentrations based on previous
publications within our laboratory and were all exclusively drip applied as all agents could be
toxic during longer exposures [125-127, 234]. 50 uM H2O2 represents the primary mitochondrial
ROS product and was used to test the effects of increased ROS presence on receptor activity.
Concentrations were identical to those used in previous experiments with turtle cortical neurons,
where 50 uM was the lowest concentration in which a significant change was detected in CM-
H2DCF ROS fluorescence [231]. H2O2 was drip perfused for 5 minutes only in order to try and
40
limit excessive reactions. Drip solutions were appropriately bubbled with 95% O2/5% CO2 or
95% N2/5% CO2 to ensure they were normoxic/anoxic.
Drugs were bulk perfused in the same manner as the normoxic and anoxic solutions for
both electrophysiological and fluorescence experiments. Separate 1 litre IV bottles were used
and filled with the respective drug solutions and transition between conditions was made using a
three way valve. All drug agents were bubbled with 95% O2/5% CO2 in order to ensure they
were normoxic. Cell permeable ROS scavengers N-2-mercaptopropionylglycine (MPG, 0.5 mM)
and n-acetylcysteine (NAC, 0.5 mM) were used to artificially decrease normoxic ROS and test
the effects of whole cell ROS decreases on NMDAR/AMPAR currents. Concentrations were
identical to those used in previous experiments with turtle cortical neurons where 30 minutes of
0.5 mM MPG/NAC decreased CM-H2DCF ROS fluorescence comparable to what was
documented for anoxia within the same time frame [231]. Rotenone (25 uM) was utilized to
inhibit mitochondrial complex I, prevent mitochondrial specific ROS production and determine
the role of mitochondrial ROS in NMDAR/AMPAR control. A rotenone concentration of 25 uM
was previously validated as safe in turtle cortical slices [156].
2.8 Measurement of oxygen tension
Oxygen concentrations within the recording chamber during induced anoxic periods were
tested using an E101 oxygen electrode (Cameron Instruments Co.) containing 1M KCl and 20
μM sodium azide connected through a Model 1900 polarographic amplifier (A-M Systems) to a
LabPro interface (Vernier Software and Technology) for computer data collection. Logger Pro
3.8.4.2 was used for data collection at a sampling rate of 2 seconds. Oxygen tension was
41
calibrated to approximately 160 mmHg by placing the electrode in a glass bottle bubbled with
21% O2/79% N2 and to 0 mmHg in a glass bottle bubbled with 95% N2/5% CO2. The electrode
was submerged in aCSF within a chamber identical to that used during fluorescent and whole-
cell patch-clamp recordings.
2.9 Chemicals
Tetrodotoxin was purchased from Tocris Bioscience (Ellisville, MO, USA) and Oregon
Green 488 was purchased from Invitrogen Life Technologies (Burlington, ON, Canada). All
other chemicals were obtained from Sigma-Aldrich Chemical Co. (Oakville, ON, Canada).
2.10 Statistical analysis
Electrophysiological and fluorescent data was analyzed using SigmaPlot software
package version 11.0 (Systat Software, Inc., San Jose, CA). NMDAR/AMPAR whole-cell peak
current data and Oregon-Green fluorescence data was analyzed following division by a factor of
1000 and root arcsine transformation with a one-way RM ANOVA (Tukey test) to compare the
means of normoxic controls and treatments within treatment groups. Results are expressed as
means and standard error of the means (S.E.M.). Significance was determined at p < 0.05.
42
CHAPTER 3: RESULTS
3.1 Pre-current recording evaluations
3.1.1 Anoxic conditions are produced within the bathing chamber
To confirm the recording chamber was anoxic during perfusion with N2/CO2 - bubbled
saline, an O2 electrode was used to measure the change in the partial pressure of oxygen during
the transition from normoxic to anoxic conditions. The solution was bubbled with room air at an
approximate partial pressure of 160 (~21% O2) mmHg, and after transition to aCSF bubbled
with 95% N2/5% CO2 the pressure fell to 0.6 mmHg (~0% O2) after 10 minutes [Figure 3.1].
This value was at the limits of detection of the O2 electrode and was not different from the level
measured in the N2/CO2 - bubbled bottle.
43
Figure 3.1: Change in oxygen tension during normoxic to anoxic transition.
Sample trace representing oxygen tension over the course of 16 minutes in the recording
chamber. The switch to anoxic conditions was made at t = 5 min and the return to normoxia was
made at t= 11 min. Complete anoxic conditions were met after approximately 10 minutes.
3.1.2 All recordings were conducted within pyramidal cells
Pyramidal neurons were ideal for this investigation due to their abundance within the
turtle cortex and their use of glutamate as their primary excitatory neurotransmitter. Previous
publications within the turtle cortex have primarily conducted experiments within these cell
types. To confirm that all neurons tested were pyramidal cells the adaptation pattern and
amplitudes of action-potential firing of all cells were tested previous to NMDAR/AMPAR
current recordings. In order to generate the necessary electrophysiological traces for comparison
cells were current clamped and then current injected for 450 ms to generate a 30 mV
depolarizing change in membrane potential and action potential firing. Cells that displayed
action potentials exhibiting marked frequency adaptation during a sustained injection of current
44
about firing threshold, as well as firing two discrete sizes of action potential were classified as
pyramidal cells. Cells that exhibited no spike frequency adaptation, uniform spike amplitudes,
and a short but large undershoot of membrane potential following action potentials were
classified as stellate cells. The difference between cell types was distinct when traces were
compared [Figure 3.2].
Figure 3.2: Spike frequency adaptation in pyramidal and stellate neurons.
A) Response of a representative pyramidal neuron to a suprathreshold pulse of current. B)
Response of a representative stellate neuron to a suprathreshold pulse of current.
3.2 Whole-cell NMDAR current experiments
3.2.1 Normoxic NMDAR currents are stable throughout the time course
The 90 minute experimental time frame utilized for all NMDAR pharmacological
experiments was based on previous experiments by our laboratory within the turtle cortex. To
confirm that currents remained constant and changes brought on by pharmacological
45
manipulation were not artifacts, normoxic NMDAR current controls were conducted for the
entirety of the experimental time course. To ensure recordings were stable throughout the time
course, cells that experienced a greater than 20mV change in membrane potential from t=0 min
were discarded. Normalized NMDAR peak currents did not change after 10, 30, 50, 70 and 90
minutes of normoxic saline perfusion (93.83 ± 8.00% [n=8], 97.55 ± 9.40% [n=8], 98.77 ±8.56
[n=8], 95.52 ± 9.67% [n=8] and 98.16 ± 10.23% [n=4], respectively) [Figure 3.3].
3.2.2 Normoxic NMDAR current amplitudes are reduced during anoxia
NMDAR whole cell currents in turtle cortical pyramidal neurons undergo a large
approximately 50% reduction in amplitude during anoxia [127, 129, 149]. In order to confirm
that these decreases could be replicated and generate comparisons for pharmacological
manipulations anoxic conditions were induced and evaluated. Following 20 and 40 minutes of
anoxia NMDAR peak current amplitudes were reduced comparable to what has been previously
documented (61.93 ± 8.57% [n=7, p= 0.004] and 58.58 ± 8.97% [n=7, p= 0.008], respectively).
The effects were mostly reversed after 20 minutes of normoxic reperfusion (82.72 ± 12.16%
[n=6]) and completely reversed after 40 minutes of normoxic reperfusion (98.17 ± 4.21% [n=5])
[Figure 3.3].
46
Figure 3.3: Effect of anoxia on whole cell NMDAR currents.
A, Percentage normalized NMDAR peak current amplitudes after 20 min (t=30, blue bars) and
40 min (t=50, red bars) of treatment followed by 20 min (t=70, green bars) and 40 min (t= 90,
purple bars) of washout. Dashed lines represent normoxic baseline controls taken at t=10 min
(red) and 40 min anoxic treatment value (blue). Data are expressed as means ± S.E.M. Asterisks
(*) indicate data significantly different from the paired normoxic control (p < 0.05). B-C, Paired
sample NMDAR currents of the initial baseline recording (t= 10 min) and following 20 min of
the indicated experimental treatment (t= 30 min).
47
3.2.3 NMDAR current amplitudes are increased by ROS scavenging
In order to test the hypothesis that decreases in intracellular ROS concentrations ([ROS]i)
during anoxia serve to trigger decreases in whole cell NMDAR currents, the ROS scavengers
MPG and NAC were administered separately under normoxic conditions. Following 20 and 40
minutes of scavenging, large increases in NMDAR peak current amplitudes were observed for
MPG (201.42 ± 8.25% [n=6, p< 0.001] and 195.06 ± 16.31% [n=4, p< 0.001], respectively) and
NAC (192.46 ± 19.86 % [n=6, p= 0.046] and 208.93 ± 37.32% [n=4 p= 0.035], respectively).
The effects of these increases often resulted in hyperactivity, depolarization and the loss of the
patch. Within the recordings that were maintained, normoxic reperfusion for 20 and 40 minutes
failed to reverse drug effects from MPG (182.84 ± 16.32% [n=3, p< 0.001] and 186.67 ± 5.33%
[n=3, p< 0.001], respectively) and only mildly reversed the effects of NAC (168.35 ± 26.05%
[n=3] and 139.55 ± 24.74% [n=3], respectively) [Figure 3.4]. In only one recording for both
experiments were the effects completely reversed after a period of 110 minutes for MPG and 120
minutes for NAC [Data Not Shown].
48
Figure 3.4: Effect of ROS scavenging on whole cell NMDAR currents.
A, Percentage normalized NMDAR peak current amplitudes after 20 min (t=30, blue bars) and
40 min (t=50, red bars) of treatment followed by 20 min (t=70, green bars) and 40 min (t= 90,
purple bars) of washout. Dashed lines represent normoxic baseline controls taken at t=10 min
(red) and 40 min anoxic treatment value (blue). Data are expressed as means ± S.E.M. Asterisks
(*) indicate data significantly different from the paired normoxic control (p < 0.05). B-C, Paired
sample NMDAR currents of the initial baseline recording (t= 10 min) and following 20 min of
the indicated experimental treatment (t= 30 min). Abbreviations: N-2-mercaptopropionyl glycine
(MPG), n-acetylcysteine (NAC).
49
3.2.4 NMDAR current amplitudes are decreased by H2O2 addition
In order to evaluate the effect of increases in [ROS]i on receptor activity the primary
mitochondrial ROS product H2O2 was administered under normoxic conditions. H2O2
administration was limited to 5 minutes prior to recordings in order to prevent/limit H2O2
toxicity. At 20 and 40 minutes of the treatment regimen significant decreases in NMDAR peak
current amplitudes were observed (80.24 ± 3.11% [n=9, p= 0.023] and 78.42 ± 4.43% [n=5, p=
0.036], respectively). The effects were completely reversed after reperfusion for 20 (93.23 ±
7.11% [n=4]) and 40 minutes (99.26 ± 7.78% [n=4]) [Figure 3.5].
3.2.5 Blocking mitochondrial ROS production increases NMDAR currents
In order to determine if the increases in NMDAR activity due to decreases in [ROS]i are
from a mitochondrial or non-mitochondrial source the mitochondrial complex I inhibitor
rotenone was used to block mitochondrial ROS production under normoxic conditions. After 20
and 40 minutes of rotenone treatment increases in NMDAR peak current amplitudes were
observed (216.57 ± 30.14% [n=5, p= 0.017] and 232.26 ± 28.47% [n=4, p= 0.01], respectively).
The effects were reversed after 20 and 40 minutes of reperfusion (100.08 ± 2.89% [n=3] and
104.56 ± 3.44% [n=2], respectively) [Figure 3.5].
50
Figure 3.5: Effect of H2O2 manipulation on whole cell NMDAR currents.
A, Percentage normalized NMDAR peak current amplitudes after 20 min (t=30, blue bars) and
40 min (t=50, red bars) of treatment followed by 20 min (t=70, green bars) and 40 min (t= 90,
purple bars) of washout. Dashed lines represent normoxic baseline controls taken at t=10 min
(red) and 40 min anoxic treatment value (blue). Data are expressed as means ± S.E.M. Asterisks
(*) indicate data significantly different from the paired normoxic control (p < 0.05). B-C, Paired
sample NMDAR currents of the initial baseline recording (t= 10 min) and following 20 min of
the indicated experimental treatment (t= 30 min). Abbreviations: hydrogen peroxide (H2O2).
51
3.2.6 Anoxia attenuated NMDAR currents are not reversed by ROS addition
In order to test if the effects of anoxia are reversed by the re-introduction of ROS, H2O2
was added after anoxic reductions of NMDAR were measured. As a result the time frame was
modified. Decreases in NMDAR peak current amplitudes were confirmed after 20 and 40
minutes of anoxia (69.62± 3.45% [n=5, p= 0.031] and 66.56 ± 5.96% [n=5, p= 0.013],
respectively). Anoxic conditions were maintained and H2O2 was administered over a further 40
minute period. H2O2 administration did not reverse anoxia induced changes after 20 or 40
minutes (58.60 ± 6.73% [n=5, p= 0.001] and 54.36 ± 5.57% [n=5, p< 0.001], respectively). The
decreases produced from H2O2 administration were not statistically significant from those
produced by anoxia alone [p >0.05]. Normoxic reperfusion only reversed the effects of anoxia
and H2O2 mildly after 20 minutes (59.00 ± 11.48% [n=3, p= 0.006]) and moderately after 40
minutes (75.36 ± 10.10% [n=3]) [Figure 3.6].
52
Figure 3.6: Effect of H2O2 addition on anoxia-attenuated whole cell NMDAR currents.
A, Normalized NMDAR peak current amplitudes after 20 and 40 min of anoxia followed by 20
and 40 min of H2O2 treatment and concluded with 20 and 40 min of washout (n=6). Data are
expressed as means ± S.E.M. Asterisks (*) indicate data significantly different from the paired
normoxic control (p < 0.05). B, paired sample NMDAR currents of a 10 min control and 20 min
of anoxia (t= 30 min), and 20 min of H2O2 + anoxia (t= 70 min). Abbreviations: hydrogen
peroxide (H2O2).
53
3.3 Whole-cell AMPAR current experiments
3.3.1 Normoxic AMPAR currents are stable throughout the time course
The time course utilized for AMPAR experiments was the same as that used for NMDAR
experiments. AMPAR currents under normoxic conditions were examined throughout the
experimental time course to confirm that current amplitudes remained stable. Normalized
AMPAR peak current amplitudes did not change after 10, 30, 50, 70 and 90 minutes of normoxic
saline perfusion (99.85 ± 4.40% [n=6], 96.92 ± 7.10% [n=6], 97.15 ± 7.01 [n=6], 104.94 ±
2.72% [n=4], 105.95 ± 0.78% [n=4] respectively) [Figure 3.7].
3.2.2 Normoxic AMPAR current amplitudes are reduced during anoxia
Like NMDARs, AMPARs also undergo large decreases in current amplitudes under
anoxic conditions. In order to generate anoxic decreases for comparisons to pharmacological
conditions anoxia was induced and currents were induced and evaluated within the established
time frame. Following 20 and 40 minutes of anoxia AMPAR peak current amplitudes were
reduced to a comparable degree of to those previously published (69.95 ± 5.69% [n=5, p= 0.006]
and 55.16 ± 6.04% [n=5, p< 0.001], respectively) [125, 234]. The effects were reversed after 20
and 40 minutes of normoxic reperfusion (98.76 ± 4.86% [n=4] and 101.32 ± 5.19% [n=4]
respectively) [Figure 3.7].
54
Figure 3.7: Effect of anoxia on whole cell AMPAR currents.
A, Percentage normalized AMPAR peak current amplitudes after 20 min (t=30, blue bars) and 40
min (t=50, red bars) of treatment followed by 20 min (t=70, green bars) and 40 min (t= 90,
purple bars) of washout. Dashed lines represent normoxic baseline controls taken at t=10 min
(red) and 40 min anoxic treatment value (blue). Data are expressed as means ± S.E.M. Asterisks
(*) indicate data significantly different from the paired normoxic control (p < 0.05). B-C, Paired
sample AMPAR currents of the initial baseline recording (t= 10 min) and following 20 min of
the indicated experimental treatment (t= 30 min).
55
3.3.3 AMPAR current amplitudes are unaffected by ROS scavenging
In order to test the hypothesis that decreased [ROS]i has an inhibitory effect on AMPAR
current amplitudes the ROS scavengers MPG and NAC were administered separately under
normoxic conditions. No changes in AMPAR peak current amplitude were found after 20 or 40
minutes for MPG (105.40 ± 4.61% [n=6] and 94.71 ± 7.30% [n=4], respectively) or NAC
(100.48 ± 3.21 % [n=7] and 103.48 ± 2.80 % [n=6], respectively). Currents remained unchanged
through 20 and 40 minutes of normoxic reperfusion for MPG (95.26 ± 3.29 % [n=4] and 97.81 ±
12.60 % [n=3]) and NAC (98.62 ± 2.81% [n= 4] and 97.04 ± 3.86 % [n=3]) [Figure 3.8].
3.3.4 AMPAR current amplitudes are unaffected by H2O2 addition
In order to evaluate if increases in [ROS] bring about changes in AMPAR currents the
primary mitochondrial ROS product H2O2 was administered under normoxic conditions.
Normoxic AMPAR peak current amplitudes were not altered by 20 or 40 minutes of H2O2
addition (100.00 ± 3.35% [n=11] and 99.19 ± 2.44% [n=7], respectively). AMPAR current
amplitudes remained unchanged through 20 and 40 minutes of normoxic reperfusion (106.33 ±
3.91% [n=5] and 100.00 ± 2.95% [n=3], respectively) [Figure 3.8].
56
Figure 3.8: Effect of ROS manipulation on whole cell AMPAR currents.
A, Percentage normalized AMPAR peak current amplitudes after 20 min (t=30, blue bars) and 40
min (t=50, red bars) of treatment followed by 20 min (t=70, green bars) and 40 min (t= 90,
purple bars) of washout. Dashed lines represent normoxic baseline controls taken at t=10 min
(red) and 40 min anoxic treatment value (blue). Data are expressed as means ± S.E.M. Asterisks
(*) indicate data significantly different from the paired normoxic control (p < 0.05). B-C, Paired
sample AMPAR currents of the initial baseline recording (t= 10 min) and following 20 min of
the indicated experimental treatment (t= 30 min). Abbreviations: n-acetylcysteine (NAC), N-2-
mercaptopropionyl glycine (MPG), hydrogen peroxide (H2O2).
57
3.4 Oregon green intracellular calcium assessments
3.4.1 Excitation scans confirmed dye loading in turtle brain sheets
Dye-loading within the cells was confirmed by running excitation scans from 450 nm to
550 nm prior to all recordings. Peak excitation was observed around 488 nm in all dye loaded
cell but was absent in non-dye loaded cells [Figure 3.8].
Figure 3.9: Oregon Green excitation scans.
A-B, sample excitation scans from 450 nm to 550 nm in Oregon Green dye-loaded (A) and non-
dye-loaded (B) neurons.
3.4.2 Normoxic calcium concentrations are stable throughout the time course
The time course for fluorescent experiments was different from that utilized for whole
cell recordings. Photo bleaching occurs after extended periods of exposure that can cause a loss
of the fluorescent signal. The experimental time frame consisted of a 10 minute normoxic time
58
period that allowed for the fluorescent signal to stabilize, followed by 10 minutes of
pharmacological treatment, and 10 minutes of normoxic recovery. To confirm that the Oregon
Green fluorescence was stable throughout the experimental time course traces were collected
under normoxic conditions through a 30 minute time period. Under normoxic conditions Oregon
Green fluorescence did not change significantly with respect to baseline (0.61 ± 0.77% [n= 6])
and decreased at a steady rate due to photo-bleaching and/or dye exocytosis [Figure 3.10].
3.4.3 Intracellular calcium concentrations are increased by anoxia
The entrance into anoxia is associated with a mild increase in intracellular calcium [126,
128, 149]. These experiments were replicated to confirm previous findings and generate rates of
change for comparison to pharmacological treatments. Anoxic administration mildly increased
Oregon Green fluorescence (16.94 ± 3.29% [n= 6, p< 0.001]) [Figure 3.10].
3.4.4 Intracellular calcium concentrations are unaffected by ROS scavenging
As previously discussed intracellular calcium is increased as a result of anoxia. In order
to determine if decreases in ROS served to bring about the mild increases in calcium associated
with anoxia the ROS scavengers MPG and NAC were separately applied during an experimental
time course. Oregon green fluorescence was not significantly altered with respect to the baseline
by MPG or NAC addition (0.56 ± 0.64% [n= 6] and 0.96 ± 1.28% [n= 6]) [Figure 3.10].
59
3.4.5 Intracellular calcium concentrations are unaffected by H2O2 addition
H2O2 was bulk perfused onto the slice as drip perfusion proved unreliable and affected
image stability. H2O2 bulk concentrations were altered to be comparable to those brought on by
drip perfusion into the recording bath during electrophysiological recordings (drip= 50 uM,
bulk= 5 uM). H2O2 addition did not significantly alter fluorescence in respect to baseline (0.84 ±
1.39% [n= 6]) [Figure 3.10].
60
Figure 3.10: Effect of oxygen and ROS manipulation on Oregon Green fluorescence.
Percentage normalized changes in Oregon Green fluorescence post treatment. Data are expressed
as means ± S.E.M. Asterisks (*) indicate data significantly different from the paired normoxic
control (p < 0.05). Abbreviations: Arbitrary Fluorescence Units (AFU), N-2-mercaptopropionyl
glycine (MPG), n-acetylcysteine (NAC), Hydrogen peroxide (H2O2).
61
Figure 3.11: Sample Oregon Green fluorescence traces.
Sample traces of Oregon Green fluorescence from neurons treated as indicated. Changes in
fluorescence were calculated between two parallel tangents, as illustrated in B. C, Illustration of
the method by which Fmin and Fmax were determined. Abbreviations: Arbitrary Fluorescence
Units (AFU), N-2-mercaptopropionyl glycine (MPG), n-acetylcysteine (NAC), Hydrogen
peroxide (H2O2).
62
3.5 Action potential frequency assessments in spontaneously active pyramidal cells
3.5.1 Action potential frequency is reduced by ROS scavenging
As previously discussed entrance into anoxia is associated with an increase in
GABAergic PSCs that serve to bring about SA and inhibit/decrease neuronal firing [115-117].
This is most clearly seen within neurons that exhibit spontaneous activity and fire continuously
at regular intervals. Recordings from these cells within turtle cortical sheets demonstrate a 70%
reduction in action potential frequency (APf) with the transition to anoxia [112]. ROS scavenging
with MPG and NAC has also been shown to bring about increases in GABAergic PSCs similar
to anoxia [231]. Therefore, in order to determine if decreases in ROS result in SA similar to that
seen during anoxia, spontaneously active pyramidal cells were treated with MPG during APf
recordings. APf decreased by approximately 97% after 18 minutes of MPG treatment (0.04 ±
0.04 [n=4, p< 0.001]) that was reversed and increased after 13 minutes of normoxic reperfusion
(2.33 ± 0.44 [n=4, p= 0.016]) [Figure 3.11].
63
Figure 3.12: Changes in spontaneous activity in response to ROS scavenging.
A, Summary of APf expressed as fold change relative to pretreatment baseline. Data are
expressed as means ± S.E.M. Dashed line represents normoxic baseline APf. Asterisks (*)
indicate data significantly different from the paired normoxic control (p < 0.05). B, Raw
spontaneous activity sample trace. Abbreviations: N-2-mercaptopropionyl glycine (MPG).
64
CHAPTER 4: DISCUSSION
4.1 Control normoxic and anoxic NMDA and AMPA currents
In agreement with previous findings normoxic NMDA and AMPA administration
produced consistent repeatable currents throughout the 90 minute time course of the experiment.
The introduction of anoxia produced a decrease of approximately 50% in both NMDA and
AMPA currents within 20 minutes consistent with previous reports [126, 149, 234]. This served
to further confirm neurological stability and pharmacological validity.
4.2 Pharmacological manipulation of ROS
4.2.1 The NMDAR modulatory redox site in turtle cortical slices
Cellular ROS scavenging with both MPG and NAC produced massive increases in
NMDAR whole cell currents contrary to the anoxia mediated decrease typically observed in the
western painted turtle cortex. Inversely, the addition of H2O2 under normoxic conditions
decreased NMDAR whole cell currents. Although this indicates that decreases in ROS are not
responsible for NMDAR attenuation during anoxia, the findings are consistent with what has
been documented within neural tissue from other vertebrates and may reveal a conserved
mechanism of NMDAR control during normoxia [190-198].
As discussed above, pharmacological increases in ROS or the addition of oxidizing
agents has been documented to decrease NMDAR activity within tissue preparations from
65
mammals, amphibians, and birds. Pharmacological decreases in ROS or the addition of reducing
agents has also been shown to produce increases in NMDAR activity within these same groups.
These changes have been previously attributed to alterations in the oxidation/reduction state of
extracellular cysteine residues found within the NMDAR GluN1 (Cys744, Cys798, Cys 79 and
Cys 308), GluN2A (Cys 87, Cys 320 and Cys 399), and/or GluN2B subunits (Cys not yet
localized) that collectively make up NMDAR redox sites [187-189]. The changes documented
within cortical slices from the turtle during ROS manipulation may also be attributed to
alterations in the NMDAR redox site. Specific regions within NMDAR subunits have been
documented to be highly conserved between vertebrate species. Comparisons of the Brown ghost
knifefish GluN1 amino acid structure found an 88% sequence similarity to the amino acid
structures in frog, duck, rat and human [235]. Similarly, the amino acid structures of the four
NR2 subtypes possess high levels of conservation between species within sequences coding the
lining of the ion pore, glutamate binding pockets, and C-terminal region [119, 236]. The redox
modulatory sites on the GluN1 and GluN2 subunits may be similarly conserved between species.
Increases and decreases in NMDAR current amplitude between ROS scavenging and
addition were not uniform. The decreases in receptor currents with H2O2 addition were only
~20% compared to the ~100% increase with ROS scavenging. This is consistent with what has
been seen in other vertebrates where the addition of reducing agents produced increases that
were 6-11 times greater than decreases produced through oxidation [190, 196, 199]. Our findings
suggest that the turtle NMDAR redox modulatory site is maintained closer to a fully oxidized
than fully reduced form under normoxic conditions, in agreement with what has been
documented in other vertebrate species. This provides support for the Cys residues being found
on the extracellular portion of the receptors, where the environment is predominantly oxidized
66
[168]. It may also indicate a maintained ROS presence in close proximity to the receptors and
redox sites.
The level of recovery from treatment varied drastically between ROS scavenging and
addition. Although recovery from H2O2 was achieved within the 40 minute reperfusion period,
MPG/NAC recovery was not and cells often depolarized and did not recover preventing the
exploration of a longer recovery period. This is consistent with what has been documented in
other vertebrates where decreases in NMDAR activity induced through the application of
oxidizing agents were consistently reversed with washout, but increases in NMDAR activity
induced through reductive agents were maintained. The effects of reducing agents have only
been documented to be reliably recovered to baseline through the addition of oxidizing agents
[196, 237]. The reasoning behind the lack of recovery from scavenging/reducing agents may be
explained by slow cellular ROS production and slow degradation of scavenging/reducing agents.
As discussed above, ROS are produced slowly in the cell (only 2-3% of all mitochondrial O2
consumed) maintained at low levels and may move slowly across the extracellular membrane
[168]. It may take a long time for baseline levels to be recovered following ROS scavenging
and/or re-oxidize cysteine residues affected by scavenging agents. Furthermore, scavenging and
reducing agents may take a long time to be removed from the cell or to be degraded through
oxidation, further delaying the re-establishment of baseline ROS levels and NMDAR activity.
The addition of oxidizing agents would help in re-oxidization of redox sites and the degradation
of scavenging/reducing agents, potentially explaining why this tactic was successful in reversing
the effects of reducing agents on NMDAR activity. Cells are likely to be better equipped to deal
with ROS increases then their absence as the complete removal of ROS is unlikely for most
aerobic life.
67
The risks of NMDAR over activation and resulting ECD are common to all vertebrates
and NMDAR activity must be tightly regulated to prevent it. Ca2+ entry must be maintained
within a range where intake concentrations are great enough to signal downstream changes but
small enough to be taken up by the mitochondria and ER so that they don’t bring on toxic
effects. Most cellular activities are maintained within a desired range via negative feedback
systems allowing products generated by specific processes to regulate the actions of the
processes themselves. NMDAR Ca2+ dependent ROS production via nNOS or mitochondrial
activation may be a mechanism through which NMDAR activity is regulated among vertebrates.
4.2.2 Mitochondrial produced ROS in NMDAR redox modulation
The mitochondrial complex I inhibitor rotenone decreases ROS production in various
vertebrate tissues including neuronal tissue [157, 200-202]. Rotenone addition in turtle cortical
slices produced increases in NMDAR currents comparable to what was seen with MPG/NAC
ROS scavenging. This suggests that changes in mitochondrial ROS production affect NMDAR
activity. Full recovery from rotenone treatment was achieved within the reperfusion period
unlike what was seen with scavenging agents. This supports the concept that the recovery delay
seen during scavenging is the result of agents taking a long period to be degraded and removed
from the cell.
The finding that mitochondrial ROS production affects NMDAR activity does not rule
out a role for NO generated from nNOS. During whole cell patch experiments, TTX application
and voltage clamping would have prevented natural glutamate release and blocked NO
68
production except during NMDA addition. Because nNOS activity directly linked to NMDAR
activation and NO can oxidize cysteine residues in a similar manner to H2O2, it is very likely that
both NO and mitochondrial ROS play a role in NMDAR control. In studies within other
vertebrates the addition of nNOS inhibitors increases baseline NMDAR currents while artificial
NO generators bring about receptor attenuation [241, 242]. However, nNOS inhibitors do not
prevent mitochondrial ROS increases triggered by NMDAR activation, suggesting that the
mechanisms function independently of each other [238, 239]. Further investigation is necessary
to determine the role and contribution of NO in NMDAR redox control.
Mitochondrial ROS generation is increased by NMDAR activation within mammalian
neural cultures. NMDA mediated ROS increases were not prevented by nNOS inhibitors but
were blocked by the removal of extracellular calcium, rotenone/ other mitochondrial uncouplers,
and Ca2+ uniporter inhibitors [238, 239]. This suggests that increased mitochondrial ROS is
generated as a result of NMDAR specific Ca2+ influx and increased mitochondrial Ca2+ uptake. If
Ca2+ influx were to exceed the capabilities of the storage systems like mitochondrial uptake, then
Ca2+ would begin to spread through the cell causing toxicity. It is therefore not unreasonable that
mitochondrial uptake might influence NMDAR activity.
Mitochondrial calcium uptake is primarily accomplished through the actions of the
mitochondrial calcium uniporter (MCU) located on the inner mitochondrial membrane (IMM).
The outer mitochondrial membrane is permeable to most large ions (<6000 DA) allowing for
unaided Ca2+ entry. The IMM is not as permeable and transporters are required to allow for Ca2+
movement [240]. The MCU becomes active in response to large concentrations of localized Ca2+
known as microdomains that in excitable cells need to be 500-600 nM in order to generate a
69
response [241]. Activity is driven by the large negative potential of the mitochondrial matrix
established by the actions of the respiratory chain. Electrophoretic uptake of Ca2+ into the
mitochondrial matrix through the uniporter partially dissipates the electrochemical gradient [242,
243]. Compensation is accomplished via Ca2+ induced increases in electron transport chain
activity that pump H+ out of the matrix helping to re-establish the electrochemical gradient and
also bringing about increases in ATP production [242]. Enhancements in ROS production can be
accomplished through the increases in ETC activity, generating ROS as a by-product. There is
also some evidence to suggest that matrix Ca2+ can partially dislocate cytochrome c from the
IMM thereby slowing down the electron transfer from complex III to complex IV and enhancing
ROS generation at the Q cycle [239, 244]. Ca2+ within the matrix is transient and the actions of
multiple Ca2+ efflux mechanisms, the primary one in excitable tissues being the Na+/Ca2+
exchanger, work in concert to move Ca2+ back into the cytosol [242, 245]. Storage within the
mitochondrial matrix can be quite high and has been documented in excess of 100 uM [246].
However, the capacity is not unlimited and if Ca2+ overload occurs and intake exceeds efflux
mitochondrial membrane potentials can collapse leading to excessive Ca2+ efflux and the
triggering of apoptosis [247, 248].
4.2.3 A potential mechanism for ROS mediated NMDAR control
For NMDAR activity to be regulated via oxidation/reduction ROS synthesis must not
only be tied directly to receptor activity, ROS production must be accomplished within close
proximity to the channel. The redox site will be maintained in a preferentially oxidized state by
likely existing within the extracellular environment, but changes in intracellular ROS changes
will only be effective signals if ROS production occurs within the vicinity of the NMDAR. As
70
discussed above, this may involve NO produced by nNOS that is tethered to NMDAR via PSD-
95. However, for mitochondrial ROS generation to be effective in modulating activity as shown
the rotenone experiments, mitochondrial localization must also take place.
ROS are produced from the mitochondria in relatively small quantities (2-3% of O2
consumed) that are maintained at low levels through the actions of cellular antioxidant defence
systems. This coupled with high reactivity leads to small and inconsistent mitochondrial ROS
levels that exist throughout the cytoplasm limiting their signalling ability to short distances [163,
249]. Furthermore, in order for ROS to affect extracellular redox sites ROS must pass into the
extracellular environment through diffusion or WTRs where it can then be further degraded by
extracellular antioxidants [168-171]. Therefore in order for ROS control to be effective and
consistent, target cysteine residues must be in close proximity to the mitochondria. NMDARs are
primarily located within post synaptic densities of the excitatory synapse and mitochondria are
similarly packed into these synapses in order to provide the ATP necessary for synaptic activities
[250-252]. These synaptic regions are high in membrane ion channels that transport water
(WTRs) potentially allowing for an efficient flux of generated ROS across the membrane [172].
Recent investigation in mammalian neurons found NMDAR specific Ca2+ entry was far more
rapidly taken up by the mitochondria than Ca2+ entry through other channels [253]. This tight
coupling of mitochondrial Ca2+ uptake to NMDAR activation is suggested to be the result of the
organelles and receptors being maintained within close proximity to each other in order to better
control cytosolic Ca2+ levels [253, 254]. There is a considerable body of work in non-excitable
cells that suggests that rapid mitochondrial Ca2+ uptake requires high levels of Ca2+ to be present
in close proximity (~2-5 uM) to the mitochondria and that the endoplasmic reticulum and
mitochondria are positioned so that excess calcium leaving the ER is high enough to signal this
71
action and be quickly sequestered [255-259]. Within excitable cells like neurons this high Ca2+
concentration has been shown not to be required to activated rapid uptake, but localized cytosolic
Ca2+ levels of above 500–600nM are still required to stimulate influx supporting the tight
coupling concept [241].
Given my experimental findings and those of similar experiments conducted in other
organisms oxidation/reduction of the NMDAR redox site appears to have a similar effect across
multiple vertebrate species suggesting a common mechanism of control. Using information
collected from other studies I submit the following potential mechanism for mitochondrial ROS
mediated NMDAR control under normoxic conditions. I do this in order to try and better explain
how ROS control might be accomplished so that that I can better explain later how anoxia
mediated mechanisms may override these processes. The proposed mechanism for is as follows:
NMDAR activation causes an influx of extracellular Ca2+ into the cell that serves to
signal changes downstream changes via activation of receptor localized Ca2+-dependent
enzymes. Excessive calcium that does not become bound is rapidly taken up by localized
mitochondria to prevent it from spreading out through the cell and undergoing unwanted
reactions. Ca2+ freely flows across the permeable mitochondrial outer membrane and is then
taken up into the matrix via the action of the MCU. Ca2+ entrance stimulates increases in ETC
activity to compensate for the inward ion flow, generating increased levels of ROS as result. The
ROS produced will rapidly move out of the mitochondria and into the cytosol. ROS may then
move across the extracellular membrane and into the extracellular environment by simple
diffusion or through WTRs where it can interact with NMDAR redox sites in close proximity,
reversibly oxidizing the modulatory redox site and decreasing the receptors open time in
72
response to stimulus. The cumulative action of multiple mitochondria undergoing these
processes will serve to generate a cloud or microdomain of ROS that will help maintain
NMDARs primarily in an oxidized state and resist the actions of antioxidant enzymes. Increases
in NMDAR activation will bring on increases in Ca2+ influx and subsequent ROS generation that
will increase receptor oxidation and decrease sensitivity. Similarly decreases in NMDAR
activation will lead to decreases in ROS allowing for enzymes like GSH/L-Cys to reduce a
greater portion of receptors and increase sensitivity. Through this negative feedback mechanism
Ca2+ influx can be maintained within a range that brings about necessary changes without
becoming high enough to overtake mitochondrial uptake mechanisms and trigger overload
[Figure 4.1].
Although this mechanism will maintain NMDAR activity within a normal range during
normoxia it will not function under anoxic conditions as mitochondrial ROS generation requires
O2. The loss of cellular oxygen will halt oxidative phosphorylation and allow for cellular ROS
levels to be quickly depleted. The method of maintained NMDAR inhibition will quickly be lost
as the receptors are rapidly reduced leading to hyper excitability. Activation of the receptors will
trigger a massive influx of Ca2+ bringing about excitotoxic cell death. In order for the western
painted turtle to survive prolonged anoxia another mechanisms must serve to override ROS
control and trigger the decreases in NMDAR activity documented to occur.
73
Figure 4.1: Diagrammatic representation of the proposed mechanism for ROS mediated
NMDAR control under normoxic conditions.
74
4.2.4 AMPAR redox insensitivity
AMPAR currents were unaffected by MPG, NAC or H2O2 consistent with what is seen in
mammalian tissue with oxidizing/reducing agents suggesting that the mechanisms for AMPAR
control in respect to ROS changes are conserved between species [196-198, 203].
The reasoning behind why there exists a mechanism for ROS mediated NMDAR control
but not AMPAR may be a direct result of how oxidation of the NMDAR leads to inhibition of its
activity. AMPARs activation produces excitatory postsynaptic potentials (EPSPs) that cause
neuronal depolarization and the removal of Mg2+ from the pore region of the NMDAR allowing
activation and Ca2+ influx [260-263]. It is through this indirect mechanism of action that
AMPARs exhibit control over intracellular Ca2+ influx. Changes to the modulatory redox site
may override AMPAR control by affecting the efficiency to which NMDARs can maintain Mg2+
within the pore region. Treatment of mammalian neural tissue with rotenone resulted in increases
in NMDAR activity as previously discussed, but also documented that NMDAR currents lost the
characteristic region of negative-slope conductance that is normally produced by voltage-
dependent block by Mg2+. NMDAR increases were mimicked by low extracellular Mg2+ and
rotenone induced increases were antagonized by a high levels of Mg2+. Rotenone addition had no
effect on AMPAR currents [203, 206]. This suggests that oxidation/reduction alters the ability of
the NMDAR to maintain Mg2+ within the pore region and overrides/circumvents AMPAR
control.
75
The lack of AMPAR down regulation in response to decreased [ROS]i further supports
the concept that ROS changes are not responsible for NMDAR/AMPAR current decreases in the
anoxic Western painted turtle brain and that a secondary mechanism must be at work.
4.2.5 Anoxia-mediated block of NMDAR redox control
Anoxic decreases in NMDAR currents were not reversed by the addition of H2O2
confirming that ROS changes are not responsible for anoxia mediated changes. H2O2 addition to
anoxia-attenuated NMDAR currents appeared to further reduce whole cell currents (~10%), but
it is not clear if this decrease was simply the result of extended anoxia. Average currents after 90
minutes of anoxia with H2O2 reached ~54% of normoxic levels that was not significantly
different from that seen after 30 minutes of anoxia in control experiments (~58%) or previous
publications (~52%) [149]. H2O2 addition alone produced decreases after 40 min of
approximately 22% from normoxic baseline compared to addition after anoxia that produced
after 40 min only an approximate 12% change. This would seem to support the concept that the
secondary mechanism for triggering anoxic NMDAR activity decreases exists that overrides the
mechanisms of ROS control. However without anoxic controls for a comparable period of time
the assumption cannot be verified. Regardless, the results confirm that another mechanism aside
from ROS decreases is responsible for the induction of NMDAR down regulation. The most
likely candidate for these changes is changes in cellular calcium from an intracellular source.
76
4.3 A role for anoxia-mediated calcium increases in blocking redox control
A mild increase in cytosolic calcium from the mitochondria is seen during anoxia in the
western painted turtle and is necessary to bring about decreases in NMDAR and AMPAR whole
cell currents [125, 264]. Investigations using Oregon Green Ca2+ fluorescence confirmed an
anoxic increase in intracellular calcium similar to what has been previously documented (~15%)
[149]. However, the addition of MPG, NAC and H2O2 failed to elicit any change in the
fluorescent Ca2+ signal demonstrating that ROS changes do not play a role in the anoxia-
mediated increase in cytosolic Ca2+. This serves to confirm that a secondary mechanism for
anoxia-mediated changes is at work and suggests that mitochondrial release of Ca2+ is central to
its activity.
Increases in mitochondrial Ca2+ release are associated with activation of the calcium-
binding messenger protein calmodulin (Ca2+/calmodulin). Ca2+/calmodulin is a small Ca2+
binding protein that is highly conserved within eukaryotes, present in high concentrations in the
brain and capable of modulating a number of different enzymes and cellular processes [265,
266]. Within the western painted turtle inhibition of Ca2+/calmodulin has been demonstrated to
prevent decreases in NMDAR activity. Ca2+/calmodulin may also activate the Ca2+/calmodulin-
dependent phosphatase PP2B (calcineurin), whose inhibition has also been shown to inhibit early
anoxia mediated NMDAR downregulation. Interestingly inhibition of PP1 and PP2A was also
shown to inhibit NMDAR decreases suggesting a dependency between both dephosphorylation
and calmodulin interaction [129]. The sites of modification and binding offer means for
NMDAR/AMPAR control that may also serve to override the mechanism of ROS mediated
NMDAR regulation.
77
Calmodulin has been shown to exert effects via binding to two identified sites (CBS1 and
CBS2) the C0 terminus of the GluN1 via destabilization of the NMDARs from the cytoskeleton
[267-269]. It has also been demonstrated that phosphorylation of the GluN1 serine residues
decreases affinity for calmodulin, an effect that can be reversed by the actions of PP1 and PP2A
[150]. This has led to the hypothesis that during anoxia, PP1 and 2A dephosphorylate the serine
residue on the C-terminus of the GluN1 subunit allowing for the binding of Ca2+/calmodulin and
an attenuation of NMDAR function [264]. As discussed above NMDAR redox modulatory sites
have been identified on the GluN1, (Cys744, Cys798, Cys 79 and Cys 308) GluN2A (Cys 87,
Cys 320 and Cys 399) and GluN2B (not yet localized) [187-189]. GluN1 dephosphorylation or
calmodulin binding both offer a potential means to block modulatory redox sites, preventing
oxidation/reduction and overriding ROS control. Furthermore, calcineurin dephosphorylates the
GluN2A subunit (not GluN1) that besides having been shown to contribute to glycine
desensitisation and NMDAR current reduction, offers a means through which modulatory redox
sites on the GluN2A subunit may be blocked [267, 270]. It is also possible that calmodulin
induced NMDAR cytoskeletal dissociation may bring about destabilization the modulatory redox
site leading to a loss of ROS control. AMPAR activity has not been demonstrated to be regulated
through calmouldin binding but has been identified to be controlled by dephosphorylation.
Increases in calcineurin activity dephosphorylate Ser845 on the GluA1 subunit triggering a
decrease in AMPAR function [271-273].
In support of this mechanism, inhibition of intracellular calcium release and subsequent
phosphatase activity has been associated with increases in NMDAR activity under anoxic
conditions. Addition of the calcium chelator BAPTA increased NMDAR whole cell currents by
78
35% after 20 minutes of anoxia. Similarly addition of the PP1/PP2A inhibitor calyculin A
increased NMDAR whole cell currents by 45% after 40 minutes of anoxia. Finally, addition of
the calmodulin inhibitor calmidazolium increased NMDAR whole cell currents by 20% after 40
minutes of anoxia [129].
The mechanism by which calcium is released from the mitochondria has yet to be
established. However recent investigation has suggested that it is the result of mKATP activation
and subsequent mitochondrial permeability transition pore (mPTP) formation. The decreases in
AMPAR and NMDAR activity seen during anoxia are mimicked by mKATP activators and
blocked by mKATP inhibitors [125, 126]. Furthermore, inhibition of mKATP prevents the mild
increase in intracellular Ca2+ characteristic of anoxia, while activation produces Ca2+ increases of
a similar magnitude[126]. mKATP channels are traditionally gated by high levels of ATP under
normoxia. During anoxia ATP production at the level of the mitochondria decreases causing
local ATP levels to be reduced potentially leading to the opening of mKATP channels. Opening
then leads to an influx of K+ that can trigger mPTP formation as a result of mitochondrial
membrane depolarization. The mPTP is a temporary channel that can be formed in the inner
mitochondrial membrane and permit Ca2+ release. The formation of a singular large mPTP often
bring on excessive Ca2+ release and toxic results, but the formation of several small mPTPs has
been shown to bring about safe levels of Ca2+ efflux [274]. Within the western painted turtle, the
addition of an mPTP activator produced decreases in NMDAR currents similar to those seen
during anoxia that were abolished by the addition of a Ca2+ chelator. The addition of the mPTP
activator also produced an increase in Ca2+ fluorescence identical to that seen during anoxia
[149]. The mPTP is naturally formed under conditions of mitochondrial stress such as K+
mediated uncoupling that can trigger mild to moderate membrane depolarization (10-20%) [275].
79
Furthermore, the pore is traditionally unstable and short lived causing it to quickly collapsing
and potentially only produce a mild increase in intracellular calcium [276].
4.4 Roles for redox sensitive GABA release
Entrance into anoxia is associated with a decrease in spontaneous AP frequency of
approximately 70%, which is attributed to increases in GABAR activity [112]. Recently
decreases in ROS were shown to produce increases in GABAergic PSC activity suggesting that
they play a role in GABAergic anoxic signalling [231]. In support of these findings MPG
addition reduced spontaneous AP frequency similar to anoxia although to a greater degree
(~98%). These results serve to confirm that although decreases in ROS during anoxia are not
responsible for NMDAR and AMPAR down regulation they do appear to be involved in the
processes controlling GABAergic activity increases.
Recovery from MPG treatment resulted in a major increase in firing above previous
baseline levels (~74%). This may be the result of NMDAR over excitation and Ca2+ influx,
triggered by ROS decreases as previously discussed. As discussed previously, MPG may take a
long time to be removed from the cell leading to NMDAR currents increases that were not
reversed after 40 minutes of normoxic reperfusion. Therefore, even though GABA mediated
ROS effects were quickly reversed the NMDAR augmentation may have remained and resulted
in excessive Ca2+ influx, increased neuronal firing, and generated the AP frequency increases
seen.
80
Interestingly, increases in GABAergic PSCs will serve to decrease/inhibit postsynaptic
glutamate release preventing NMDAR and AMPAR activation. Through this mode of action
GABA changes in response to low oxygen may serve as a preliminary level of protection,
preventing NMDAR from activating in a reduced/overexcited and triggering ECD long enough
for the mechanisms controlled via mitochondrial Ca2+ release to take effect.
4.5 Potential mechanism of NMDAR/AMPAR control under anoxic conditions
The mechanisms proposed above for control of NMDAR activity through redox changes
will maintain intracellular Ca2+ influx within a normal range under normoxic conditions.
However, during anoxia O2 and ROS depletion will produce a state of NMDAR hyper
excitability that has the potential to produce massive increases in Ca2+ intake. Within the brain
of the western painted turtle ECD is not seen to occur and NMDAR/AMPAR receptors
experience major down regulations in activity. This suggests that a secondary mechanism of
control is activated by the anoxic state that can override the mechanism of ROS/redox control.
Given my and others findings in the brain of the western painted turtle during oxygen
deprivation I propose the following mechanism of anoxia-mediated NMDAR/AMPAR control:
as oxygen concentrations are depleted within neuronal tissue oxidative phosphorylation will
begin to cease. This will result in mitochondrial localized decreases in ATP triggering the
opening of mKATP channels. Influx of K+ through these channels will trigger mild/moderate loss
of the mitochondrial membrane potential and bring about the formation of multiple small
mPTPs. Ca2+ will move rapidly through these channels that do to their instability quickly
collapse. The resulting mild cytosolic Ca2+ increase will bind to calmodulin activating it.
81
Ca2+/calmodulin can then bind directly to the NMDAR GluN1 subunit either with or without the
aid of dephosphorylation via PP1 and PP2A. This binding will serve to inhibit receptor activity
as well as block the GluN1 redox modulatory sites overriding the mechanisms of ROS control.
The activation of calcineurin via calmodulin binding will serve to dephosphorylate the GluN2A
subunit contributing to receptor down regulation and GluN2A modulatory redox site
destabilization. Calcineurin will also dephosphorylate the Ser845 on the GluA1 subunit of the
AMPAR down regulating activity. While these changes are occurring increases in GABAergic
activity triggered by decreasing ROS levels will serve to prevent glutamate release and
subsequent NMDAR/AMPAR activation [Figure 4.2].
82
Figure 4.2: Diagrammatic representation of the proposed mechanism for anoxia mediated
NMDAR/AMPAR downregulation.
83
4.6 Concluding remarks and directions for future research
These results suggest that changes in oxidation/reduction state of the NMDAR
modulatory redox site lead to increases and decreases in activity respectively, without effect on
AMPAR. This is in agreement with what has been demonstrated within investigations in other
vertebrates. To our knowledge this is the first case where evidence for this mechanism has been
demonstrated in reptiles. Our findings also support the hypothesis that these redox changes are
regulated by natural ROS levels, and in this way receptor function can be conservatively
regulated between vertebrates under normoxic conditions. Our research demonstrated that this
mechanism is not responsible for NMDAR/AMPAR downregulations seen during anoxia within
the western painted turtle, but found evidence to support an alternative mechanism involving
mitochondrial Ca2+ release.
In order for a better determination of the validity of the mechanisms proposed, a number
of questions still need to be answered and different experiments should be conducted. (1) It has
yet to be conclusively established how anoxia-mediated changes override ROS control, though
there is evidence to suggest it occurs through mitochondrial Ca2+ release. The mKATP activator
diazoxide has been previously documented to produce anoxic decreases in NMDAR currents
similar to those seen during anoxia. Diazoxide mediated decreases under normoxia should be
evaluated in conjunction with MPG/NAC in order to determine if scavenging effects are
mitigated. (2) The effects of phosphatases or calmodulin on AMPAR activity have yet to be
evaluated in the western painted turtle brain. Anoxia mediated AMPAR currents should be
evaluated in the presence of inhibitors for PP1, PP2A, calmodulin, and calcineurin in order to
determine if which if any of these groups is responsible for anoxia mediated AMPAR
84
downregulation. (3) The role of NO in NMDAR control has yet to be firmly established. nNOS
inhibitors should be used in the same manner as roteone to determine if their blockade produces
comparable increases to ROS scavenging.
Understanding neurological anoxia tolerance is critical to many of the medical problems
faced within our society today. Stroke represents the third leading cause of death in North
America and with both childhood and adult obesity rates increasing across North America the
number of incidents is predicted to increase [277]. The development of preventative and fast
acting measures, like seen in the overwintering capabilities of the western painted turtle, may
offer a means to save thousands of lives.
85
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