THE ROLE OF CARBONIC ANHYDRASE IN THE MODULATION OF ... · ii The Role of Carbonic Anhydrase in the...
178
THE ROLE OF CARBONIC ANHYDRASE IN THE MODULATION OF CENTRAL RESPIRATORY-RELATED PH/CO 2 CHEMORECEPTOR-STIMULATED BREATHING IN THE LEOPARD FROG (RANA PIPIENS) FOLLOWING CHRONIC HYPOXIA AND CHRONIC HYPERCAPNIA by Kajapiratha Srivaratharajah A thesis submitted in conformity with the requirements for the degree of Master of Science Graduate Department of Cell and Systems Biology (Zoology) University of Toronto © Copyright by Kajapiratha Srivaratharajah (2008)
Transcript of THE ROLE OF CARBONIC ANHYDRASE IN THE MODULATION OF ... · ii The Role of Carbonic Anhydrase in the...
THE ROLE OF CARBONIC ANHYDRASE IN THE MODULATION OF CENTRAL
RESPIRATORY-RELATED PH/CO2 CHEMORECEPTOR-STIMULATED BREATHING IN
THE LEOPARD FROG (RANA PIPIENS) FOLLOWING CHRONIC HYPOXIA AND
CHRONIC HYPERCAPNIA
by
Kajapiratha Srivaratharajah
A thesis submitted in conformity with the requirements
for the degree of Master of Science
Graduate Department of Cell and Systems Biology (Zoology)
University of Toronto
© Copyright by Kajapiratha Srivaratharajah (2008)
ii
The Role of Carbonic Anhydrase in the Modulation of Central Respiratory-Related pH/CO2
Chemoreceptor-Stimulated Breathing in the Leopard Frog (Rana pipiens) Following Chronic
Hypoxia and Chronic Hypercapnia
Kajapiratha Srivaratharajah Master of Science
Graduate Department of Cell and Systems Biology University of Toronto
2008
ABSTRACT
The aim of this thesis was to elucidate the role of carbonic anhydrase (CA) in the modulation of
central pH/CO2-sensitive fictive breathing (measured using in vitro brainstem-spinal cord
preparations) in leopard frogs (Rana pipiens) following exposure to chronic hypercapnia (CHC)
and chronic hypoxia (CH). CHC caused an augmentation in fictive breathing compared to the
controls (normoxic normocapnic). Addition of acetazolamide (ACTZ), a cell-permeant CA
inhibitor, to the superfusate reduced fictive breathing in the controls and abolished the CHC-
induced augmentation of fictive breathing. ACTZ had no effect on preparations taken from frogs
exposed to CH. Addition of bovine CA to the superfusate did not alter fictive breathing in any
group, suggesting that the effects of ACTZ were due to inhibition of intracellular CA. Taken
together, these results indicate that CA is involved in central pH/CO2 chemoreception and the
CHC-induced increase in fictive breathing in the leopard frog.
iii
ACKNOWLEDGMENTS
First and foremost, I would like to thank my supervisor, Dr. Stephen Reid, without whom this
thesis project would not be possible. Dr. Reid, thank you for encouraging me to pursue this
accelerated Masters program and for your support and guidance throughout the course of this
project. I also wish to thank Dr. Herbert Kronzucker, Dr. Rene Harrison and Dr. Les Buck for
their input and suggestions. I am indebted to many student colleagues for their assistance at
various stages of this thesis project and they are: Jessica McAneney for her technical expertise
with the in vitro brainstem-spinal cord preparation and histological analysis of amphibian brain
sections; Balinda Phe for her knowledge on gelatin-chrome-alum coating of slides and tissue
staining protocols as well as her assistance with the analysis of stained brain tissue and finally,
Sherri Thiele for her input with regards to histochemical analysis of stained brain tissue. In
addition, I wish to thank the entire Reid lab (Balinda, Jeff, Jessica and Alex) for providing me
with an enjoyable and stimulating environment to conduct my thesis in. I would also like to
thank the Natural Sciences and Engineering Research Council of Canada (NSERC) for funding
this thesis project. Finally, to my family and friends, thanks for putting up with my long hours in
the laboratory and I am ever grateful for your love and support throughout the years.
Permission has been obtained from the copyright owners (Elsevier and Permissions Department
of Annual Reviews) and or authors (Dr. Stephen Reid) for the inclusion of figures (Figures 2 and
3 in Chapter 1) from published manuscripts and for the use of this thesis by the National Library.
iv
TABLE OF CONTENTS
ABSTRACT....................................................................................................................................ii
ACKNOWLEDGEMENTS............................................................................................................iii
TABLE OF CONTENTS................................................................................................................iv
LIST OF FIGURES......................................................................................................................viii
LIST OF ABBREVIATIONS..........................................................................................................x
CHAPTER 1: GENERAL INTRODUCTION................................................................................1
Preamble..............................................................................................................................2
Breathing in Anuran Amphibians........................................................................................3
The Mechanics of Breathing....................................................................................3
Bimodal Breathing...................................................................................................9
Discontinous Breathing..........................................................................................10
Respiratory Control Systems in Anuran Amphibians........................................................12
Central Control of Breathing.................................................................................12
Olfactory Chemoreceptors.....................................................................................14
Pulmonary Stretch Receptors................................................................................15
Peripheral (Arterial) Chemoreceptors....................................................................16
v
Central Chemoreceptors.........................................................................................17
The In Vitro Brainstem-Spinal Cord Preparation..............................................................18
Environmental Hypoxia and Hypercapnia.........................................................................22
Chronic Hypercapnia Studies............................................................................................24
Chronic Hypoxia Studies...................................................................................................30
Carbonic anhydrase............................................................................................................34
Discovery and Kinetics..........................................................................................34
Role of Carbonic Anhydrase in CO2 Chemoreception..........................................36
Hypothesis & Goals of the thesis.......................................................................................41
CHAPTER 2: EFFECTS OF CARBONIC ANHYDRASE INHIBITION
WITH ACETAZOLAMIDE ON FICTIVE BREATHING
IN CHRONICALLY HYPOXIC AND HYPERCAPNIC
LEOPARD FROGS (RANA PIPIENS)...................................................................43
Introduction........................................................................................................................44
Materials and Methods.......................................................................................................46
Experimental Animals...........................................................................................46
Exposure to Chronic Hypoxia and Hypercapnia...................................................46
In Vitro Brainstem Spinal Cord Preparations........................................................47
Acetazolamide........................................................................................................49
Experimental Protocol...........................................................................................50
vi
Data and Statistical Analysis.................................................................................51
Results................................................................................................................................53
Discussion..........................................................................................................................76
Conclusion.........................................................................................................................83
CHAPTER 3: EFFECTS OF EXOGENOUS CARBONIC ANHYDRASE
APPLICATION ON FICTIVE BREATHING IN ISOLATED
IN VITRO BRAINSTEM-SPINAL CORD PREPARATIONS
TAKEN FROM CHRONICALLY HYPOXIC AND
HYPERCAPNIC LEOPARD FROGS (RANA PIPIENS)......................................84
Introduction........................................................................................................................85
Materials and Methods.......................................................................................................87
Experimental Animals...........................................................................................87
Exposure to Chronic Hypoxia and Hypercapnia...................................................87
In Vitro Brainstem Spinal Cord Preparations........................................................87
Carbonic Anhydrase...............................................................................................87
Experimental Protocol...........................................................................................88
Data and Statistical Analysis.................................................................................88
Results................................................................................................................................89
Discussion........................................................................................................................110
Conclusion.......................................................................................................................114
CHAPTER 4: HISTOCHEMICAL ANALYSIS OF ACTIVE CARBONIC
ANHYDRASE IN BRAINSTEMS TAKEN FROM CONTROL,
CHRONICALLY HYPERCAPNIC AND CHRONICALLY
vii
HYPOXIC LEOPARD FROGS...............................................................................116
Introduction......................................................................................................................117
Materials and Methods.....................................................................................................120
Histochemical Localization of Active Carbonic Anhydrase...............................120
Results..............................................................................................................................122
Discussion........................................................................................................................125
Conclusion.......................................................................................................................128
CHAPTER 5: GENERAL DISCUSSION...................................................................................129
Goals of the Thesis...........................................................................................................130
Critique of In Vitro Brainstem-Spinal Cord Preparation.................................................131
CO2-Sensitive Respiratory Control Systems...................................................................133
Role of Carbonic Anhydrase in CO2 Chemoreception
Following Chronic Hypercapnia.....................................................................................135
Different Effects of Chronic Hypoxia in
Terrestrial Versus Aquatic Amphibians...........................................................................136
Perspectives......................................................................................................................138
REFERENCES CITED................................................................................................................141
viii
LIST OF FIGURES
Figure 1: Phases of anuran amphibian lung ventilation.
Figure 2: Buccal and lung pressure traces from an intact cane toad (Bufo marinus).
Figure 3: Comparison of vertebrate breathing traces.
Figure 4: The in vitro brainstem-spinal cord preparation.
Figure 5: The effect of chronic hypercapnia on fictive breathing in cane toads (Bufo marinus).
Figure 6: Effects of chronic hypercapnia on fictive breathing in leopard frogs (Rana pipiens).
Figure 7: The effect of chronic hypoxia on fictive breathing in cane toads (Bufo marinus).
Figure 8: Effects of chronic hypoxia on fictive breathing in leopard frogs (Rana pipiens).
Figure 9: Model of central respiratory-related pH/CO2 chemoreception.
Figure 10: Electroneurogram of vagal motor output from control, CHC and CH frogs.
Figure 11: Effects of chronic hypercapnia on fictive breathing in leopard frogs (Chapter 2).
Figure 12: Effects of chronic hypoxia on fictive breathing in leopard frogs (Chapter 2).
Figure 13: Effects of acetazolamide on fictive breathing frequency in leopard frogs.
Figure 14: Effects of acetazolamide on the number of fictive episodes per minute.
Figure 15: Effects of acetazolamide on the number of fictive breaths per episode.
Figure 16: Effects of acetazolamide on fictive breath duration in leopard frogs.
ix
Figure 17: Effects of acetazolamide on integrated fictive breath area in leopard frogs.
Figure 18: Effects of acetazolamide on total fictive ventilation in leopard frogs.
Figure 19: Effects of chronic hypercapnia on fictive breathing in leopard frogs (Chapter 3)
Figure 20: Effects of chronic hypoxia on fictive breathing in leopard frogs (Chapter 3).
Figure 21: Effects of carbonic anhydrase on fictive breathing frequency in leopard frogs.
Figure 22: Effects of carbonic anhydrase on the number of fictive episodes per minute.
Figure 23: Effects of carbonic anhydrase on the number of fictive breaths per episode.
Figure 24: Effects of carbonic anhydrase on fictive breath duration in leopard frogs.
Figure 25. Effects of carbonic anhydrase on integrated fictive breath area.
Figure 26: Effects of carbonic anhydrase on total fictive ventilation in leopard frogs.
Figure 27: Histochemical localisation of carbonic anhydrase using the cobalt-phosphate method.
x
LIST OF ABBREVIATIONS
aCSF: artificial cerebral spinal fluid
ACTZ: acetazolamide
ANOVA: analysis of variance
CA: carbonic anhydrase
CH: chronic hypoxia
CHC: chronic hypercapnia
cn V: cranial nerve V/ trigeminal nerve
cn VII: cranial nerve VII/ facial nerve
cn VIII: cranial nerve VIII/ auditory nerve
cn IX: cranial nerve IX/ glossopharyngeal nerve
cn X: cranial nerve X/ vagus nerve
CSF: cerebral spinal fluid
DMSO: dimethyl sulfoxide
E: enzyme
GPI: glycosyl-phosphatidyl-inositol
NI: nucleus isthmus
xi
pFRG: parafacial respiratory group
pHi: intracellular pH
PIP-C: phosphatidylinositol-specific phospholipase C
preBotC: pre-Bötzinger complex
preI: preinspiratory area
PSR: pulmonary stretch receptor
RM: repeated measures
SEM: standard error of the mean
SNK: Student–Newman–Keuls
1
CHAPTER 1
GENERAL INTRODUCTION
2
1. INTRODUCTION
1.1 Preamble
In air-breathing animals, the most important stimulus to breath is the level of CO2 in the
cerebral spinal fluid (CSF). The CO2 level in the CSF is monitored by central (brain) respiratory-
related pH/CO2-sensitive chemoreceptors (chemosensors) located on the ventral surface of the
medulla (Mitchell et al., 1963). Previous studies on the control of breathing in anuran
amphibians have demonstrated that exposure to chronic hypercapnia (elevated inspired CO2
levels) and chronic hypoxia (decreased inspired O2 levels) augments, and decreases, respectively,
the function of these central chemoreceptors (Gheshmy et al., 2006; McAneney and Reid, 2007).
It has been suggested that the hypercapnia-induced augmentation and hypoxia-induced
attenuation of these chemoreceptors results from alterations in the function of the enzyme
carbonic anhydrase (CA; the enzyme that catalyzes the reversible hydration/dehydration of CO2;
Roughton and Meldrum, 1933). The overall goal of this study was to address the hypothesis that
changes in CA function can account for the changes in central chemoreceptor function that occur
during chronic hypercapnia and chronic hypoxia in an aquatic anuran amphibian, the Northern
leopard frog (Rana pipiens).
The following introduction will cover the basics of anuran amphibian breathing, while
highlighting similarities and differences with respect to mammalian breathing. In addition,
respiratory control mechanisms and specifically, the topic of this thesis, central pH/CO2
chemoreceptor function following chronic hypoxia and hypercapnia, will also be discussed.
Finally, I will speculate on the role of carbonic anhydrase in the modulation of breathing
following chronic hypoxia and chronic hypercapnia.
3
1.2. Breathing in Anuran Amphibians
Although mammals, birds, reptiles and amphibians are all air-breathing animals, there are
distinct differences amongst all of them especially with respect to the mechanics of breathing.
The study of breathing in amphibians is also complicated due to the bimodal nature of their
breathing (that changes with development) and the presence of discontinuous breathing. Hence,
what follows is a discussion of these three differences between amphibians and other air-
breathers: use of a positive pressure pump to drive respiratory air flow (i.e., breathing
mechanics), anuran bimodal breathing and a discontinuous pattern of breathing.
1.2.1. The Mechanics of Breathing
Unlike mammals who breathe using a negative pressure aspiration pump driven by the
diaphragm and intercostal muscles, anuran amphibians (which lack a diaphragm and intercostal
muscles) use a positive pressure pump (also known as the buccal force pump) to force air into
their lungs (Gans et al., 1969; West and Jones, 1975). This buccal pump is driven by muscles in
the buccal cavity (i.e., the mouth) that force air from the mouth into the lungs. Laryngeal and
pharyngeal muscles are also used to control the amount of air flow, through the glottis, into the
lungs during lung ventilation (Sakakibara, 1984a; Gans et al., 1969).
Gans and colleagues (1969) outlined the mechanics of lung ventilation in Rana
catesbeiana (the American bullfrog) using pressure recordings from the lungs, buccal cavity and
abdominal cavity. West and Jones (1975) documented the mechanics of breathing in Rana
pipiens via similar measurement of changes in pressure and volume in the lungs and buccal
cavity accompanying breathing. Both of these papers provide the basis for the following
discussion on the mechanics of breathing in anuran amphibians.
4
The four sites involved in anuran amphibian breathing are the nares, buccal cavity, glottis
and lungs (refer to Fig. 1). The nares control the entrance and exit of air from the atmosphere
into the mouth (or buccal cavity) of the animal. The pumping of the buccal cavity via elevator
and depressor muscles located on the floor of the mouth allows air to be drawn into the buccal
cavity and later pumped into the lungs. The valve that controls air flow into and out of the lungs
is the glottis which is opened and closed with dilator and constrictor muscles, respectively. The
final component is the amphibian lung, which can account for up to 20-30% of the whole body
volume of bullfrogs when inflated (Gans et al., 1969).
Figure 1 depicts the various phases of lung ventilation in anuran amphibians. In the first
phase (Fig. 1A), the nares (which regulate air flow from the environment into the anuran’s
buccal cavity) and glottis (which regulates the air flow in and out of the lungs) are open and
closed, respectively. Depression of the buccal cavity (Fig. 1A) creates a negative pressure within
the buccal cavity causing air to flow through the nares and into the ventral region of the buccal
cavity (Fig. 1B). Note that the mouth is not involved in breathing and remains closed throughout
this process. Subsequent contraction of the laryngeal dilator muscles (m. dilatator laryngis)
causes the glottis to open (Fig. 1C). At this point, the air within the lungs (which is at a higher
pressure than that of ambient air) flows from the lungs via the glottis, through the upper region of
the buccal cavity and out the nares (Fig. 1C). Positioning of O2-rich air entering via the nares in
the posterior compartment of the buccal cavity and the flow of O2-poor air from the lungs
directly forward and out the nares, minimizes the admixture of these two gases. Closure of the
nares (achieved via contraction of m. submentalis in frogs; Gans and Pyles, 1983) and elevation
of the buccal cavity forces the fresh air from the ventral/posterior region of the buccal cavity
through the glottis and into the lungs (Fig. 1D).
5
Figure 1: Phases of anuran amphibian lung ventilation. Anuran amphibian lungs are ventilated
through a sequence of events represented here in panels A through D. Buccal depression with the
nares open and glottis closed (A) draws air into the buccal cavity (B). The glottis then opens (C)
allowing air under high pressure in the lungs to exit through the nares. Subsequent closure of the
nares and elevation of the buccal cavity forces air from the buccal cavity to enter the lungs (D).
The cycle then repeats itself as shown.
6
Following this, the closure of the glottis (which has been shown to be passive in Rana pipiens
(West and Jones, 1975) while active in Rana catesbieana (Gans et al., 1969) and the opening of
the nares (also passive in Rana pipiens; West and Jones, 1975) leads into the next respiratory
cycle (starting again at Panel A of Fig. 1).
The sequence of events described above can manifest as three different cycles (Gans et
al., 1969; West and Jones, 1975; see Figure 2). First, there is a pattern of air movement through
the nares, into the buccal cavity, that occurs with the glottis closed so that no air enters the lungs.
It has been suggested that this buccal oscillation cycle might serve to “flush out” the buccal
cavity and fill it with fresh air. Alternately, it has been suggested that these buccal oscillations
serve an olfactory purpose (Gans et al., 1969). Second, a lung ventilation cycle refers to the
ventilation of the lungs as described above with the lung volume returning to the pre-breathe
level at the end of each breath. Note that the pause between breaths occurs with the lungs inflated
in anurans unlike mammals, in which the pause occurs with the lung volume at functional
residual capacity (Reid and West, 2004). Third, sequential lung inflation breaths can occur
rapidly in succession to create a lung inflation cycle during which time the lungs are “pumped
up” or inflated. Similarly, when the lungs are in an inflated state there can be several deflation
breaths in sequence that lower lung volume. These consist of back-to-back breaths in which the
volume of air expired is greater than the volume inspired.
Figure 2 illustrates lung and buccal pressure traces from an intact (i.e., in vivo) cane toad
(Bufo marinus).
7
Figure 2: Buccal (top) and lung (bottom) pressure traces from an intact cane toad. Panel A
illustrates 4 breathing episodes, one of which is represented on a magnified scale in panel B. The
breathing sequence shown in panel B is that of a few balanced breaths (lung volume post-breath
is equivalent to that of pre-breath), followed by inflation breaths (successive breaths that result in
incremental increases in lung volume), breath holding (i.e., during which period the glottis is
closed) and subsequent deflation breaths (successive breaths that result in the gradual reduction
of lung volume). Consecutive inflation breaths constitute a lung inflation cycle (enclosed within
dotted lines) and result in complete inflation of the lungs. Figure reproduced with permission
from Reid (2006).
8
9
Panel A shows 4 distinct breathing episodes whereas panel B is a magnification of the trace in
panel A and shows only a single breathing episode. A lung inflation cycle, composed of several
inflation breaths and cumulating in the complete inflation of the lungs, is highlighted (i.e., using
dotted lines) in panel B. Subsequent breath holding (representative of a period of glottal closure)
and gradual deflation of the lungs leads into another breathing episode.
1.2.2. Bimodal Breathing
Anuran amphibians are bimodal breathers both as tadpoles and adults (Burggren and
West, 1982; Burggren and Doyle, 1986; Burggren and Infantino, 1994). As tadpoles, anuran
amphibians use both their gills and skin for gas (O2 and CO2) exchange, with cutaneous gas
exchange accounting for about 60% of O2 uptake and CO2 excretion (the remaining 40%
occurring across the gills, i.e., branchial gas exchange) in aquatic tadpoles of Rana catesbeiana
(the American bullfrog; Burggren and West, 1982). In air-breathing tadpoles, the gills start to
degenerate, and account for a relatively smaller (approximately 15%) portion of O2 uptake. This
reduction in the branchial contribution is compensated for by the involvement of the developing
lungs (accounting for approximately 15% of O2 uptake), with the remaining 70% occurring via
cutaneous gas exchange. Contrary to the changes in partitioning of O2 uptake seen at this stage,
CO2 excretion remains as before with the gills contributing about 40% and the skin 60% of CO2
excretion; the lungs still play no role. During metamorphosis, the lungs enlarge and develop
further while the gills begin to be reabsorbed. Post-metamorphic bullfrogs rely on their lungs and
skin for oxygen uptake (approximately 75 and 25%, respectively), with CO2 excretion occurring
predominately through the skin (about 90%, the remaining 10% is through the lungs). Following
metamorphosis, when the gills are no longer present, the adult anuran relies on its lungs as the
10
primary site for (approximately 90% of) O2 uptake (Burggren and West, 1982) while the skin
continues to play a significant role in CO2 excretion (Burggren and West, 1982; Pinder and
Burggren, 1986). Burggren and West (1982) reported that 80% of CO2 excretion in adult
American Bullfrogs (Rana catesbeiana), a semi-aquatic species, occurs through cutaneous
exchange. Far less data is available on the partitioning of O2 uptake and CO2 elimination across
the three respiratory gas exchanges surfaces (i.e., skin, lungs and gills) in Northern leopard frogs
(Rana pipiens; the subject of my thesis project). Pinder and Burggren (1986) stated that, in Rana
pipiens, cutaneous gas exchange can account for 23% of total oxygen uptake when the animals
are floating (i.e., inactivity) in well-aerated water at 25ºC. However, this amount decreases to
approximately 19% in active R. pipiens in well-aerated water at 25ºC (Pinder and Burggren,
1986). Both values reported for R. pipiens are slightly higher than that reported for R.
catesbieana (Burggren and West, 1982; Pinder and Burggren, 1986). However, I assume that
CO2 excretion will be similar in both aquatic species since experimental data is sparse in this
area for R. pipiens.
1.2.3. Discontinuous Breathing
Most air-breathing fish, reptiles and amphibians do not exhibit a continuous breathing
pattern as seen in euthermic mammals and most water-breathing fish (Milsom, 1991). Anuran
amphibian breathing tends to be discontinuous, consisting of either single breaths or doublets
(separated by periods of apnea), due to their relatively lower metabolic requirements (Milsom,
1991; McAneney and Reid, 2007; Gargaglioni and Milsom, 2007; see Figure 3).
11
Figure 3: Comparison of vertebrate breathing traces. These breathing traces, obtained from
various vertebrates, illustrate both continuous and discontinuous breathing patterns.
Discontinuous breathing can take the form of “randomly” distributed breaths or clusters of
breaths (episodes). Figure reproduced (with permission) from Milsom (1991).
12
In fact, the discontinuous nature of anuran breathing was recorded as early as 1891 and
characterized at that point as Cheyne-Stokes breathing (i.e., periodic breathing) in the frog
(Sherrington, 1891). As respiratory drive increases (i.e., during exposure to hypoxia or
hypercapnia), breathing becomes more episodic (i.e., several breaths are clustered together and
separated by periods of apnea). Eventually, with a high enough respiratory drive, the apneic
periods decrease as the number of breaths within each episode increase, resulting in continuous
breathing (Milsom, 1991; Gargaglioni and Milsom, 2007).
1.3 Respiratory Control Systems in Anuran Amphibians
Anuran amphibian breathing is regulated by various complex control systems located
both peripherally and centrally (i.e., within the brain). As in mammals, breathing is produced in
brainstem respiratory centers located in the medulla (McLean et al., 1995a,b; Reid et al., 2000a;
Wilson et al., 2002). However, much less in known about the central control of breathing (i.e.,
neural components responsible for rhythm generation and underlying mechanisms driving these
rhythm generators) in amphibians compared to mammals.
1.3.1. Central Control of Breathing
Two coupled, synchronous respiratory oscillators have been identified in the mammalian
rostro-ventral medulla. These are the pre-Bötzinger complex (preBotC) and the parafacial
respiratory group (pFRG; also known as the pre-inspiratory or pre-I area; Smith et al., 1991;
Onimaru and Homma, 2003; Feldman and Del Negro, 2006; Wilson et al., 2006).
Recently, Wilson and colleagues (2002) provided evidence to support the notion that
there are also two distinct respiratory-related neuronal oscillators in the bullfrog, a rostral site
13
controlling lung ventilation rhythm that is located between cranial nerves VIII and IX and a
caudal one controlling buccal rhythm located near cranial nerve X. Gill rhythm in earlier stages
of development in the amphibian (i.e., in tadpoles) seems to be driven by the same neural
oscillator as buccal rhythm (Vasilakos et al., 2006). Interaction between these two oscillators
(i.e., lung and buccal/gill) generates the overall respiratory rhythm (Vasilakos et al., 2006).
Further similarity between mammalian and amphibian respiratory rhythmogenesis is
shown in the responses of each respective pair of neural oscillators to application of μ opioid
receptor agonists. Takeda and colleagues (2001) showed that only the preBotC neurons are
depressed by opiates and that the pFRG/pre-I neurons are not affected with respect to their
oscillatory activity. A similar disparity is observed between buccal and lung oscillators in
amphibians. In frogs for example, lung rhythm is preferentially inhibited by opioids while buccal
rhythm remained unchanged (Vasilokos et al., 2006). These results, taken together, suggest that
the mammalian pFRG area is homologous to the amphibian buccal oscillator and the mammalian
preBotC is homologous to the amphibian lung oscillator (Wilson et al., 2006).
A recent study questioned whether or not there is a third respiratory-related oscillator in
amphibians that is responsible for clustering of lung breaths together into episodes (Wilson et al.,
2006). Earlier studies have shown that episodic breathing in bullfrogs can be eliminated by
transections made behind the optic lobe (Oka, 1958a,b). More recently, Reid and colleagues
(2000a) showed that a transection made slightly caudal to the optic chiasma resulted in the
conversion of an episodic to a continuous breathing pattern in in vitro bullfrog brainstem-spinal
cord preparations. Overall respiratory drive (the integration of all peripheral afferent input and
central processes) determines whether breathing is continuous or episodic in amphibians (as
discussed earlier in section 1.2.3). The nucleus isthmus (NI), a mesenchephalic structure located
14
between the roof of the midbrain and the cerebellum (Gargaglioni and Branco, 2004), appears to
be involved in integrating respiratory drive since its absence (via chemical lesions) converts
episodic breathing into a pattern of sparsely distributed single breaths (Kinkead et al., 1997;
Gargaglion and Branco, 2004). This region is however, not responsible for clustering of breaths
into episodes since Kinkead et al. (1997) showed that bilateral kainic acid lesions to the NI did
not alter breathing frequency or amplitude following increased respiratory drive (i.e., elicited by
pulmonary stretch receptor feedback).
To recap then, respiratory rhythm is generated in brainstem respiratory centres and is
modified by the overall respiratory drive (i.e., the result of integrating peripheral afferent inputs
and central processes in brain regions such as the NI). These modifications result in respiratory
breathing patterns (i.e., episodic vs. continuous) generated via efferent output from brainstem
respiratory centres to motor neurons controlling respiratory muscles. A brief discussion of the
various central processes and peripheral afferent inputs alluded to in the previous statements is
presented below.
1.3.2. Olfactory Chemoreceptors
Olfactory chemoreceptors, located in the nasal epithelium, are sensitive to CO2 (Coates
and Ballam, 1990; Coates, 2001). Olfactory chemoreceptors inhibit breathing when stimulated
by high levels of inspired CO2 (i.e., ranging from 0.4- 4% CO2) via olfactory nerve input to the
brain (Sakakibara, 1978; Coates and Ballam, 1990; Coates, 2001). In support of this, olfactory
denervation has been shown to increase breathing frequency following hypercapnia compared to
olfactory intact control animals (Kinkead and Milsom, 1996). The mechanism via which
15
olfactory chemoreceptors sense and respond to CO2 is not clear but Coates and colleagues (1998)
suggest that carbonic anhydrase may be involved. The role of carbonic anhydrase in CO2
chemoreception will be discussed later in this introduction.
1.3.3. Pulmonary Stretch Receptors
Pulmonary stretch receptors (PSR) are located in the walls of the lungs and monitor lung
inflation and deflation (Reid and West, 2004). PSR input is sent to respiratory centres in the
brainstem via pulmonary vagi (Reid et al., 2000b). There are three types of PSRs in anurans:
slow-adapting (tonic) receptors, rapidly-adapting (phasic) receptors or a mixture of the two (Reid
and West, 2004). The rapidly-adapting receptors increase firing during lung inflation and
deflation but are far less active during sustained lung inflation (i.e., phasic PSR activity). On the
other hand, slow-adapting receptors show a delayed, but lasting, activity in response to lung
inflation (in other words, tonic activity). Studies on leopard frogs and bullfrogs have shown that
pulmonary stretch receptors are also CO2 sensitive and are inhibited by increasing levels of CO2
(Milsom and Jones, 1977; Kuhlman and Fedde, 1979). Removal of PSR feedback to central
respiratory centres via pulmonary vagotomy in decerebrate, paralyzed and uni-directionally
ventilated bullfrogs resulted in reduced breathing frequency in bullfrogs (Kogo et al., 1994;
Kinkead and Milsom, 1997). Similarly, studies on in vitro brainstem-spinal cord preparations
(which lack afferent input and will be discussed in a later portion of this introduction) of anurans
have shown a reduced hypercapnic response compared to that observed in vivo (Kinkead et al.,
1994; McLean et al., 1995a,b; Kinkead and Milsom, 1996; Reid and Milsom, 1998; Gheshmy et
al., 2006). However, electrical stimulation of the pulmonary vagi in these in vitro preparations
(mimicking PSR feedback) increases respiratory motor output in response to lowered superfusate
16
pH (Kinkead et al., 1994; Reid and West, 2004). Such studies have shown the importance of
pulmonary stretch receptor feedback on breathing and the hypercapnic response in anurans.
1.3.4. Peripheral (Arterial) Chemoreceptors
Peripheral chemoreceptors, located in the carotid labyrinth (the amphibian equivalent of
the carotid body but occupying a much more diffuse location) and aortic and pulmocutaneous
arches (Hoffmann and DeSousa, 1982; West et al., 1987; Smatresk and Smits, 1991), are
sensitive to both CO2 and O2 (Smatresk and Smits, 1991). Under normoxic, normocapnic
conditions, these receptors provide a baseline level of tonic, stimulatory afferent input to the
brain. However, under hypoxic and hypercapnic conditions, the level of input increases,
contributing to increases in breathing (West et al., 1987; Smatresk and Smits, 1991). In
amphibians, the carotid labyrinth arises from the first extant gill arch and is innervated by the
glossopharyngeal nerve (cn IX; Milsom and Burleson, 2007). The discharge from the
glossopharyngeal nerve acts on brainstem respiratory centres to modulate breathing.
The sensitivity of these arterial chemoreceptors for O2 is dependent on the level of CO2
and vice versa. For example, in Bufo marinus, a significant reduction in arterial CO2 levels will
cause breathing to cease regardless of the O2 level (West et al., 1987). Therefore, if CO2 levels in
blood are low, breathing doesn’t occur even if O2 levels are lower than normal. Hence, O2 is only
a stimulus for breathing if CO2 levels are sufficiently high enough to allow breathing to proceed.
On the other hand, hyperoxic conditions will also prevent breathing even in the presence of
elevated arterial CO2 levels (West et. al., 1987). Taken together, these observations suggest that
signals from O2 and CO2-sensitive chemoreceptors are integrated to produce an overall level of
respiratory drive (Smatresk and Smits, 1991; Reid, 2006). For a mathematical model and
17
graphical representation of the interaction between PO2 and PCO2 sensitivity of peripheral
chemoreceptors in mammals, refer to Duffin (2005).
1.3.5. Central Chemoreceptors
Central pH/CO2 chemoreceptors trigger an increase in breathing when they are stimulated
by a reduction in cerebral spinal fluid (CSF) pH or increases in CSF CO2 levels (Smatresk and
Smits, 1991; Kinkead and Milsom, 1994; Milsom, 2002; Lahiri and Forster, 2003). This was first
documented by Leuson (1950) via perfusion of artificial cerebrospinal fluid (aCSF) containing
high PCO2 levels into the cerebral ventricles (lateral to fourth ventricles) of dogs, thus stimulating
ventilation (Leuson, 1972). The earliest evidence for the existence of a central chemoreceptor
drive to breathing in ectothermic air breathers was obtained by Hitzig and Jackson (1978) who
perfused cerebral ventricles (lateral to fourth ventricles) of turtles (Pseudemys scripta) with
aCSF of low pH and showed hyperventilation that persisted for the duration of perfusion. More
than 40 years ago, central chemoreceptors were localized to the ventrolateral surface of the
medulla oblongata by Mitchell and colleagues (1963) who showed that application of filter paper
soaked in hypercapnic solutions to this region in anesthetised dogs stimulated ventilation. These
central chemoreceptive regions are now thought to be widely distributed throughout the
brainstem (Coates et al., 1993). Focal tissue acidification using acetazolamide (an inhibitor of
carbonic anhydrase; will be discussed later in this introduction) application to the following
regions stimulated breathing: the locus coeruleus in mammals (Coates et al., 1993) as well as
amphibians (Noronha-de-Souza et al., 2006), the nucleus tractus solitarii, the medullary raphe,
the retrotrapezoid nucleus, and the pre-Bötzinger complex in mammals (Coates et al., 1993;
Nattie, 1999). A current review by Nattie and Li (2008) indicates that the presence of multiple
18
sites of central CO2 chemoreception produce an enhanced response to simultaneous stimulation.
In addition to this, multiple neuronal types (for example serotonergic and glutamatergic) and
parallel PCO2 sensing mechanisms with multiple pH/CO2 sensors (i.e., inhibition of inward
rectifying K+ channels, Na+/Ca2+ exchangers, TASK-1 channel, gap junctions, etc) are suggested
to exist (Jiang et al., 2005).
Decreases in cerebral spinal fluid pH, caused by increased PCO2 levels, elicit an
immediate (activation occurs within a second; Gray, 1971) response from the central
chemoreceptors resulting in increased ventilation. Despite this, the stimulus for pH/CO2
chemoreception remains elusive. There are two different mechanisms postulated to explain CO2
chemoreceptor stimulation and subsequent signal transduction; the membrane potential
hypothesis and the Na+/Ca2+ exchange hypothesis (see below).
1.4 The In Vitro Brainstem-Spinal Cord Preparation
This thesis will focus on central pH/CO2 chemoreceptors and their sensitivity to changes in
cerebral spinal fluid pH following chronic hypercapnia (CHC) and chronic hypoxia (CH). The
central control of breathing is frequently studied using in vitro preparations such as the isolated
brainstem-spinal cord preparation. This is a reduced, decerebrate preparation which includes the
midbrain, brainstem and a short segment of the spinal cord. This preparation is discussed now
(rather than in the Materials and Methods) because many of the studies that form the basis for
this thesis used this preparation. Given this, it is necessary to have an appreciation of this
preparation prior to reading sections of the introduction below. In this preparation, motor output
recorded from respiratory-related cranial nerves (i.e., nerves that control respiratory muscles)
serves as an index of breathing termed fictive breathing.
19
Figure 4. The in vitro brainstem-spinal cord preparation. Panel A is a picture of the experimental
set-up used to record motor output (i.e., fictive breathing) from cranial nerves in the in vitro
brainstem-spinal cord preparation taken from leopard frogs. The brainstem-spinal cord
preparation is pinned into the recording chamber, superfused with artificial cerebrospinal fluid
(aCSF) and whole-nerve recordings made via a suction electrode attached to the
micromanipulator. Panel B depicts the relative size of a frog brain (ventral view). Panel C is an
illustration of the frog brain (dorsal view) with the cranial nerves labelled and dashed lines
indicating the transections that were made in order to produce the brainstem-spinal cord
preparation.
20
Figure 4
21
Figure 4 shows a photograph and accompanying illustration of this preparation. The brainstem-
spinal cord preparation, once removed from the cranial case of the animal, is pinned into a
recording chamber and continually superfused with oxygenated artificial cerebrospinal fluid
(aCSF; a modified Ringer’s solution). Motor output from cranial nerve rootlets that innervate
respiratory muscles in the intact animal is used as an index of breathing termed fictive breathing.
Gassing the superfusate with increasing or decreasing CO2, acidifies or alkalinizes it,
respectively. Hence, acidification of aCSF is representative of respiratory acidosis.
This technique was first developed by Suzue (1984) as an alternative to the brain slice for
studies examining respiratory central pattern generator(s) in neonatal rat brains. Using this
technique, Suzue (1984) recorded spontaneous periodic activity from the phrenic, hypoglossal
and other spinal nerves. Synchrony between spontaneous discharge from the phrenic nerve and
contraction of the diaphragm measured in vivo and a comparison of this discharge with phrenic
nerve root activity in vitro suggested that the in vitro spontaneous motor output corresponds to
respiratory-related nerve discharge in the intact animal (Suzue, 1984). The main advantage of
this in vitro brainstem-spinal cord preparation is that it contains much more of the neural
circuitry that controls breathing than is present in a brain slice preparation. Furthermore, this
preparation serves as a means to study the central control of breathing in the absence of any
peripheral input (Reid and Milsom, 1998). Since its initial use in studies on neonatal rats (Suzue,
1984), this preparation has been used to study the central control of breathing in a wide variety of
animals including air- and water-breathing fish (McClellan,1984; Wilson et al., 2000; Bongianni
et al., 2006), amphibians (Galante et al., 1996; McLean and Remmers, 1997; Reid and Milsom,
1998; Torgerson et al., 1998; Delvolvé et al., 1999; Gdovin et al., 1999), reptiles (Douse and
Mitchell, 1990; Johnson and Mitchell, 1998; Johnson and Mitchell, 2000) and hibernating
22
mammals (Bagust et al., 1985; Keifer and Kalil, 1989; Zimmer and Milsom, 2004). These in
vitro brainstem-spinal cord studies have confirmed that central pH/CO2 sensing chemoreceptors
increase in vitro fictive breathing in response to low cerebral spinal fluid pH, which is indicative
of high blood CO2 levels (Kinkead et al., 1994; Lahiri and Forster, 2003; Gheshmy et al., 2006).
In vitro brainstem-spinal cord preparations from ectotherms such as amphibians remain
viable for longer periods of time compared to those from mammals due to their lower tissue
metabolic rates, maintenance of normoxia in the regions containing respiratory control centres
via diffusion of oxygen from the bath solution (artificial cerebrospinal fluid), greater
anoxia/hypoxia tolerance and proximity of the superfusion conditions to their normal body
temperature ranges (Reid and Milsom, 1998; Morales and Hedrick, 2002).
1.5 Environmental Hypoxia and Hypercapnia
Prior to examining laboratory studies involving chronic hypoxic and hypercapnic
exposure, it is prudent to first point out circumstances under which amphibians encounter such
situations in their natural environments. Anuran amphibians can experience environmental
hypoxia and/or hypercapnia during overwintering under ice-covered bodies of water (i.e., aquatic
species) or in underground burrows (i.e., terrestrial species) as observed in leopard frogs (R.
pipiens) and cane toads (Bufo marinus), respectively (Pinder et al., 1992).
Data on the gas levels encountered by burrowing amphibians has not been well
documented. Boggs and colleagues (1984) have reported CO2 levels as high as 6% and O2 levels
as low as 14% in mammalian burrows. Given the similarity between the burrow systems of
23
amphibians and mammals, it is likely that such fossorial amphibians are also exposed to similar
conditions of hypoxia and hypercapnia during their estivation and/or overwintering period.
Terrestrial and semi-terrestrial anurans such as the cane toad (Bufo marinus) usually
burrow under moist soil and remain dormant during the winter (Pinder et al., 1992).
Hypoventilation, in addition to a reduced capacity for cutaneous CO2 exchange to occur (as their
skin is covered in soil) is said to double blood PCO2 in these terrestrial species (Pinder et al.,
1992). However, plasma pH compensation (mainly via HCO3- gain from the environment) is
relatively low (a species dependent range of 0-30% during chronic hypercapnia) in amphibians
(Toews and Boutilier, 1986; Boutilier and Heisler, 1988).
Ranid frogs such as leopard frogs (Rana pipiens), which I conducted my thesis
experiments on, typically “hibernate” while submerged under water during which time they
tolerate a certain level of hypoxia and hypercapnia via a reduction in metabolism and with the
aid of cutaneous gas exchange (Stewart et al., 2004). Stewart and colleagues (2004) further state
that the ability to retain aerobic metabolism, at low PO2 levels for a period of months, in addition
to anoxia tolerance for a period of days, is sufficient for frogs to overwinter with limited chance
of death.
Rana pipiens occasionally overwinter on land (Rand, 1950; Emery et al., 1972). In this
case they burrow into soil as do terrestrial amphibians and hence experience the same difficulties
faced by species such as the cane toad. Reduced metabolism (to conserve energy),
hypoventilation and reduced cutaneous gas exchange will lead to respiratory acidosis (Pinder et
al., 1992). Simkiss (1968) states that some amphibians maybe able to liberate CaCO3 from their
bones or nodules found within endolympathic sacs in order to buffer arterial pH changes. In
24
addition, studies have shown that the haemoglobin-O2 binding affinity in overwintering bullfrogs
is high during hypoxic exposure (Pinder et al., 1992).
1.6 Chronic Hypercapnia (CHC) Studies
Exposure to acute hypercapnia causes an increase in ventilation. This hypercapnic
ventilatory drive arises mainly from central pH/CO2 chemoreceptor stimulation (which can
account for up to 80% of overall hypercapnic ventilatory drive) in vivo in Bufo paracnemis
(Branco et al., 1992) with a lesser contribution from the peripheral (arterial) CO2
chemoreceptors. Numerous studies have examined the effects of acute hypercapnia on amphibian
respiration (Boutilier et al., 1979; Toews and Heisler, 1982; Toews and Stiffler, 1990; Smatresk
and Smits, 1991; Kinkead and Milsom, 1994; Kinkead and Milson, 1996). However, the
responses to chronic hypercapnia and the mechanisms that underlie them have only recently been
investigated. A previous study from this laboratory (Gheshmy et al., 2006) reported that
exposure of cane toads to CHC (3.5% CO2 for 9 days) caused an increase in central pH/CO2
chemoreceptor-stimulated fictive breathing measured using the in vitro brainstem-spinal cord
preparation (Fig. 5).
Further studies were conducted in our laboratory in order to determine the mechanisms
that may be responsible for the CHC-induced increase in central pH/CO2-sensitive fictive
breathing. Hypotheses were put forth suggesting that the central pH/CO2 chemoreceptor function
was altered either by descending central input (i.e., from the midbrain) or by afferent input (i.e.,
from olfactory chemoreceptors, pulmonary stretch receptors, or arterial chemoreceptors).
25
Figure 5: Fictive breathing frequency (fictive breaths·min-1) as a function of artificial
cerebrospinal fluid (aCSF) pH recorded from brainstem spinal cord preparations taken from
chronically hypercapnic (CHC; closed circles) and normocapnic control (open circles) cane toads
(Bufo marinus). The data are plotted as mean values ± 1 SEM. Letters (a, b, and c) indicate a
significant difference amongst pH levels in any one group. A plus sign (+) indicates a significant
difference between CHC and controls. Figure modified from Gheshmy et al., 2006.
aCSF pH
7.4 7.6 7.8 8.0 8.2
Fic
tive
Bre
athi
ng F
requ
ency
(br
eath
s·m
in-1
)
0
5
10
15
20
25
30
35b, +
ba
a a
Chronic Hypercapnia
Control
26
To elucidate the role of higher brain centres (i.e., midbrain regions) in the chronic
hypercapnia (CHC)-induced increase in central pH/CO2-sensitive fictive breathing, midbrain
transections were performed at a level previously shown to alter episodic breathing (i.e., caudal
to optic chiasm; Reid et al., 2000a; Gheshmy et al., 2006). Gheshmy and colleagues (2006)
reported that midbrain transections, at a level slightly caudal to the optic chiasma, did not alter
the chronic hypercapnia-induced increase in central pH/CO2-sensitive fictive breathing, leading
to the conclusion that descending input from the rostral half of the midbrain does not contribute
to the chronic hypercapnia-induced increase in fictive breathing.
Next, the role of olfactory chemoreceptors was determined via olfactory nerve
denervation. Gheshmy and colleagues (2007) determined that the CHC-induced increase in
central pH/CO2 chemoreceptor function (fictive breathing) was abolished by denervation (prior
to CHC) of the CO2-sensitive olfactory chemoreceptors which inhibit breathing when stimulated
by high levels of CO2. In other words, it seems that the CHC-induced increase in central pH/CO2
chemoreceptor function is a compensatory response to offset the increase in inhibitory input
from olfactory CO2 chemoreceptors to brainstem respiratory centres, which would presumably
occur during CHC.
In order to examine any influence of arterial chemoreceptors on the CHC-induced
increase in fictive breathing, Gheshmy et al. (2007) exposed cane toads to simultaneous CHC
(3.5 % CO2) and chronic hyperoxia (30% O2). Since peripheral chemoreceptor sensitivity to PO2
and PCO2 are interdependent, with high levels of PO2 leading to a relatively low peripheral CO2
respiratory drive, simultaneous CHC and chronic hyperoxia would result in minimal input from
the peripheral CO2 chemoreceptors to the brainstem respiratory centres. Reducing afferent input
from peripheral O2/CO2 chemoreceptors within the carotid labyrinth prevented the CHC-induced
27
increase in fictive breathing that occurred in response to central chemoreceptor stimulation in in
vitro brainstem-spinal cord preparations of cane toads (Gheshmy et al., 2007). Hence, arterial
chemoreceptor input is important in the modulation of central pH/CO2 chemosensitivity
following CHC.
These studies do not however, delve into mechanisms at the chemoreceptor level that
may have induced the augmentation in fictive breathing following CHC exposure. This topic will
be dealt with in this thesis in an aquatic anuran amphibian species, the Northern leopard frog
(Rana pipiens). However, prior to the commencement of the experiments in this thesis, the
effects of chronic hypercapnic exposure on fictive breathing was examined in leopard frogs
(Srivaratharajah and Reid, unpublished). This study showed a similar augmentation in fictive
breathing following CHC (Fig. 6; unpublished data).
28
Figure 6: (A) Fictive breathing frequency (fictive breaths·min-1), (B) the number of fictive
episodes per minute, (C) the number of fictive breaths per episode, (D) fictive breath duration,
(E) the integrated area of fictive breaths (V·s) and (F) total fictive ventilation (V·s·min-1) as a
function of artificial cerebrospinal fluid (aCSF) pH in chronically hypercapnic (closed squares)
and normocapnic control (open circles) leopard frogs (Rana pipiens). The data are plotted as
mean values ± 1 SEM. Letters (a, b, and c) indicate a significant difference amongst pH levels in
any one group. A plus sign (+) indicates a significant difference between CHC and controls.
29
A. Fictive Breathing FrequencyF
ictiv
e B
reat
hing
Fre
quen
cy (
brea
ths·
min
-1)
0
10
20
30
Chronic Hypercapnia
Control ab
a
a, +
b, +b, +
B. Fictive Episodes per Minute
Num
ber
of F
ictiv
e E
piso
des
per
Min
ute
0
5
10
15
20
25
30
Chronic Hypercapnia
Control ab
a
+
+
C. Fictive Breaths per Episode
aCSF pH
7.4 7.6 7.8 8.0 8.2Num
ber
of F
ictiv
e B
reat
hs p
er E
piso
de
0.5
1.0
1.5
2.0
2.5
Chronic Hypercapnia
Control
D. Fictive Breath Duration
Fic
tive
Bre
ath
Dur
atio
n (s
)
0.1
0.2
0.3
0.4
0.5
0.6
Control
Chronic Hypercapnia
E. Integrated Fictive Breath AreaIn
tegr
ated
Fic
tive
Bre
ath
Are
a (V
·s)
0.00
0.01
0.02
0.03
0.04
Control
Chronic Hypercapnia
F. Total Fictive Ventilation
aCSF pH
7.4 7.6 7.8 8.0 8.2Tot
al F
ictiv
e V
entil
atio
n (V
·s·m
in-1
)
0.0
0.1
0.2
0.3
0.4
0.5
Control
Chronic Hypercapnia
a, +
b, +b
aa
b
Figure 6
30
1.7 Chronic Hypoxia Studies
Responses to chronic hypoxia in anuran amphibians have also been investigated in past
studies. A previous study from this laboratory by McAneney et al. (2006) showed that, following
acclimatisation to chronic hypoxia (CH; 10% O2) for 10 days, the acute hypoxic ventilatory
response, but not resting ventilation, of cane toads was blunted in comparison to normoxic
controls. More recently, McAneney and Reid (2007) determined that CH led to a decrease in the
sensitivity of central (brainstem) respiratory-related pH/CO2 chemoreceptors (which normally
trigger an increase in breathing when stimulated by elevated CO2/low pH), measured using in
vitro brainstem-spinal cord preparations of cane toads (Fig. 7). This reduction in pH/CO2
chemoreceptor function was reversed by a midbrain transection, suggesting that descending
inhibitory inputs may be involved in blunting the sensitivity of central chemoreceptors in
response to CH (McAneney and Reid, 2007). Although potential cellular mechanisms underlying
O2 chemoreception have been documented, the involvement of carbonic anhydrase remains to be
determined.
The study on the effects of CH on fictive breathing was then replicated in leopard frogs
(Srivaratharajah and Reid, unpublished) prior to beginning the experiments in this thesis. The
results of this study were not consistent with those of McAneney and Reid (2007) as exposure to
CH did not alter fictive breathing in this species (Fig. 8).
31
Figure 7: Fictive breathing frequency (fictive breaths·min-1) as a function of artificial
cerebrospinal fluid (aCSF) pH in chronically hypoxic (CH; closed circles) and normoxic control
(open circles) cane toads (Bufo marinus). The data are plotted as mean values ± 1 SEM. Letters
(a, b, and c) indicate a significant difference amongst pH levels in any one group. A plus sign (+)
indicates a significant difference between CHC and controls. Figure modified from McAneney
and Reid (2007).
7.6 7.8 8.0
2
4
6
8
10
12
14
aCSF pH
Fic
tive
Bre
athi
ng F
requ
ency
(br
eath
s/m
in)
Control
Chronic Hypoxia
b
a, b
+
Fictive Breathing Frequency
a
32
Figure 8: (A) Fictive breathing frequency (fictive breaths·min-1), (B) the number of fictive
episodes per minute, (C) the number of fictive breaths per episode, (D) fictive breath duration
(E) the integrated area of fictive breaths (V·s) and (F) total fictive ventilation (V·s·min-1) as a
function of artificial cerebrospinal fluid (aCSF) pH in chronically hypoxic (closed circles) and
normoxic control (open circles) leopard frogs (Rana pipiens). The data are plotted as mean
values ± 1 SEM. Letters (a, b, and c) indicate a significant difference amongst pH levels in any
one group. A plus sign (+) indicates a significant difference between CH and controls.
33
Fic
tive
Bre
athi
ng F
requ
ency
(br
eath
s·m
in-1
)
0
10
20
30
Chronic Hypoxia
Controlb
b
aa
b
a
A. Fictive Breathing FrequencyN
umbe
r of
Fic
tive
Epi
sode
s pe
r M
inut
e
0
5
10
15
20
25
30
Control
Chronic Hypoxia
aa
b
ab
b
B. Fictive Episodes per Minute
aCSF pH
7.4 7.6 7.8 8.0 8.2Num
ber
of F
ictiv
e B
reat
hs p
er E
piso
de
0.5
1.0
1.5
2.0
2.5
Control
Chronic Hypoxia
C. Fictive Breaths per Episode
Fic
tive
Bre
ath
Dur
atio
n (s
)
0.1
0.2
0.3
0.4
0.5
0.6
Control
Chronic Hypoxia
+
D. Fictive Breath Duration
Inte
grat
ed F
ictiv
e B
reat
h A
rea
(V·s
)
0.00
0.01
0.02
0.03
0.04
Control
Chronic Hypoxia
E. Integrated Fictive Breath Area
aCSF pH
7.4 7.6 7.8 8.0 8.2Tot
al F
ictiv
e V
entil
atio
n (V
·s·m
in-1
)
0.0
0.1
0.2
0.3
0.4
0.5
Control
Chronic Hypoxia
a a
b
F. Total Fictive Ventilation
Figure 8
34
1.8 Carbonic Anhydrase
1.8.1 Discovery and Kinetics
The uncatalyzed hydration and dehydration of CO2 (equation 1) are relatively fast
reactions with first order reaction rate constants of approximately 0.035 and 20 sec-1,
respectively (Henry and Swenson, 2000). Lahiri and Forster (2003) report a half-time of 3.7s for
the uncatalyzed hydration/dehydration reaction of CO2 under physiological conditions (i.e., pH
7.4 and 37ºC).
Equation 1: CO2 + H2O H+ + HCO3-
However, early in the 1930s it was determined that these reaction rates were not sufficient to
explain the rate at which CO2 was eliminated from blood plasma during the approximately 1
second transit time through pulmonary capillaries (Meldrum and Roughton, 1933). This
discrepancy lead to the discovery of a group of enzymes called carbonic anhydrases. Carbonic
anhydrases (CA) are ubiquitous enzymes that catalyze the reversible hydration of carbon dioxide
Meldrum and Roughton, 1933). Since its discovery approximately 70 years ago, CA has been
under extensive study and has been associated with a wide range of physiological processes such
as breathing; acid-base balance; bone resorption; production of aqueous humour, cerebrospinal
fluid, gastric acid, and pancreatic juice; and possibly cell growth (Sly and Hu, 1995; Chegwidden
and Carter, 2000). There are 3 families (α, β and γ) of CA, each with several isoforms; however,
I will only be discussing the αCAs.
αCAs are zinc metalloenzymes that are found predominately in animals and possess an
average molecular weight of approximately 29 kDa (Chegwidden and Carter, 2000). The first
αCA isoform was discovered in 1962 (Rickli and Edsall, 1962). Currently, there are at least 15
35
isoforms of αCA (which from here on will be referred to simply as CA) in mammals, ranging
from CA I and II found in mammalian erythrocytes to sulfonamide-resistant CA III found in rat
liver, membrane-bound CA IV, mitochondrial CA V, cell-secreted CA VI to the recently
discovered isozyme CA XV (Dodgson, 1991; Fernley, 1990; Hilvo et al., 2007). CA II is the
most commonly found isozyme and is also the most highly active with an overall enzymatic
catalytic rate (kcat) greater than 1 ×106 s-1 (Chegwidden and Carter, 2000). CA II is present in
human red blood cells (Dodgson, 1991) and a related isozyme was purified from bullfrog
erythrocytes (Chegwidden, 1991). In fact, studies done on anuran amphibians have shown the
presence of an isozyme that closely resembles mammalian CA II in reaction rate and inhibition
characteristics (Bundy and Cheng, 1976; Ziegler et al., 1974; Rosen and Friedley, 1973; Scott
and Skipski, 1979). Other possible isozymes in anurans have not been discussed in the literature.
The 3 main steps in the CA catalyzed CO2 hydration mechanism have been described in
several studies (Lindskog et al., 1984; Liljas et al., 1994; Vince et al., 2000; Chegwidden and
Carter, 2000) and is as follows (refer to equation 2 below). The zinc atom in the active site of the
CA enzyme (E-Zn) reacts with water to form a E-Zn-OH- complex (equation 2a). Nucleophilic
attack on this E-Zn-OH- complex by CO2 in the vicinity of the active site forms E-Zn-HCO3
(equation 2b). The bicarbonate bound to the Zn atom is then displaced by a water molecule
(equation 2c). The rapid transfer of an H+ from E-Zn-H2O to buffers (B) in the surrounding
solution regenerates E-Zn-OH- (equation 2d) and the cycle continues (from equation 2b through
2d). For a more detailed review of this catalytic mechanism refer to Lindskog and Silverman
(2000).
36
Equation 2:
a. E-Zn + H2O + B → E-Zn-OH- + B-H+
b. E-Zn-OH- + CO2 → E-Zn-HCO3-
c. E-Zn-HCO3- + H2O → E-Zn-H2O + HCO3
-
d. E-Zn-H2O + B → E-Zn-OH- + B-H+
1.8.2 Role of Carbonic Anhydrase in CO2 Chemoreception
With chemoreceptor activation occurring within a second of hypercapnic or acidic
stimulation, it is not feasible for the uncatalyzed hydration of CO2 to account for any major
aspect of CO2 chemoreception (Gary, 1971). Hence, CA was implicated in the mechanism of
CO2 chemoreception. In support of this, studies have shown that inhibition of CA reduced
chemoreceptor response (Black et al., 1966; Travis, 1971; Gary, 1971). These initial studies
drew a connection between carotid body CO2 chemoreceptors and CA. The distribution of the
isozymes within the brain, however, remains an area in need of further research. It was initially
thought that CA isozymes were restricted to glial cells and generally not present in neurons
(Giacobini, 1961; Parthe, 1981). However, later studies have proven otherwise (Brown, 1980;
Wong et al., 1983; Wong et al., 1987; Neubauer, 1991). It has been suggested that CA is
involved in CO2 chemoreception by providing a link between intracellular CO2/H+ and neuronal
signalling (Neubauer, 1991; Torrence, 1993). Several studies have provided evidence in support
of this notion via localization of CA activity to previously identified central CO2 chemoreceptive
regions. For example, Ridderstråle and Hanson (1985) and Torrance (1993) reported CA activity
near the ventral surface of the cat medulla, the long accepted location of central respiratory-
related chemoreceptors. Furthermore, Coates and colleagues (1998) reported the presence of CA
37
in olfactory CO2 chemoreceptors in bullfrogs (R. catesbeiana) and Lahiri (1991) described CA
localization in the glomus cells of the cat carotid body.
The standard mechanistic model for CO2 chemoreception according to previous studies is
as follows. CO2 in the blood enters the cerebral spinal fluid and diffuses into chemoreceptor cells
(Fig.9, step 1). Within the chemoreceptor cells, carbonic anhydrase catalyzes the reversible
hydration of CO2 (Fig. 9 step 2), where it is converted into a bicarbonate ion and a proton (Fig. 9,
step 3). The rapid accumulation of H+ ions in the cell leads to a number of ion exchange
processes (Fig. 9, step 4) that lead to Ca2+ influx. Two different hypotheses have been postulated
to explain the link between chemoreceptor stimulation and the rise in intracellular Ca2+ which is
required to initiate neurotransmitter release (Rocher et al., 1991; Buckler and Vaughan-Jones,
1993; Peers and Buckler, 1995). The first is the Na+/Ca2+ -exchange hypothesis and the second is
the membrane potential hypothesis (Peers and Buckler, 1995).
Non-chemoreceptive cells possess means for intracellular pH (pHi ) regulation using
Na+/H+ antiporters that counteract a decrease in pHi via electroneutral exchange of intracellular
H+ for Na+ (Putnam, 2001) and HCO3-/Cl- antiporters that counteract an increase in pHi via
efflux of HCO3-. The presence of CA in chemosensitive cells, on the other hand, expedites H+
production, which may overwhelm these pHi regulatory mechanisms, leading to a maintained
intracellular acidosis and chemoreceptor firing as long as PCO2 remains elevated (Necakov,
2002). The Na+/Ca2+ -exchange hypothesis (Rocher et al., 1991; Gonzalez et al. 1992; Peers and
Buckler, 1995) proposes that a fall in pHi activates Na+/H+ antiporters, which result in the efflux
of H+, and Na+/HCO3- symporters that further increase the influx of Na+ (Fig. 9, step 4).
38
Figure 9. Carbonic anhydrase (CA) and the mechanistic model of central pH/CO2
chemoreception. Arterial CO2 crosses the blood brain barrier and enters neuronal pH/CO2
chemosensitive cells (step 1). CA within pH/CO2 chemoreceptor cells catalyze the reversible
hydration of CO2 (step 2), forming H+ and HCO3- (step 3). Accumulation of H+ leads to a drop in
intracellular pH and subsequent ion exchange processes (step 4). Ion exchange leads to Ca2+
influx and neurotransmission (step 5), ultimately stimulating ventilation.
39
The rise in intracellular Na+ (Fig. 9, step 4) then activates Ca2+/Na+ exchangers which result in
the influx of Ca2+ in exchange for the efflux of Na+. The resultant increase in intracellular Ca2+
triggers neurotransmitter release from the chemoreceptor cells via exocytosis (Fig. 9, step 5).
This signal then reaches the brainstem respiratory centres, which then increase ventilation via
efferent motor output to respiratory muscles.
The basis for the membrane potential hypothesis (Peers and Buckler, 1995; Peers, 2004),
which has recently garnered support, is the acid-induced inhibition of K+ channels. Such an
inhibition would reduce K+ efflux and result in cell membrane depolarization, subsequent Ca2+
influx and signalling of respiratory centres. The role of CA in the rapid accumulation of
intracellular H+ could also be applied to this hypothesis, whereby the resultant rapid decrease in
pH would inhibit K+ channels.
In order to determine the physiological function of CA in the chemoreceptive process,
many studies (Coates et al., 1991; Erlichman et al., 1994; Necakov et al, 2002; Taylor et al.,
2003) have used CA inhibitors to examine the effects of reducing or eliminating CA activity
(yielding stronger evidence than histochemical co-localization studies). Some CA inhibitors
include sulfanomides, metal ions such as Cu (II) and Hg (II), imidazole, phenol, nitrate and
perchlorate (Eriksson and Liljas, 1991). Sulfanomides are a class of strong CA inhibitors and
include acetazolamide (membrane permeant), benzolamide (poorly permeant), dichlorphenamide
and methazolamide (Dodgson, 1991). By far, the most commonly used CA inhibitor is
acetazolamide (ACTZ) which has pKa values of 7.2 and 8.8 (i.e., what these values mean is that
ACTZ is a diprotic weak acid and exists in a deprotenated, ionic form at physiological pH) and
an IC50 value of 9.9 nM for CA II (Maren, 1967). Sulfonamide inhibitors of CA such as ACTZ
reversibly inhibit CA activity by displacing the water molecule coordinated to the Zn atom at the
40
active site of the enzyme (refer back to equation 2). Hence, the Zn-OH complex is prevented
from forming and the proceeding nucleophilic attack by CO2 cannot occur (Liljas et al., 1994).
Taylor and colleagues (2003) applied 25 μM ACTZ to the superfusate of in vitro
brainstem-spinal cord preparations taken from bullfrogs (R. catesbeiana) of various
developmental stages in order to test central CO2 chemosensitivity. 25 μM is a concentration that
is approximately 250 times the IC50 value of ACTZ for CA II and, at which, 99% of CA II
activity is said to be inhibited (Maren, 1967; Taylor et al., 2003). Taylor et al. (2003) reported a
local acidification-induced increase in fictive breathing in all stages of development except for
that of late stage tadpoles. Erlichman and colleagues (1994) showed that bath application of
ACTZ to the isolated brain-pneumostome preparation of the pulmonate snail (Helix aspersa)
slowed the ventilatory response (i.e., pneumostomal opening) to rapid CO2 changes. ACTZ
application to the ventrolateral medullary surface in cats also delayed the ventilatory response to
CO2 (Coates et al., 1991). These studies seem to indicate that CA participates in central CO2
chemoreception. However, a study undertaken by Necakov and colleagues (2002) showed that
bath application of 1 mM ACTZ to transverse medullary slices from neonatal rats did not seem
to alter breathing (in this case, fictive breathing as measured from hypoglossal nerve activity).
Recently, Nattie and Li (2006) proposed that central chemoreception need not involve a single
neuronal type but rather involves an assortment of neurons (such as glutamatergic and
serotonergic neurons). Note that glial cells may also be involved in CO2 sensing since they also
possess carbonic anhydrase isozymes.
41
1.9 Hypothesis and Goals of the Thesis
Building upon the foundation of these previous studies that have linked CA to central
chemoreception, the aim of my thesis was to examine the role of CA in the CHC-induced
augmentation in fictive breathing (central pH/CO2 chemosensitivity) in amphibian in vitro
brainstem-spinal cord preparations. To accomplish this, the first series of experiments used bath
application of the membrane permeant CA inhibitor, acetazolamide (ACTZ), to access whether
or not CHC acclimatization and the subsequent increase in in vitro fictive breathing in leopard
frogs (R. pipiens) was altered by blocking CA. If this occurred, the data would suggest that CHC
has caused an increase in CA activity/amount that in turn augmented the central chemoreceptor
function.
Due to the fact that ACTZ is a relatively permeable CA inhibitor and hence will inhibit
both intracellular and extracellular CA, I sought a means to distinguish between the two. In the
second series of experiments, I examined the effect of a bath application of bovine CA (isozyme
type II) in the in vitro brainstem-spinal cord preparations from control and CHC leopard frogs.
Assuming that intracellular CA is responsible for central CO2 chemoreception (on the basis that
intracellular pH changes trigger these chemoreceptors; Ritucci et al., 1997; 1998; Wang et al.,
2002; Putnam et al., 2004) and the CHC-induced augmentation in fictive breathing, I expected
exogenous CA application to have no effect on fictive breathing (given that CA is relatively cell
impermeable).
These two series of experiments were also performed on in vitro brainstem-spinal cord
preparations taken from frogs exposed to chronic hypoxia in order to access the potential
involvement of CA in O2 chemoreception following this chronic respiratory challenge. Finally, I
42
performed a histochemical comparison between the intensity of staining for CA in brains taken
from frogs exposed to control conditions, CHC and CH using a modified version of Hansson’s
method (also referred to as the Cobalt-Phosphate method; Hansson, 1967). Assuming that the
CHC-induced augmentation in fictive breathing frequency is a result of CA upregulation, I
hypothesized that the stain intensity would be greater in the CHC compared to control and CH
frog brains.
43
CHAPTER 2
EFFECTS OF CARBONIC ANHYDRASE INHIBITION WITH ACETAZOLAMIDE ON
FICTIVE BREATHING IN CHRONICALLY HYPOXIC AND HYPERCAPNIC LEOPARD
FROGS (RANA PIPIENS)
44
2.1 INTRODUCTION
Previous studies have shown that exposure to chronic hypercapnia (CHC) augments
(Gheshmy et al. 2006) while exposure to chronic hypoxia (CH) attenuates (McAneney and Reid,
2007) fictive breathing measured from in vitro brainstem spinal cord preparations taken from a
terrestrial amphibian, the cane toad (Bufo marinus). The CHC-induced augmentation was due to
alterations in afferent input from olfactory and peripheral chemoreceptors (Gheshmy et al. 2007)
while the CH-induced attenuation was due to central influences, from the midbrain, to the
medullary respiratory centres. Exposure of a semi-aquatic anuran (the leopard frog; Rana
pipiens) to CHC also caused an increase in fictive breathing measured in vitro while exposure to
CH had no effect (Srivaratharajah and Reid, unpublished).
The above-mentioned studies indicate a role for altered afferent input and central
processes in the CHC-induced and CH-induced, respectively, changes in fictive breathing.
However, these studies do not address the possible mechanisms responsible for this effect at the
central chemoreceptor level. This chapter addresses whether a change in carbonic anhydrase
(CA) amount/activity is involved in the changes in fictive breathing that occur following CHC.
Although exposure to CH did not cause an attenuation of fictive breathing in Rana pipiens
(Srivaratharajah and Reid, unpublished) I also investigated whether changes in CA
amount/activity occurred following exposure to CH.
I hypothesised that an augmentation in the activity/amount of CA was responsible for the
CHC-induced increase in fictive breathing. To test this hypothesis, I examined whether inhibition
of CA with acetazolamide (ACTZ), a potent cell-permeant CA inhibitor, would alter the CHC-
induced increase in fictive breathing. The assumption here is that ACTZ will block both
45
intracellular and extracellular CA (although we assume the vast majority of CA is intracellular;
Ritucci et al., 1997; 1998; Wang et al., 2002; Putnam et al., 2004), hence slowing down CO2
hydration. Consequently, as more CO2 is bubbled into the superfusate, the drop in intracellular
pH (pHi) will occur at a slower rate than it would in the absence of ACTZ. This would allow
time for pHi regulation mechanisms to stabilize any changes in pHi, leading to a reduction in the
output of central CO2 chemoreceptors and an eventual reduction in fictive breathing (Necakov et
al., 2002). Given that CH did not alter fictive breathing in this species, I hypothesised that ACTZ
would have no effect on fictive breathing following CH.
46
2.2 MATERIALS & METHODS
2.2.1 Experimental Animals
Adult leopard frogs (Rana pipiens; N = 26; approximately 5 to 8 cm in length) were
obtained from a commercial supplier (Boreal Scientific, St. Catharines, Ontario) and transported
to the University of Toronto, Scarborough. Frogs were housed in large tanks supplied with
running dechlorinated water as well as dry terrestrial landings. The animals were held at room
temperature (20–22ºC) with the photoperiod maintained at 12 h light:12 h dark. Frogs were fed
live crickets and/or earthworms twice per week. Holding conditions and experimental protocols
were approved by the University of Toronto Animal Care Committee and conform to the
guidelines established by the Canadian Council for Animal Care.
2.2.2 Exposure to Chronic Hypoxia and Hypercapnia
Leopard frogs were exposed to chronic hypoxia (CH; N = 7) or chronic hypercapnia
(CHC; N = 11) for a 10-day period in a Plexiglas chamber. To establish conditions of CHC, the
inspired CO2 within the chamber was maintained at 3.5% using a ProCO2 120 control unit
(Biospherix, NY, USA). A CO2 electrode within the chamber measured the CO2 level. When it
fell below 3.5%, the ProCO2 unit delivered a small amount of CO2 into the chamber to raise the
level back to 3.5%. To establish CH, the inspired O2 within the chamber was maintained at 10%
using a ProOx 110 control unit (Biospherix, NY, USA). An O2 electrode in the chamber
monitored the level of O2. When the O2 level rose above 10%, the ProOx unit delivered a small
amount of N2 to lower the level back to 10%. A level of 3.5% CO2 (CHC) and 10% O2 (CH)
were selected based on previous studies (Gheshmy et al., 2006; 2007; McAneney et al., 2007;
47
Srivaratharajah et al., 2007). The hypercapnic and hypoxic acclimatisation chambers were
maintained at room temperature and exposed to a 12 h light:12 h dark cycle. Normoxic
normocapnic leopard frogs (exposed to room air; 20.97% O2; 0.03% CO2; 79% N2) maintained
under the same temperature and light conditions were used as controls.
2.2.3 In vitro Brainstem-Spinal Cord Preparations
Leopard frogs were anaesthetised via emersion in a solution of 3-aminobenzoic acid ethyl
ester (MS222, 0.6 g l−1
; Sigma–Aldrich Inc., Oakville, Ontario, Canada) neutralised to pH 7.4
with sodium bicarbonate (Reid and Milsom, 1998). Animals were kept in the anaesthetic until
eye-blink (or corneal) and toe-pinch (or withdrawal) reflexes were eliminated. The brain and
spinal cord were removed en bloc from the rest of the animal (Taylor et al., 2003). Bone shears
and rongeurs were used to remove the cranial case, exposing the brain from the olfactory bulb to
the spinal cord. Immediately following exposure, the brain was superfused continually with ice-
cold oxygenated artificial cerebrospinal fluid (aCSF) (in mmol l−1
; NaCl, 103; KCl, 4.05; MgCl2,
1.38; glucose, 10; NaHCO3, 25; CaCl2, 2.45; Sigma–Aldrich Inc.; pH 7.8; Taylor et al., 2003;
Gheshmy et al., 2006; 2007; McAneney and Reid, 2007; Srivaratharajah et al., 2008). Following
decerebration (i.e., removal of the forebrain region, leaving just enough to pin the preparation
into the recording chamber), the spinal cord was severed at the level of the third spinal nerve, the
cranial nerves were cut distal to their roots and the brainstem–spinal cord, with the midbrain
attached, was removed. The brainstem–spinal cord preparation was then transferred onto a
Sylgard-coated dissecting dish and immobilised using insect pins.
The dura matter surrounding the brain was partially removed in order to free the cranial
nerve roots and the nerve tips were cut to provide a clean surface for recording. The preparation
48
was then pinned in place over a fine nylon mesh within a recording chamber. The brainstem–
spinal cord preparation, in the recording chamber, was continuously superfused with oxygenated,
room temperature aCSF (at a rate of 10 ml/min) using peristaltic pumps that delivered and
removed the aCSF from the chamber. The nylon mesh, onto which the brainstem–spinal cord
preparation was pinned (ventral side up), divided the chamber into upper and lower
compartments, which facilitated simultaneous superfusion of both the dorsal and ventral surfaces
of the preparation (Kinkead et al., 1994; Reid and Milsom, 1998; Gheshmy et al., 2006; 2007;
McAneney and Reid, 2007).
Using a micro-manipulator, two narrow diameter suction electrodes were positioned near
the end of the vagus nerve root (cranial nerve X; cnX) and the trigeminal nerve root (cranial
nerve V; cnV) and the nerves were aspirated into the electrode such that a tight seal was
obtained. The suction electrodes were formed from thin-walled capillary glass pulled to a fine tip
using a vertical pipette puller (Kopf model 720; Tujunga, CA, USA) and polished using a
grinding stone. Electroneurogram recordings were taken of whole-nerve discharge from the
vagus nerve root and trigeminal nerve root. In the intact animal, the laryngeal branch of the
vagus nerve innervates the glottis, which opens and closes with each breath (Sakakibara, 1984;
Kogo et al., 1997), whereas the trigeminal nerve innervates the nasal mucosa (Sakakibara, 1978)
and buccal elevator muscles (Sakakibara, 1984). Since brainstem–spinal cord preparations lack
afferent input and breathing is an inherently rhythmic process generated in the brainstem, all
spontaneous rhythmic activity recorded from the vagus and trigeminal nerve roots was assumed
to represent motor output to the respiratory muscles (Kinkead et al., 1994; McLean et al., 1995a,
b; Reid and Milsom, 1998; Gheshmy et al., 2006; 2007; McAneney and Reid, 2007). This nerve
activity is the neural correlate of breathing and is termed fictive breathing (Kinkead et al., 1994;
49
McLean et al., 1995a, b; Galante et al., 1996; Reid and Milsom, 1998; Reid et al., 2000a, b;
Gheshmy et al., 2006; 2007; McAneney and Reid, 2007).
Nerve activity from the suction electrodes was initially amplified 10X and filtered (30
Hz, high pass; 1 kHz, low pass) using a DAM50 AC amplifier (World Precision Instruments;
Sarasota, FL, USA) the output of which was sent to a second AC amplifier (ISO8A, WPI) and
amplified a further 100X. The amplified, filtered nerve signal was sent to a moving averager
(CWE MA821/RSP; CWE Inc., Ardmore, PA, USA; time constant = 200 ms) for integration and
to an audio monitor (AM Systems Model 3300; Carlsborg, WA, USA). The amplified/filtered
and integrated traces were monitored and stored using a data acquisition system (Biopac
Systems, MP150; Goleta, CA, USA). The sampling rate of analogue to digital conversion was
2000 Hz. Gassing the aCSF with varying levels of CO2 (0–5% CO2; balance O2) altered the
aCSF pH. The levels of O2 and CO2 gassing the aCSF were set using digital mass flow
controllers (Smart-Trak 100, Sierra Instruments; Monterey, CA, USA). The pH level of the aCSF
was monitored using a pH electrode (VWR) calibrated with standard buffers (pH 7.0 and 10.0)
and placed within the aCSF reservoir.
2.2.4 Acetazolamide
Two different acetazolamide (ACTZ; Sigma-Aldrich Inc., Oakville, Ontario, Canada)
concentrations were used in this study (1 and 10 µM). ACTZ is only slightly soluble in water
(approximately 0.7 mg/ml; Kaur et al., 2002), hence dimethyl sulfoxide (DMSO; Sigma-Aldrich
Inc., Oakville, Ontario, Canada) was used to dissolve ACTZ. Since ACTZ has an optimum
stability at pH 4 (Parasrampuria and Gupta, 1989) and is present in its ionized form at
physiological pH, its transport across lipid membranes may be limited. Hence, DMSO (an
50
amphipathic molecule) also served to enhance the cell permeability (Yu and Quinn, 1998) of
ACTZ at the pH levels used in this study. Due to earlier speculation on the possible stimulatory
effect of DMSO on respiration (De la Torre and Rowed, 1974) and recent evidence for increases
in brain metabolic rate at concentrations of DMSO as low as 0.000025% (v/v) (Nasrallah et al.,
2008), a equivalent amount of DMSO was added to the aCSF reservoir containing no ACTZ.
Furthermore, the concentrations used in this study were derived from a 100X stock solution of
ACTZ made fresh each day and diluted to 1 and 10 µM.
2.2.5 Experimental Protocol
Prior to recording, the brainstem–spinal cord preparation was allowed to stabilise at room
temperature in aCSF (containing DMSO) of pH 7.8 for 1 h. This pH approximates plasma pH of
anuran amphibians under normoxic normocapnic conditions (Reid and Milsom, 1998) while
room temperature is within the temperature range in which fictive breathing is consistently active
from these preparations (Morales and Hedrick, 2002). Following the 1-h stabilisation period and
the observation of stable levels of fictive breathing, each preparation was exposed to 3 different
levels of aCSF pH (7.6, 7.8, and 8.0 which correspond to acute hypercapnia, normocapnia and
hypocapnia, respectively). Each pH change was achieved over a period of 5-10 min and the
preparations were allowed to acclimatise to each new pH level for a further 5-10 min before
fictive breathing was monitored for an additional 20 min period of data-collection. The different
aCSF pH levels were delivered in random order (Gheshmy et al., 2006; 2007; McAneney and
Reid, 2007). Once the abovementioned aCSF pH changes were complete, the preparation was
superfused with aCSF containing 1 µM ACTZ (which was first dissolved in DMSO). After 30
min of superfusion with 1 µM ACTZ, the pH changes (7.6, 7.8 and 8.0) were repeated as
51
described above. The preparation was then superfused, for 30 min, with aCSF containing no
ACTZ following which it was superfused with aCSF containing 10 µM ACTZ. After 30 min of
superfusion with 10 µM ACTZ, the pH changes (7.6, 7.8 and 8.0) were repeated again as
described above. Since the effects of ACTZ and its half-life are not known for this in vitro
preparation (a biological half-life of 8.5 ± 2.5 hours in plasma is reported for ACTZ suspensions
administered to humans; Schoenwald et al., 1978), the order of exposure to various ACTZ levels
was not changed; 0 µM ACTZ + DMSO was followed by 1 µM ACTZ + DMSO and finally 10
µM ACTZ + DMSO. These experiments were performed on brainstem–spinal cord preparations
taken from leopard frogs exposed to CHC (N = 11), CH (N =7) as well as control (normoxic
normocapnic) conditions (N = 8).
2.2.6 Data and Statistical Analysis
AcqKnowledge 3.7.3 (Biopac Systems) software was used to acquire and store nerve
recordings. Data were analysed for the last 10 min of the recording period at each aCSF pH level
and values are reported as the mean ± one standard error of the mean (S.E.M.). Respiratory-
related neural activity was distinguished using criteria described by Reid and Milsom (1998).
Fictive breathing traces were analysed to determine fictive breathing frequency (fictive
breaths·min-1
), the number of fictive breathing episodes per min, the number of fictive breaths
per episode, fictive breath duration (s), and the integrated area of fictive breaths (i.e., the area
under the curve of the integrated vagal motor activity trace; V·s) which is considered a correlate
of breath amplitude or volume (Sakakibara, 1984; McAneney and Reid, 2007). Episodes were
designated as breaths occurring within 2 s of each other (Kinkead et al., 1994; Reid and Milsom,
1998; Gheshmy et al., 2006; 2007; McAneney and Reid, 2007). Total fictive ventilation
52
(V·s·min−1
) was calculated as the product of fictive breathing frequency and the integrated area
of fictive breaths and is used here as an index of overall breathing (i.e., fictive breathing).
All statistical analyses were performed using commercial software (SigmaStat 3.0; SPSS,
Chicago, IL, USA). The effects of changing aCSF pH in any given group (i.e., controls, CHC or
CH), were analysed using a one-way repeated measures (RM) analysis of variance (ANOVA)
followed by a Student–Newman–Keuls (SNK) multiple comparison test. The effects of CHC or
CH were analysed using a two-way non-repeated measures ANOVA (control/CHC or
control/CH X aCSF pH) followed by a SNK multiple comparison test. In any given group
(controls; CHC; CH) the effect of ACTZ was analysed using a two-way RM ANOVA (aCSF pH
X ACTZ level) followed by an SNK multiple comparison test. In all cases, p < 0.05 was taken to
be the limit of statistical significance.
53
2.3 RESULTS
Figure 10 illustrates fictive breathing traces recorded, at an aCSF pH of 7.6, from the
vagus nerve root of brainstem-spinal cord preparations taken from normoxic normocapnic (Fig.
10A), chronically hypercapnic (CHC; Fig. 10B) and chronically hypoxic (CH; Fig. 10C) animals
during treatment with 0, 1 and 10 1µM acetazolamide (ACTZ).
2.3.1 Effects of Chronic Hypercapnia
In both control and CHC groups, fictive breathing frequency (Fig. 11A) was elevated,
compared to pH 8.0, as the aCSF pH level was reduced (control: pH 7.6, p= 0.042; CHC: pH
7.6, p= 0.001; pH 7.8, p= 0.027). Fictive breathing frequency was significantly greater in the
CHC preparations compared to controls at an aCSF pH 7.6 (p= 0.028).
The components of fictive breathing frequency are number of fictive episodes per minute
(Fig. 11B) and number of fictive breaths per episode (Fig. 11C). In the CHC group, the number
of fictive episodes per minute increased significantly at pH 7.6 compared to the value at pH 8.0
(p= 0.006). However, the number of fictive episodes per minute remained unaltered as aCSF pH
was changed in the controls (p= 0.064). The number of fictive breaths per episode did not
change as a function of aCSF pH in either CHC or control preparations (control: p= 0.967; CHC:
p= 0.674). The number of fictive episodes per minute was greater in the CHC group, compared
to controls, at pH 7.6 (p= 0.016). The number of fictive breaths per episode remained unchanged
following CHC exposure (p= 0.934).
In both groups, fictive breath duration was unaltered by changes in aCSF pH (Fig 11D;
controls, p= 0.531; CHC, p= 0.077). Compared to controls, CHC blunted fictive breath duration
54
at pH 7.6 (p= 0.042). The integrated area of the fictive breaths (Fig. 11E) was also unaltered by
changes in aCSF pH in both the control and CHC preparations (controls, p= 0.828; CHC, p=
0.913). The integrated area of the fictive breaths was not significantly altered by exposure to
CHC (p = 0.132).
In the control group, total fictive ventilation (Fig. 11F) increased significantly as aCSF
pH was lowered to 7.6 from 8.0 (p= 0.022). Similarly, total fictive ventilation increased in the
CHC group following acidification of the aCSF from pH 8.0 to pH 7.8 (p= 0.028) and 7.6 (p=
0.004). Total fictive ventilation was significantly higher in CHC preparations compared to
controls at aCSF pH 7.6 (p= 0.03).
2.3.2 Effects of Chronic Hypoxia
Note, the control data plotted in Figure 12 is the same as the control data plotted in Figure
11. The effects of pH on fictive breathing in this group were described in the section above. In
preparations taken from chronically hypoxic frogs, fictive breathing frequency (Fig.12A) was
elevated at aCSF pH 7.6 compared to pH 8.0 (p= 0.021). However, there was no significant
difference in fictive breathing frequency between the control and CH preparations (p=0.138).
The components of fictive breathing frequency, fictive episodes per minute (E/M; Fig.12B) and
the number of fictive breaths per episode (B/E: Fig. 12C), were not altered by changes in aCSF
pH (E/M: p= 0.620; B/E: p= 0.172) nor where they altered by exposure to CH (E/M: p = 0.061;
B/E: p=0.883).
In the CH group, fictive breath duration (Fig. 12D) did not changes as aCSF pH was
lowered (p= 0.366). Compared to the control group, CH blunted fictive breath duration at aCSF
55
pH levels of 7.6 (p= 0.044) and 8.0 (p= 0.022) but not 7.8 (p= 0.058). The integrated area of
fictive breaths (Fig. 12E) was not altered by changes to aCSF pH (p= 0.489). However,
integrated fictive breath area was significantly lower in the CH group, compared to controls, at
aCSF pH 7.6 (p= 0.045).
In the CH group, total fictive ventilation (Fig. 12F) was augmented in response to a
reduction in aCSF pH from 8.0 to 7.8 (p= 0.032) and 7.6 (p= 0.006). Total fictive ventilation
was not significantly different between the control and CH preparations (p= 0.876).
56
Fig. 10. Electroneurograms of vagal motor output recorded at an artificial cerebrospinal fluid pH
of 7.6 from in vitro brainstem-spinal cord preparations taken from a normoxic normocapnic
control frog (column A), a chronically hypercapnic frog (column B) and a chronically hypoxic
(column C) frog. Within each group (column), there are 3 sets of electroneurograms: Pre-ACTZ,
following 1µM ACTZ addition and finally 10µM ACTZ addition. The top trace within each
electroneurogram is the raw vagal nerve activity (X) and the bottom trace is the integrated one (∫
X).
57
58
Fig. 11. Fictive breathing frequency (A), number of fictive episodes per minute (B), number of
fictive breaths per episode (C), fictive breath duration (D), integrated fictive breath area (E) and
total fictive ventilation (F), as a function of artificial cerebrospinal fluid (aCSF) pH, recorded
from in vitro brainstem-spinal cord preparations taken from normoxic normocapnic control (n=8;
open circles) and chronically hypercapnic (CHC; n=11; closed squares) leopard frogs. The data
are plotted as mean values ± 1 SEM. Letters (a and b) indicate a significant difference amongst
pH levels in any one group. A plus sign (+) indicates significant differences between CHC and
controls.
59
Fic
tive
Bre
ath
ing
Fre
qu
ency
(bre
aths·
min
-1)
0
5
10
15
20
25
30
A. Fictive Breathing Frequency
Chronic Hypercapnia
Control
a
b
b, +
a, bb
a
Nu
mber
of
Fic
tiv
e E
pis
odes
per
Min
ute
0
5
10
15
20
25
30
B. Fictive Episodes per Minute
a
a,b
b, +
Control
Chronic Hypercapnia
C. Fictive Breaths per Episode
aCSF pH
7.6 7.8 8.0
Num
ber
of
Fic
tive
Bre
ath
s
per
Epis
ode
0
1
2
3
4
5
6
Control
Chronic Hypercapnia
Fic
tive
Bre
ath D
ura
tion (
s)
0.1
0.2
0.3
0.4
0.5
D. Fictive Breath Duration
Control
Chronic Hypercapnia+
Inte
gra
ted F
icti
ve
Bre
ath A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
0.016
E. Integrated Fictive Breath Area
Control
Chronic Hypercapnia
aCSF pH
7.6 7.8 8.0
Tota
l F
icti
ve
Ven
tila
tio
n (
V·s
·min
-1)
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
Control
Chronic Hypercapnia
F. Total Fictive Ventilation
a, bb
a
b, +
b
a
Figure 11
60
Fig. 12. Fictive breathing frequency (A), number of fictive episodes per minute (B), number of
fictive breaths per episode (C), fictive breath duration (D), integrated fictive breath area (E) and
total fictive ventilation (F) as a function of cerebrospinal fluid (aCSF) pH, recorded from in vitro
brainstem spinal cord preparations taken from normoxic normocapnic control (n=8; open circles)
and chronically hypoxic (CH; n=7; closed circles) leopard frogs. The data are plotted as mean
values ± 1 SEM. Letters (a and b) indicate a significant difference amongst pH levels in any one
group. A plus sign (+) indicates significant differences between CH and controls.
61
Fic
tive
Bre
ath
ing
Fre
quen
cy
(bre
ath
s·m
in-1
)
0
5
10
15
20
25
30
Chronic Hypoxia
Control
aa, b
b
a
a, b
b
A. Fictive Breathing Frequency
B. Fictive Episodes per Minute
Nu
mb
er o
f F
icti
ve
Ep
isod
es
per
Min
ute
0
5
10
15
20
25
30
Control
Chronic Hypoxia
aCSF pH
7.6 7.8 8.0
Num
ber
of
Fic
tive
Bre
ath
s
per
Epis
od
e
0
1
2
3
4
5
6
Control
Chronic Hypoxia
C. Fictive Breaths per Episode
Fic
tive
Bre
ath
Du
rati
on
(s)
0.1
0.2
0.3
0.4
0.5
Control
Chronic Hypoxia+ +
D. Fictive Breath Duration
Inte
gra
ted F
icti
ve
Bre
ath A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
0.016
Control
Chronic Hypoxia+
E. Integrated Fictive Breath Area
aCSF pH
7.6 7.8 8.0
To
tal
Fic
tive
Ven
tila
tio
n (
V·s
·min
-1)
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
Control
Chronic Hypoxia a
a, b
b
a
bb
F. Total Fictive Ventilation
Figure 12
62
2.3.3 Effects of Acetazolamide
Fictive Breathing Frequency: Neither dose of ACTZ (1µM or 10µM) had any effect on
fictive breathing frequency in preparations taken from control frogs (Fig. 13A; p= 0.306). In
preparations taken from chronically hypercapnic (CHC) animals (Fig. 13B), addition of both 1
and 10µM ACTZ reduced fictive breathing frequency at an aCSF pH level of 7.6 (1µM; p =
0.04; 10µM; p = 0.011). In the CH group (Fig. 13C), neither dose of ACTZ had any statistically-
significant effect on fictive breathing frequency although there was a trend for the 10µM dose to
reduce breathing frequency (p= 0.068). In the CH group there was a significant difference in
fictive breathing frequency between the two doses of ACTZ at aCSF pH 7.8 (p= 0.048).
Fictive Episodes per Minute: ACTZ treatment (1 and 10 µM) had no effect on the
number of fictive episodes per minute in preparations taken from control (Fig. 14A; p= 0.797)
and CH (Fig. 14C; p= 0.122) animals. In the CHC group (Fig. 14B), both doses of ACTZ
reduced the number of episodes per minute at pH 7.6 (1µM: p= 0.026; 10µM: p= 0.004).
Fictive Breaths per Episode: In the control (Fig. 15A) and CHC (Fig. 15B) groups, there
was no effect of either dose of ACTZ on the number of fictive breaths per episode (controls: p=
0.930; CHC: p= 0.533). In the CH (Fig. 15C) group, 1 µM ACTZ caused an increase in the
number of fictive breaths per episode at an aCSF pH of 7.8 (p = 0.005) while 10 µM caused a
decrease at pH 7.6 (p= 0.026). Additionally, a significant difference between the 1 and 10 µM
doses was observed in the CH group at aCSF pH 7.6 (p= 0.026).
Fictive Breath Duration: Neither dose of ACTZ had any effect on fictive breath duration
in the control (Fig. 16A; p= 0.440), CHC (Fig. 16B; p= 0.302) or CH (Fig. 16C; p= 0.613)
groups.
63
Integrated Fictive Breath Area: In the control group (Fig. 17A) 1 µM ACTZ had no
effect on integrated fictive breath area (p= 0.165) while 10 µM ACTZ caused a reduction in
integrated fictive breath area at pH 7.8 (p= 0.02). Neither dose of ACTZ had any effect on fictive
breath duration in the CHC (Fig. 17B; p= 0.151) or CH (Fig. 17C; p= 0.188) groups.
Total Fictive Ventilation: In the control group (Fig. 18A) 1 µM ACTZ had no effect on
total fictive ventilation (p = 0.066) while 10 µM caused a significant decrease at pH 7.6 (p=
0.045). In the CHC group (Fig. 18B), both doses of ACTZ caused a significant reduction in total
fictive ventilation at pH 7.8 (1 µM; p= 0.015; 10 µM, p= 0.003) and pH 7.6 (1 µM; p= 0.002; 10
µM, p <0.001). Neither dose of ACTZ had any effect on total fictive ventilation in the CH group
(Fig. 18C; 1 µM: p= 0.558; 10 µM: p= 0.073).
64
Fig. 13. Fictive breathing frequency (breaths·min-1
) as a function of artificial cerebrospinal fluid
(aCSF) pH in preparations taken from (A) normoxic normocapnic control frogs (n=8), (B)
chronically hypercapnic frogs (CHC; n=11) and (C) chronically hypoxic frogs (CH; n=7) prior to
ACTZ addition (open circles), following 1µM ACTZ addition (closed circles) and following
10µM ACTZ addition (closed squares). The data are plotted as mean values ± 1 SEM. Letters (a,
and b) indicate a significant difference amongst pH levels in any one group. A number sign (#)
indicates significant differences between the Pre-ACTZ condition and 1 or 10µM ACTZ
addition. An ampersand (@) indicates a significant difference between the 1 and 10µM ACTZ
additions.
65
Fic
tiv
e B
reat
hin
g F
req
uen
cy
(bre
ath
s·m
in-1
)
0
5
10
15
20
25
30
Fic
tiv
e B
reat
hin
g F
req
uen
cy
(bre
ath
s·m
in-1
)
0
5
10
15
20
25
30
aCSF pH
7.5 7.6 7.7 7.8 7.9 8.0 8.1
Fic
tiv
e B
reat
hin
g F
req
uen
cy
(bre
ath
s·m
in-1
)
0
5
10
15
20
25
30
Pre-ACTZ
Pre-ACTZ
Pre-ACTZ
1µM ACTZ
1µM ACTZ
1µM ACTZ
10µM ACTZ
10µM ACTZ
10µM ACTZ
b
b
a
abb, #
a
a, b
b
a
a, b
b
a
b
b
#
@
A. Control
B. Chronic Hypercapnia
C. Chronic Hypoxia
Figure 13
66
Fig. 14. Fictive episodes per minute as a function of artificial cerebrospinal fluid (aCSF) pH in
preparations taken from (A) normoxic normocapnic control frogs (n=8), (B) chronically
hypercapnic frogs (CHC; n=11) and (C) chronically hypoxic frogs (CH; n=7) prior to ACTZ
addition (open circles), following 1µM ACTZ addition (closed circles) and following 10µM
ACTZ addition (closed squares). The data are plotted as mean values ± 1 SEM. Letters (a, and b)
indicate a significant difference amongst pH levels in any one group. A number sign (#) indicates
significant differences between the Pre-ACTZ condition and 1 or 10µM ACTZ addition.
67
Nu
mb
er o
f F
icti
ve
Ep
iso
des
p
er M
inu
te
0
5
10
15
20
25
30
Nu
mb
er o
f F
icti
ve
Ep
iso
des
per
Min
ute
0
5
10
15
20
25
30
aCSF pH
7.6 7.8 8.0
Nu
mb
er o
f F
icti
ve
Ep
iso
des
per
Min
ute
0
5
10
15
20
25
30
A. Control
B. Chronic Hypercapnia
C. Chronic Hypoxia
Pre-ACTZ
Pre-ACTZ
Pre-ACTZ
1µM ACTZ
1µM ACTZ
1µM ACTZ
10 µM ACTZ
10 µM ACTZ
10 µM ACTZ
b
b
b
a, b
ab, # b
#
a
a
Figure 14
68
Fig. 15. Fictive breaths per episode as a function of artificial cerebrospinal fluid (aCSF) pH in
preparations taken from (A) normoxic normocapnic control frogs (n=8), (B) chronically
hypercapnic frogs (CHC; n=11) and (C) chronically hypoxic frogs (CH; n=7) prior to ACTZ
addition (open circles), following 1µM ACTZ addition (closed circles) and following 10µM
ACTZ addition (closed squares). The data are plotted as mean values ± 1 SEM. A number sign
(#) indicates significant differences between the Pre-ACTZ condition and 1 or 10µM ACTZ
addition. An ampersand (@) indicates a significant difference between the 1 and 10µM ACTZ
additions.
69
Nu
mb
er o
f F
icti
ve
Bre
ath
s
per
Ep
iso
de
0
1
2
3
4
5
6
Nu
mb
er o
f F
icti
ve
Bre
ath
s p
er E
pis
od
e
0
1
2
3
4
5
6
aCSF pH
7.6 7.8 8.0
Nu
mb
er o
f F
icti
ve
Bre
ath
s p
er E
pis
od
e
0
1
2
3
4
5
6
#
#, @
A. Control
B. Chronic Hypercapnic
C. Chronic Hypoxic
Figure 15
70
Fig. 16. Fictive breath duration (s) as a function of artificial cerebrospinal fluid (aCSF) pH in
preparations taken from (A) normoxic normocapnic control frogs (n=8), (B) chronically
hypercapnic frogs (CHC; n=11) and (C) chronically hypoxic frogs (CH; n=7) prior to ACTZ
addition (open circles), following 1µM ACTZ addition (closed circles) and following 10µM
ACTZ addition (closed squares). The data are plotted as mean values ± 1 SEM. Letters (a, and b)
indicate a significant difference amongst pH levels in any one group.
71
Fic
tiv
e B
reat
h D
ura
tio
n (
s)
0.1
0.2
0.3
0.4
0.5
Fic
tiv
e B
reat
h D
ura
tio
n (
s)
0.1
0.2
0.3
0.4
0.5
aCSF pH
7.5 7.6 7.7 7.8 7.9 8.0 8.1
Fic
tiv
e B
reat
h D
ura
tio
n (
s)
0.1
0.2
0.3
0.4
0.5
A. Control
B. Chronic Hypercapnia
C. Chronic Hypoxia
a
b b
aa, b
b
Pre-ACTZ
Pre-ACTZ
Pre-ACTZ1µM ACTZ
1µM ACTZ
1µM ACTZ
10µM ACTZ
10µM ACTZ
10µM ACTZ
Figure 16
72
Fig. 17. Integrated fictive breath area (V·s) as a function of artificial cerebrospinal fluid (aCSF)
pH in preparations taken from (A) normoxic normocapnic control frogs (n=8), (B) chronically
hypercapnic frogs (CHC; n=11) and (C) chronically hypoxic frogs (CH; n=7) prior to ACTZ
addition (open circles), following 1µM ACTZ addition (closed circles) and following 10µM
ACTZ addition (closed squares). The data are plotted as mean values ± 1 SEM. A number sign
(#) indicates significant differences between the Pre-ACTZ condition and 1 or 10µM ACTZ
addition.
73
Inte
gra
ted
Fic
tiv
e B
reat
h A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
0.016In
teg
rate
d F
icti
ve
Bre
ath
Are
a (V
·s)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
0.016
aCSF pH
7.6 7.8 8.0
Inte
gra
ted
Fic
tiv
e B
reat
h A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
0.016
A. Control
B. Chronic Hypercapnia
C. Chronic Hypoxia
Pre-ACTZ
Pre-ACTZ
Pre-ACTZ
1µM ACTZ
1µM ACTZ
1µM ACTZ
10µM ACTZ
10µM ACTZ
10µM ACTZ #
Figure 17
74
Fig. 18. Total fictive ventilation (V·s·min-1
) as a function of artificial cerebrospinal fluid (aCSF)
pH in preparations taken from (A) normoxic normocapnic control frogs (n=8), (B) chronically
hypercapnic frogs (CHC; n=11) and (C) chronically hypoxic frogs (CH; n=7) prior to ACTZ
addition (open circles), following 1µM ACTZ addition (closed circles) and following 10µM
ACTZ addition (closed squares). The data are plotted as mean values ± 1 SEM. Letters (a, and b)
indicate a significant difference amongst pH levels in any one group. A number sign (#) indicates
significant differences between the Pre-ACTZ condition and 1 or 10µM ACTZ addition.
75
To
tal
Fic
tiv
e V
enti
lati
on
(V
·s·m
in-1
)
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
To
tal
Fic
tiv
e V
enti
lati
on
(V
·s·m
in-1
)
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
aCSF pH
7.6 7.8 8.0
To
tal
Fic
tiv
e V
enti
lati
on
(V
·s·m
in-1
)
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
Pre-ACTZ
Pre-ACTZ
Pre-ACTZ
1µM ACTZ
1µM ACTZ
10µM ACTZ
10µM ACTZ
A. Control
B. Chronic Hypercapnia
C. Chronic Hypoxia
a
a, b
b
a
b
b
b
a
b
aa, #
b, #
b, # b, #a
abb
#
1µM ACTZ
10µM ACTZ
Figure 18
76
2.4 Discussion
2.4.1 Summary
The major observations of this study are: 1) Decreasing aCSF pH caused an increase in
total fictive ventilation (i.e., fictive breathing) in all groups. 2) The increase in total fictive
ventilation, with lowered pH, was due to increases in fictive breathing frequency not the
integrated area of the fictive breaths. 3) The increases in fictive breathing frequency were, for the
most part, due to increases in the number of fictive episodes per minute not the number of fictive
breaths per episode. 4) Chronic hypercapnia (CHC) caused an augmentation of fictive breathing.
This was mediated by increases in fictive breathing frequency via an increase in the number of
fictive episodes per minute. 5) Chronic hypoxia (CH) had no effect on fictive breathing. 6) The
CHC-induced increase in fictive breathing was abolished by ACTZ treatment. 7) There was a
non-significant trend for the higher dose of ACTZ to reduce fictive breathing in the CH group.
2.4.2. Effects of Altering aCSF pH
Total fictive ventilation (an index of overall breathing in these in vitro brainstem-spinal
cord preparations) was augmented by acidification of the superfusate bathing the control
preparations. This was due to an increase in fictive breathing frequency rather than the integrated
area of fictive breaths (an index of breath amplitude or volume) as aCSF pH was lowered. The
other components of fictive breathing (i.e., number of fictive episodes per minute, breaths per
episode, breath duration and amplitude) analysed in this study did not change in response to
acidification/alkalisation of the aCSF pH.
77
Chronically hypoxic (CH) brainstem-spinal cord preparations showed similar responses
to changes in aCSF pH. In other words, as aCSF pH was acidified, CH preparations exhibited an
augmentation in total fictive ventilation mediated by an augmentation in fictive breathing
frequency and no further changes in any other components of fictive breathing.
Chronically hypercapnic (CHC) brainstem-spinal cord preparations also exhibited a
similar change in total fictive ventilation and fictive breathing frequency as pH of the aCSF was
lowered. However, as seen in Gheshmy et al. (2006; 2007)’s results and the preliminary data
(Srivaratharajah and Reid, unpublished) presented in chapter 1, an increase in fictive breathing
frequency in CHC preparations was mediated by an increase in the number of fictive episodes
per minute as aCSF pH was lowered. The number of fictive breaths per episode, fictive breath
duration and integrated area of fictive breaths (i.e., amplitude) recorded from CHC preparations
remained unaltered in response to acidification/alkalisation of the superfusate (consistent with
earlier studies; Gheshmy et al., 2006; Srivaratharajah and Reid, unpublished).
2.4.3. Effects of Chronic Hypercapnia
Exposure to chronic hypercapnia (CHC) augmented fictive breathing at lowered aCSF
pH levels compared to the controls. This CHC-induced augmentation of fictive breathing is
consistent with previous studies from this laboratory conducted on a terrestrial amphibian species
(Bufo marinus, the cane toad; Gheshmy et al., 2006; 2007) as well as preliminary studies done
on leopard frog in vitro brainstem-spinal cord preparations (Srivaratharajah and Reid,
unpublished). As was the case in those studies, the augmentation of fictive breathing was
mediated by an augmentation in fictive breathing frequency rather than the integrated area of the
fictive breaths (an index of fictive breath amplitude or volume). In turn, the changes in fictive
78
breathing frequency were mediated by changes in the number of fictive breathing episodes per
minute rather than the number of fictive breaths per episode. Again, this is consistent with the
results from previous studies on leopard frogs (Srivaratharajah and Reid, unpublished) and cane
toads (Gheshmy et al., 2006; 2007).
2.4.4. Effects of Chronic Hypoxia
Chronic hypoxic (CH) exposure did not alter fictive breathing compared to control
preparations. However, there was a significant decrease in fictive breath duration and integrated
fictive breath area following CH compared to controls. This blunting of fictive breath duration
and amplitude were offset by a trend for fictive breathing frequency to increase, resulting in no
overall change in total fictive ventilation (i.e., fictive breathing) in brainstem-spinal cord
preparations taken from CH frogs. These results are different from those observed in previous
studies from this laboratory on a terrestrial anuran species (Bufo marinus; McAneney and Reid,
2007). McAneney and Reid reported a blunting of fictive breathing frequency following
exposure of cane toads to CH. One possible explanation for this difference is that, unlike the
terrestrial anuran species that must adapt to periods of hypoxia while overwintering in
underground burrows, leopard frogs, a semi-aquatic species that overwinters under ice-covered
bodies of water (Pinder et al., 1992; Hermes-Lima and Zenteno-Savín, 2002), are more tolerant
of hypoxia. Moreover, the current study exposed leopard frogs to terrestrial normobaric hypoxia.
Since aquatic species, to a great extent, rely on cutaneous gas exchange (Burggren and West,
1982), perhaps exposure to aquatic hypoxia and hypercapnia may have produced greater changes
in fictive breathing or changes more along the line of those observed in the cane toads. This will
be discussed further in chapter 5.
79
2.4.5. Effects of Acetazolamide (ACTZ)
Bath application of acetazolamide (ACTZ) was used to inhibit carbonic anhydrase (CA)
within the central respiratory-related pH/CO2 chemoreceptor cells. Inhibition of CA would be
expected to result in slower CO2 hydration and a subsequent slowdown in the rate of H+ ion
production and accumulation within these chemoreceptor cells. Intracellular pH (pHi) regulatory
mechanisms (i.e., Na+/H
+ exchangers and HCO3
-/Cl
- exchangers) that regulate acid-base
disturbances, are therefore able to compensate for the intracellular acidification by removing the
accumulated protons. Therefore, the significant drop in pHi that stimulates CO2 chemoreceptor
firing (when ACTZ has not been added) does not occur or occurs to lesser degree, leading to
absent or reduced chemosensitive neurotransmission and a subsequent blunted ventilatory
response. The effects of ACTZ addition in this study support this interpretation.
Addition of 1µM ACTZ to control brainstem-spinal cord preparations had no effect on
fictive breathing while addition of 10 µM ACTZ caused a significant reduction in fictive
breathing. The blunting of fictive breathing (i.e., total fictive ventilation) following addition of
10µM ACTZ was due to a reduction in the integrated area of fictive breaths (i.e., an index of
breath amplitude or volume) rather than fictive breathing frequency (which remained unaltered).
The observation that 10µM ACTZ caused an alteration (reduction) in fictive breathing in control
preparations supports the notion that CA is involved in central pH/CO2 chemoreception in this
species.
If the CHC-induced increase in fictive breathing frequency was caused by an up-
regulation of CA amount/activity of CA one might assume that preparations taken from frogs
exposed to CHC would be less-sensitive to ACTZ treatment because their chemoreceptors would
80
have greater amounts of CA. However, even though this was not observed, the data still support
the idea that CA is involved in the CHC-induced augmentation of fictive breathing in leopard
frogs since blockade of CA activity abolished that effect. In essence, the application of ACTZ to
preparations taken from animals exposed to CHC caused the CHC preparations (with ACTZ) to
respond in a very similar manner to the control preparations without ACTZ. Indeed the CHC
preparations with ACTZ were less responsive to changes in aCSF pH than were the control
preparations without ACTZ.
A previous study demonstrated that either olfactory denervation prior to exposure to CHC
or simultaneous exposure to hyperoxia and hypercapnia prevented the CHC-induced
augmentation in fictive breathing (Gheshmy et al, 2007). Although this has not yet been
investigated, it is likely that the altered afferent input, during exposure to CHC, from olfactory
and arterial chemoreceptors was in some way responsible for triggering the changes in CA that
led to the increase in fictive breathing. If this were the case, then olfactory denervation prior to
CHC or exposure to hyperoxic hypercapnia should prevent the effects of ACTZ on fictive
breathing seen in this current study.
Addition of ACTZ to brainstem-spinal cord preparations taken from frogs exposed to CH
resulted in different responses at the two different doses of ACTZ. At the lower concentration of
ACTZ (i.e., 1 µM), there was a tendency for fictive breathing frequency and total fictive
ventilation to increase although these increases were small and not statistically-significant. This
was mediated by an increase in the number of breaths per episode at aCSF pH 7.8. However,
with the addition of 10µM ACTZ to the superfusate of these preparations, fictive breathing
showed a tendency to decrease. In other words, the response, to ACTZ, in the preparations from
81
animals exposed to CH was similar to the response in the control and CHC groups although the
changes in the CH group did not reach statistical significance. The observation that there was no
statistically-significant change in fictive breathing with the high dose of ACTZ suggests that
exposure of leopard frogs to CH has not led to a change in CA activity/amount. It is possible that
the CH-induced decrease in fictive breathing seen in cane toads (McAneney and Reid, 2007) was
due to a down-regulation/decrease in CA amount/activity. If this were the case then the effects of
CH on CA are different in these two species (i.e., a down-regulation/reduction in toads and no
change in frogs). However, this remains speculative as neither the olfactory denervation nor
hyperoxic hypercapnia experiments have been performed on frogs and similarly, the ACTZ
experiments have yet to be performed on cane toads.
2.4.6 In Vivo Versus In Vitro Effects of Acetazolamide (ACTZ)
The results of this study are different from those of focal acidification studies in which
ACTZ was applied to a localized region of the brain in vivo. In vivo microinjection studies use
ACTZ to induce focal acidification of tissue by inhibition of CA and subsequent slowing of the
dehydration reaction whereby HCO3- and H
+ form CO2. This slows CO2 removal from tissue
resulting in acidification. Studies have shown that such in vivo ACTZ microinjections increase
ventilatory responses in rats (Xu et al., 2001; Nattie and Li, 1996). As argued by Necakov et al.
(2002), the in vivo and in vitro situations are quite different. In vivo, tissue pH is affected by both
the metabolic production and elimination of H+. CA is involved in both H
+ production from the
hydration of CO2 in tissue and the elimination of CO2 at the lungs by combining H+ and HCO3
-.
Hence, addition of ACTZ will not only slow down H+ production but will also slow down the
82
excretion of CO2 at the lungs. Subsequent accumulation of CO2 in the blood would ultimately
lead to increases in intracellular H+, which would stimulate pH/CO2 chemoreceptors. This
sequence of reactions could possibly explain the in vivo hyperpnea following ACTZ
administration.
In vitro, however, tissue pH no longer relies on the role of CA in the elimination of CO2
but on aCSF flow rate. Due to the short diffusion distance across frog brain tissue and a high
superfusion rate (10 ml/min), the tissue pH is dependent on the pH of the superfusion medium
which in turn depends on the amount of CO2 gassed into it. Hence, the results from this in vitro
bath application study are not comparable to in vitro focal acidification studies (although the
same agent, ACTZ, is used in both).
In vitro studies using bath application of ACTZ are not consistent either. Taylor et al.
(2003) found that bath application of 25µM ACTZ to adult bullfrog (Rana catesbeiana) in vitro
brainstem-spinal cord preparations resulted in increased fictive breathing. On the other hand,
Erlichman and colleagues (1994) found that bath application of ACTZ to pulmonate snail in vitro
brain-pneumostome preparations increased ventilation during normocapnia but slowed
ventilatory responses to rapid changes in CO2. Finally, Necakov and colleagues (2002) found no
changes in fictive breathing following addition of ACTZ to the superfusate of transverse rat
brainstem slice preparations. Whether these differences reflect differences in chemosensory
processes within these different species is unclear. Nevertheless, the results from this current
study clearly show the importance of CA, not only in central CO2 chemosensitivity, but also in
the CHC-induced increase in fictive breathing in leopard frogs.
83
2.5 Conclusion
The results of this study show that acetazolamide (ACTZ) addition reduces fictive
breathing in preparations taken from control and chronically hypercapnic (CHC) animals but
generally has no significant effect on preparations taken from chronically hypoxic (CH) animals.
Preparations taken from CHC animals were affected to a greater extent than control preparations.
Therefore, I conclude that carbonic anhydrase (CA) plays an important role in the signal
transduction pathway leading to the CHC-induced augmentation in fictive breathing. However,
whether this CHC-induced augmentation in fictive breathing is due to an increase in the amount
or activity of CA available for hydration of CO2 cannot be determined from these results.
Furthermore, since ACTZ is a cell permeant CA inhibitor, it will inhibit both intracellular and
extracellular CA. The results from this study therefore, cannot distinguish whether the changes in
amount/activity were that of intracellular or extracellular CA. The following chapters will delve
further into the role of CA in the modulation of fictive breathing in CHC compared to control
and CH frogs and address these aforementioned issues (i.e., intracellular vs. extracellular and
amount vs. activity).
84
CHAPTER 3
EFFECTS OF EXOGENOUS CARBONIC ANHYDRASE APPLICATION ON FICTIVE
BREATHING IN ISOLATED IN VITRO BRAINSTEM-SPINAL CORD PREPARATIONS
TAKEN FROM CHRONICALLY HYPOXIC AND HYPERCAPNIC LEOPARD FROGS
(RANA PIPIENS)
85
3.1 INTRODUCTION
The previous series of experiments (chapter 2) illustrated that the chronic hypercapnia
(CHC)-induced increase in fictive breathing was abolished by treatment with acetazolamide
(ACTZ). This suggests that the CHC-induced increase in fictive breathing was caused, at least in
part, by an increase in the activity/amount of carbonic anhydrase (CA), presumably within the
chemoreceptor cells. ACTZ is a cell permeant inhibitor of CA and therefore will inhibit both
extracellular and intracellular CA. However, it is reasonable to assume that the majority of CA in
the brain is intracellular rather than extracellular suggesting that the effects of ACTZ in chapter 2
were due, primarily, to its inhibition of intracellular CA (Ritucci et al., 1997; 1998; Wang et al.,
2002; Putnam et al., 2004). This would be consistent with the model of cellular CO2
chemoreception outlined in chapter 1.
Ideally, the application of an extracellular CA inhibitor (i.e., a CA inhibitor that is cell
impermeant) could be used to determine if intracellular or extracellular CA was involved in the
CHC-induced increase in fictive breathing. One such inhibitor is benzolamide. However,
benzolamide is not commercially available and has to be synthesised at considerable cost. Given
this, I took a different approach to determine whether the changes following CHC were due to
changes in intracellular or extracellular CA. In these experiments, exogenous CA was added to
the aCSF bathing the in vitro brainstem-spinal cord preparations. Given its low permeability, this
exogenous CA would be expected to remain in the extracellular domain and not enter the cells. I
hypothesised that the CHC-induced changes in fictive breathing were due to changes in
intracellular CA function. Based on this, I predicted that application of exogenous CA would
either have no effect on fictive breathing or may even reduce fictive breathing. The last
86
prediction is based on the assumption that extracellular CA would reduce the availability of CO2
to diffuse into the chemoreceptor cells by converting it to H+ and HCO3
- ions outside of the cell.
87
3.2 MATERIALS & METHODS
3.2.1 Experimental Animals
Leopard frogs (Rana pipiens; N = 24; approximately 5 to 8 cm in length) were obtained
from a commercial supplier (Boreal Scientific, St. Catharine’s, Ontario) and housed under
conditions identical to those previously described in Chapter 2.
3.2.2 Exposure to Chronic Hypoxia and Hypercapnia
Leopard frogs (Rana pipiens) were exposed to control (N = 8) chronic hypoxia (CH; N =
6) and chronic hypercapnia (CHC; N = 10) as described in Chapter 2.
3.2.3 In Vitro Brainstem-Spinal Cord Preparations
In vitro brainstem-spinal cord preparations were prepared from control (chronically
normoxic normocapnic), chronically hypercapnic and chronically hypoxic frogs as described in
Chapter 2.
3.2.4 Carbonic Anhydrase
Bovine carbonic anhydrase (CA; 4688 W/A units/mg; Sigma-Aldrich Inc., Oakville,
Ontario, Canada) was used at a concentration of 10mg/L. This particular concentration was
chosen on the basis of a previous study by Huang et al. (1995) which showed that exogenous CA
at this concentration had an effect on extracellular pH shifts in hippocampal rat brain slices and
no further effects were noticed at 100mg/L.
88
3.2.5 Experimental Protocol
Following a 1 hour stabilisation period at room temperature and an aCSF pH of 7.8 (see
chapter 2), recordings, from cnV and cnX, were made prior to CA addition at aCSF pH levels of
7.6, 7.8 and 8.0. The pH changes were made as described in chapter 2. The preparations were
then superfused with aCSF containing 10 mg/L CA. Following a 30 min stabilisation period, the
pH changes were repeated in random order as described in chapter 2. Following the recording
periods with CA in the aCSF, the preparations were superfused, for 30 min, with aCSF
containing no CA. At the end of this 30 min washout period, the aCSF pH changes were made
once again and fictive breathing was recorded as described in chapter 2.
3.2.6 Data Analysis & Statistics
Data were recorded using the BIOPAC MP150 in conjunction with the AcqKnowledge
3.7.3 software. Data were analysed for the last 10 min of each recording period at each aCSF pH
level. The values are reported as the mean ± one S.E.M. The same components of fictive
breathing were measured as described in chapter 2. The effects of changing aCSF pH in any
given group (i.e., control, CHC or CH), were analysed using a one-way repeated measures
ANOVA followed by a SNK multiple comparison test. The effects of CHC and CH were
analysed using a two-way non-repeated measures ANOVA (control/CHC or control/CH x aCSF
pH) followed by a SNK multiple comparison test. The effects of CA addition in any given group
(control, CHC or CH) were analysed using a two-way repeated measures ANOVA (aCSF pH X
pre-CA/CA/washout) followed by a SNK multiple comparison test. In all cases, p < 0.05 was
taken to be the limit of statistical significance.
89
3.3 RESULTS
3.3.1 Effects of Chronic Hypercapnia
In both the control and chronically hypercapnic (CHC) groups, fictive breathing
frequency (Fig. 19A) was elevated, compared to pH 8.0, as the aCSF pH level was reduced to 7.6
(control: p= 0.031; CHC: p= 0.025). Fictive breathing frequency was significantly greater in the
CHC preparations compared to controls at aCSF pH 7.6 (p= 0.001) and 7.8 (p= 0.026).
The two components of fictive breathing frequency are the number of fictive episodes per
min (Fig. 19B) and the number of fictive breaths per episode (Fig. 19C). Neither of these
variables were altered as pH was lowered from 8.0 to 7.6 (episodes/minute: control, p= 0.533;
CHC, p= 0.515; breaths/episode: control, p= 0.0587; CHC, p= 0.285). The number of fictive
episodes per minute was significantly greater in the CHC group compared to the controls at
aCSF pH levels of 7.6 (p= 0.007) and 7.8 (p= 0.019). The number of fictive breaths per episode
did not change following CHC acclimatization (p = 0.258).
In both groups, fictive breath duration was unaltered by changes in aCSF pH (Fig 19D;
controls, p = 0.518; CHC, p = 0.498). Compared to controls, CHC did not change fictive breath
duration at any of the aCSF pH levels (p= 0.636). The integrated area of the fictive breaths (Fig.
19E) decreased as aCSF pH was lowered from 7.8 to 7.6 (p= 0.027) in the control group but
remained unaltered by changing pH in the CHC group (p = 0.957). The integrated area of the
fictive breaths was not significantly altered by exposure to CHC (p = 0.117).
Total fictive ventilation remained unaltered in both CHC and control groups following
acidification of the aCSF from pH 8.0 to pH 7.6 (control: p= 0.227; CHC: p= 0.183). However,
90
CHC exposure increased total fictive ventilation at aCSF pH 7.6 over the corresponding control
value (p= 0.006).
91
Fig. 19. Fictive breathing frequency (A), the number of fictive episodes per minute (B), the
number of fictive breaths per episode (C), fictive breath duration (D), integrated fictive breath
area (E) and total fictive ventilation (F), as a function of artificial cerebrospinal fluid (aCSF) pH,
recorded from in vitro brainstem-spinal cord preparations taken from normoxic normocapnic
control (n=10; open circles) and chronically hypercapnic (CHC; n=8; closed squares) leopard
frogs. The data are plotted as mean values ± 1 SEM. Letters (a and b) indicate a significant
difference amongst pH levels in any one group. A plus sign (+) indicates a significant difference
between the CHC and control group at any given pH.
92
Fic
tiv
e B
reat
hin
g F
requ
ency
(bre
ath
s·m
in-1
)
0
10
20
30Chronic Hypercapnia
Control
a
a, b, +
b, +
aa,bb
A. Fictive Breathing Frequency
B. Fictive Episodes per Minute
Nu
mber
of
Fic
tiv
e E
pis
odes
per
Min
ute
0
5
10
15
20
25
30
Control
Chronic Hypercapnia
+
+
C. Fictive Breaths per Episode
aCSF pH
7.6 7.8 8.0
Nu
mb
er o
f F
icti
ve
Bre
aths
per
Epis
ode
0
2
4
6
8
Control
Chronic Hypercapnia
Fic
tive
Bre
ath D
ura
tio
n (
s)
0.10
0.15
0.20
0.25
0.30
0.35
0.40
D. Fictive Breath Duration
Control
Chronic Hypercapnia
Inte
gra
ted
Fic
tiv
e B
reat
h A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
Control
Chronic Hypercapnia
E. Integrated Fictive Breath Area
aa
b
aCSF pH
7.6 7.8 8.0
Tota
l F
icti
ve
Ven
tila
tion
(V
·s·m
in-1
)
0.00
0.05
0.10
0.15
0.20
Control
Chronic Hypercapnia
F. Total Fictive Ventilation
+
Figure 19
93
3.3.2 Effects of Chronic Hypoxia
Note, the control data plotted in Figure 20 is the same as the control data plotted in Figure
19. The effects of pH on fictive breathing in this group were described in the section above. In
preparations taken from chronically hypoxic frogs, fictive breathing frequency (Fig.20A) was
elevated at aCSF pH 7.6 and 7.8 compared to pH 8.0 (p= 0.029). Moreover, fictive breathing
frequency was significantly higher following CH exposure at aCSF pH 7.6 compared to controls
(p= 0.005). The two components of fictive breathing frequency are the number of fictive
episodes per minute (Fig.20B) and the number of fictive breaths per episode (Fig. 20C). Neither
of these variables were altered by changes in pH (episodes per minute: control, p= 0.533; CH,
p= 0.101; breaths per episode: control, p= 0.587; CH, p= 0.264). The number of fictive episodes
per min was, however, higher in the CH group compared to controls at aCSF pH 7.6 (p= 0.042),
while breaths per episode remained unaltered following CH exposure (p= 0.524).
In the CH group, fictive breath duration (Fig. 20D) did not change as aCSF pH was
lowered (p=0.07). CH exposure did not alter fictive breath duration compared to the controls (p=
0.830). The integrated area of fictive breaths (Fig. 20E) was not altered by changes to aCSF pH
(p= 0.084) in the CH group nor following CH exposure when compared to controls (p= 0.320).
Total fictive ventilation (Fig. 20F) remained unaltered in response to changes in aCSF pH within
the CH group (p= 0.173) as well as following CH exposure compared to controls (p= 0.092).
94
Fig. 20. Fictive breathing frequency (A), the number of fictive episodes per minute (B), the
number of fictive breaths per episode (C), fictive breath duration (D), integrated fictive breath
area (E) and total fictive ventilation (F), as a function of artificial cerebrospinal fluid (aCSF) pH,
recorded from in vitro brainstem-spinal cord preparations taken from normoxic normocapnic
control (n=8; open circles) and chronically hypercapnic (CH; n=10; closed circles) leopard frogs.
The data are plotted as mean values ± 1 SEM. Letters (a and b) indicate a significant difference
amongst pH levels in any one group. A plus sign (+) indicates a significant difference between
the CH and control groups at any given pH.
95
Fic
tive
Bre
athin
g F
requen
cy
(bre
aths·
min
-1)
0
10
20
30
Control
ab
b, +
Chronic Hypoxia
aa,bb
A. Fictive Breathing Frequency
Num
ber
of
Fic
tive
Ep
iso
des
per
Min
ute
0
5
10
15
20
25
30
Control
Chronic Hypoxia
+
B. Fictive Episodes per Minute
aCSF pH
7.6 7.8 8.0
Num
ber
of
Fic
tive
Bre
aths
per
Ep
isod
e
0
2
4
6
8
Control
Chronic Hypoxia
Fic
tiv
e B
reat
h D
ura
tion
(s)
0.10
0.15
0.20
0.25
0.30
0.35
0.40
Control
Chronic Hypoxia
D. Fictive Breath Duration
Inte
gra
ted
Fic
tive
Bre
ath A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
Control
Chronic Hypoxia
aa
b
E. Integrated Fictive Breath Area
aCSF pH
7.6 7.8 8.0
Tota
l F
icti
ve
Ven
tila
tion (
V·s
·min
-1)
0.00
0.05
0.10
0.15
0.20
Control
Chronic Hypoxia
F. Total Fictive VentilationC. Fictive Breaths per Episode
Figure 20
96
3.3.3 Effect of Exogenous Carbonic Anhydrase Application
Fictive Breathing Frequency: Exogenous bovine carbonic anhydrase (CA) addition
reduced fictive breathing frequency at aCSF pH 7.6 in preparations taken from control frogs
(Fig. 21A; p= 0.023). However, this effect was reversible and fictive breathing frequency
returned to pre-CA addition levels following the 30 min washout period. In preparations taken
from chronically hypercapnic (CHC; Fig. 21B) and chronically hypoxic (CH; Fig 21C) animals,
however, addition of CA did not alter fictive breathing frequency (CHC: p= 0.161; CH: p=
0.270).
Fictive Episodes per Minute: Bath application of bovine CA did not significantly affect
the number of fictive episodes per minute in preparations taken from control (Fig. 22A;
p=0.307), chronically hypercapnic (CHC; Fig 22B; p= 0.547) and chronically hypoxic (CH; Fig.
22C; p= 0.222) frogs.
Fictive Breaths per Episode: In the control (Fig. 23A), CHC (Fig. 23B) and CH (Fig.
23C) groups, there was no affect of CA addition on the number of fictive episodes per minute
(controls: p= 0.161; CHC: p= 0.093; CH: p= 0.852).
Fictive Breath Duration: CA bath application did not have any effect on fictive breath
duration in preparations taken from control (Fig. 24A; p = 0.067), chronically hypercapnic (Fig.
24B; p= 0.621) and chronically hypoxic (Fig. 24C; p= 0.761) frogs.
Integrated Fictive Breath Area: In the control group (Fig. 25A) CA addition had no
effect on integrated fictive breath area (p= 0.532). However, when comparing data after washout
of CA to that of pre-CA addition, integrated fictive breath area was significantly reduced in the
97
control group at aCSF pH 7.8 (p= 0.007) and 8.0 (p= 0.039). In addition, after washout of CA,
integrative fictive breath area was also significantly reduced at aCSF pH 7.6 (p= 0.021) and 7.8
(p= 0.004) compared to data following CA addition to the superfusate of control preparations.
Integrative fictive breath area remained unaltered following CA addition and following washout
of CA in the preparations taken from chronically hypercapnic (Fig. 25B; p= 0.911) and
chronically hypoxic (Fig. 25C; p= 0.856) frogs.
Total Fictive Ventilation: In the control group (Fig. 26A) CA addition had no effect on
total fictive ventilation (p = 0.095). Similarly, no effects on total fictive ventilation were
observed following CA addition in the chronically hypercapnic (Fig 26B; p= 0.109) and
chronically hypoxic (Fig 26C; p=0.266) groups.
98
Fig. 21. Fictive breathing frequency (breaths·min-1
) as a function of artificial cerebrospinal fluid
(aCSF) pH in preparations taken from (A) normoxic normocapnic control frogs (n=10), (B)
chronically hypercapnic frogs (CHC; n=8) and (C) chronically hypoxic (CH; n=6) frogs prior to
carbonic anhydrase (CA) addition (open circles), following CA addition (closed circles) and
following washout of CA (closed squares). The data are plotted as mean values ± 1 SEM. Letters
(a, and b) indicate a significant difference amongst pH levels in any one group. A number sign
(#) indicates significant differences between Pre-CA addition and CA addition or after CA
washout addition.
99
Fic
tive
Bre
ath
ing
Fre
quen
cy
(bre
ath
s·m
in-1
)
0
10
20
30
Fic
tive
Bre
athin
g F
req
uen
cy
(bre
ath
s·m
in-1
)
0
10
20
30
aCSF pH
7.6 7.8 8.0
Fic
tiv
e B
reat
hin
g F
requ
ency
(bre
aths·
min
-1)
0
10
20
30
Pre-CA
Pre-CA
Pre-CA
CA addition
CA addition
CA addition
After CA washout
After CA washout
After CA washout
A. Control
#
aa, bb
a, b
b
a
ab
b
B. Chronic Hypercapnia
C. Chronic Hypoxia
Figure 21
100
Fig. 22. Fictive episodes per minute as a function of artificial cerebrospinal fluid (aCSF) pH in
preparations taken from (A) normoxic normocapnic control frogs (n=10), (B) chronically
hypercapnic frogs (CHC; n=8) and (C) chronically hypoxic (CH; n=6) frogs prior to carbonic
anhydrase (CA) addition (open circles), following CA addition (closed circles) and following
washout of CA (closed squares). The data are plotted as mean values ± 1 SEM.
101
A. Control
Nu
mb
er
of
Fic
tiv
e E
pis
od
es
per
Min
ute
0
5
10
15
20
25
30
Nu
mb
er o
f F
icti
ve
Ep
iso
des
per
Min
ute
0
5
10
15
20
25
30
aCSF pH
7.6 7.8 8.0
Nu
mb
er o
f F
icti
ve
Ep
iso
des
per
Min
ute
0
5
10
15
20
25
30
Pre-CA
Pre-CA
Pre-CA
CA addition
CA addition
CA addition
After CA washout
After CA washout
After CA washout
B. Chronic Hypercapnia
C. Chronic Hypoxia
Figure 22
102
Fig.23. Fictive breaths per episode as a function of artificial cerebrospinal fluid (aCSF) pH in
preparations taken from (A) normoxic normocapnic control frogs (n=10), (B) chronically
hypercapnic frogs (CHC; n=8) and (C) chronically hypoxic (CH; n=6) frogs prior to carbonic
anhydrase (CA) addition (open circles), following CA addition (closed circles) and following
washout of CA (closed squares). The data are plotted as mean values ± 1 SEM.
103
A. Control
Num
ber
of
Fic
tive
Bre
aths
per
Epis
ode
0
2
4
6
8
Nu
mber
of
Fic
tive
Bre
ath
s
per
Ep
isod
e
0
2
4
6
8
aCSF pH
7.6 7.8 8.0
Num
ber
of
Fic
tive
Bre
aths
per
Epis
ode
0
2
4
6
8
Pre-CA
Pre-CA
Pre-CA
CA addition
CA addition
CA addition
After CA washout
After CA washout
After CA washout
B. Chronic Hypercapnia
C. Chronic Hypoxia
Figure 23
104
Fig. 24. Fictive breath duration (s) as a function of artificial cerebrospinal fluid (aCSF) pH in
preparations taken from (A) normoxic normocapnic control frogs (n=10), (B) chronically
hypercapnic frogs (CHC; n=8) and (C) chronically hypoxic (CH; n=6) frogs prior to carbonic
anhydrase (CA) addition (open circles), following CA addition (closed circles) and following
washout of CA (closed squares). The data are plotted as mean values ± 1 SEM.
105
Fic
tiv
e B
reat
h D
ura
tio
n (
s)
0.10
0.15
0.20
0.25
0.30
0.35
0.40
Fic
tiv
e B
reat
h D
ura
tio
n (
s)
0.10
0.15
0.20
0.25
0.30
0.35
0.40
aCSF pH
7.6 7.8 8.0
Fic
tiv
e B
reat
h D
ura
tio
n (
s)
0.10
0.15
0.20
0.25
0.30
0.35
0.40
A. Control
Pre-CA
Pre-CA
Pre-CACA addition
CA addition
CA addition
After CA washout
After CA washout
After CA washout
B. Chronic Hypercapnia
C. Chronic Hypoxia
Figure 24
106
Fig. 25. Integrated fictive breath area (V·s) as a function of artificial cerebrospinal fluid (aCSF)
pH in preparations taken from (A) normoxic normocapnic control frogs (n=10), (B) chronically
hypercapnic frogs (CHC; n=8) and (C) chronically hypoxic (CH; n=6) frogs prior to carbonic
anhydrase (CA) addition (open circles), following CA addition (closed circles) and following
washout of CA (closed squares). The data are plotted as mean values ± 1 SEM. Letters (a, and b)
indicate a significant difference amongst pH levels in any one group. A number sign (#) indicates
significant differences between Pre-CA addition and CA addition or after CA washout addition.
An ampersand (@) indicates a significant difference between the CA addition and CA washout
conditions.
107
Inte
gra
ted
Fic
tiv
e B
reat
h A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
Inte
gra
ted
Fic
tiv
e B
reat
h A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
aCSF pH
7.6 7.8 8.0
Inte
gra
ted
Fic
tiv
e B
reat
h A
rea
(V·s
)
0.000
0.002
0.004
0.006
0.008
0.010
0.012
0.014
Pre-CA
Pre-CA
Pre-CA
CA addition
CA addition
CA addition
After CA washout
After CA washout
After CA washoutb
a a
A. Control
#, @ #@
B. Chronic Hypercapnia
C. Chronic Hypoxia
Figure 25
108
Fig. 26. Total fictive ventilation (V·s·min-1
) as a function of artificial cerebrospinal fluid (aCSF)
pH in preparations taken from (A) normoxic normocapnic control frogs (n=10), (B) chronically
hypercapnic frogs (CHC; n=8) and (C) chronically hypoxic (CH; n=6) frogs prior to carbonic
anhydrase (CA) addition (open circles), following CA addition (closed circles) and following
washout of CA (closed squares). The data are plotted as mean values ± 1 SEM. Letters (a, and b)
indicate a significant difference amongst pH levels in any one group.
109
To
tal
Fic
tiv
e V
enti
lati
on
(V
·s·m
in-1
)
0.00
0.05
0.10
0.15
0.20
To
tal
Fic
tiv
e V
enti
lati
on
(V
·s·m
in-1
)
0.00
0.05
0.10
0.15
0.20
aCSF pH
7.6 7.8 8.0
To
tal
Fic
tiv
e V
enti
lati
on
(V
·s·m
in-1
)
0.00
0.05
0.10
0.15
0.20
Pre-CA
Pre-CA
CA addition
CA addition
After CA washout
After CA washoutaa, b
b
A. Control
Pre-CA
CA additionAfter CA washout
a, b b
a
B. Chronic Hypercapnia
C. Chronic Hypoxia
Figure 26
110
3.4 Discussion
3.4.1 Goals and Predictions
The results of chapter 2 indicated that the CHC-induced increase in fictive breathing was
abolished by treatment with acetazolamide (ACTZ) which is a cell permeant inhibitor of
carbonic anhydrase. Those results suggest that during CHC, the activity or amount of CA is up-
regulated or increased and that it is this change in CA that accounts for the CHC-induced
increase in fictive breathing. The primary goal of the experiments in this current chapter was to
determine whether the CHC-induced augmentation of fictive breathing was due to changes in
intracellular or extracellular carbonic anhydrase (CA). In order to do this, exogenous CA was
added to the aCSF bathing the in vitro brainstem-spinal cord preparations. Given its low
permeability, this exogenous CA would be expected to remain in the extracellular domain and
not enter the cells. The hypothesis was that the CHC-induced changes in fictive breathing were
due to changes in intracellular CA amount/activity. Based on this, one would predict that
application of exogenous CA would either have no effect on fictive breathing or may even
reduce fictive breathing. The last prediction is based on the assumption that extracellular CA
would reduce the availability of CO2 to diffuse into the chemoreceptor cells by converting it to
H+ and HCO3
- ions outside of the cell.
3.4.2 The Effects of Chronic Hypercapnia
As seen in the previous chapter, exposure to chronic hypercapnia (CHC) augmented
fictive breathing at the lower aCSF pH levels (7.8 and 7.6). This CHC-induced augmentation is
111
consistent with previous studies from this laboratory conducted on a terrestrial amphibian species
(Bufo marinus, the cane toad; Gheshmy et al., 2006; 2007) as well as preliminary studies done
on leopard frogs (Srivaratharajah and Reid, unpublished). The CHC-induced augmentation of
fictive breathing was mediated by an increase in fictive breathing frequency rather than
integrated area of fictive breaths (an index of fictive breath amplitude). In turn, the changes in
fictive breathing frequency were mediated by changes in the number of fictive episodes per
minute rather than fictive breaths per episode. These results are also consistent with those from
previous studies on leopard frogs (chapter 2; Srivaratharajah and Reid, unpublished) and cane
toads (Gheshmy et al., 2006; 2007).
3.4.3 The Effects of Chronic Hypoxia
Exposure to chronic hypoxia (CH) did not alter total fictive ventilation compared to
control preparations. However, there was a CH-induced increase in fictive breathing frequency,
at pH 7.6, mediated by an increase in the number of fictive episodes per minute. These results are
slightly different, although generally consistent, with the results seen in chapter 2. In that
chapter, exposure to chronic hypoxia did not alter total fictive ventilation (the same result as in
this chapter) but there was a non-significant trend for CH to augment fictive breathing frequency.
As such, the effects of CH on fictive breathing frequency were generally the same in chapters 2
and 3 although the increase at pH 7.6 was statistically significant in chapter 3 but not in chapter
2.
112
The effects of CH on fictive breathing frequency in this chapter are also different from
those observed in a previous study from this laboratory on a terrestrial anuran species (the cane
toad, Bufo marinus; McAneney and Reid, 2007). McAneney and Reid (2007) reported a
reduction of fictive breathing frequency following exposure to CH in cane toads. As discussed in
chapter 2 (and later discussed in chapter 5), this difference may in fact be due to greater hypoxia
tolerance and lower cutaneous O2 and CO2 exchange capabilities during terrestrial hypoxia in
leopard frogs.
3.4.4 Effects of Carbonic Anhydrase (CA) Addition
Addition of carbonic anhydrase (CA) to the superfusate of brainstem-spinal cord
preparations taken from control leopard frogs had no effect on overall total fictive ventilation in
any of the groups (control; CHC; CH) although there was a non-statistically significant trend for
total fictive ventilation to decrease at pH 7.6 in the CHC and CH groups. This trend was the
result of a similar trend for fictive breathing frequency to be reduced at pH 7.6 in all groups.
Indeed in the control group, but not the CHC or CH groups, fictive breathing frequency was
significantly reduced at pH 7.6. These results support my hypothesis that addition of
extracellular CA would either not change or reduce fictive breathing in these preparations. Given
this, the results of this chapter support the contention that the majority of endogenous CA in the
brain is intracellular and that the ACTZ treatment in chapter 2 exerted its effects via inhibition
primarily of intracellular rather than extracellular CA.
113
Assuming that intracellular and, not extracellular, CA is involved in the CO2 transduction
pathway responsible for respiratory-related central pH/CO2 chemoreception (Ritucci et al., 1997;
1998; Wang et al., 2002) and that changes in intracellular CA are responsible, at least in part, for
the CHC-induced augmentation in fictive breathing, addition of extracellular CA would not have
been expected to have any effect on fictive breathing. Indeed, the addition of exogenous CA
could be expected to reduce rather than augment fictive breathing as exogenously added CA may
disrupt the diffusion of CO2 into chemoreceptor cells by catalyzing the production of H+ and
HCO3- from CO2 in the superfusate. As a result of this, less CO2 would be available to diffuse
across the cell membrane into central CO2 chemoreceptor cells, hindering or attenuating the
signal transduction pathway leading to CO2 chemoreceptor firing as described in chapter 1. The
results of this study suggest that this has occurred. Exogenous CA had no statistically-significant
effect on total fictive breathing although there was a trend for CA to cause it to decrease at the
lower pH levels. This is consistent with the notion that the exogenous CA had disrupted the
diffusion of CO2 into the chemoreceptor cells.
If extracellular CA was important for central respiratory-related pH/CO2 chemoreception
or the CHC-induced augmentation of fictive breathing, one would expect that preparations taken
from control frogs would exhibit an augmentation of fictive breathing following the addition of
exogenous CA. In other words, addition of CA should cause a control preparation to mimic the
increase in fictive breathing observed following CHC without any addition of CA to the
superfusate. However, this was not the case, as in the control group, fictive breathing frequency
was actually reduced upon addition of CA and overall fictive breathing remained unaltered.
114
One may argue that since CA IV isozymes are tethered to the plasma membrane of
respective cells via a glycosyl-phosphatidyl-inositol (GPI) linkage (Brown and Waneck, 1992),
the addition of free-floating exogenous CA may not mimic the natural distribution of this
enzyme. In fact, those who support the notion that extracellular pH shifts are the stimulus for
pH/CO2 chemoreception may argue that the GPI linkage serves as an activator of downstream
ion-channels that in turn covert the extracellular change in pH into an influx of Ca2+
and
subsequent signal transduction. If this were the case, addition of free floating CA would not have
the same effect as anchored CA in activating central CO2 chemoreceptor cells. Our results cannot
discount this point. However, on the basis of parsimony, it would seem that the effects of CA on
respiratory modulation following CHC and CH are predominately due to intracellular isozymes
(particularly CA-II like isozymes).
3.5 Conclusion
The results of this chapter suggest, but cannot definitively confirm, that extracellular CA
is not involved in central pH/CO2 chemoreception or the CHC-induced augmentation of fictive
breathing. However, based upon the results of chapter 2 and this chapter, I suggest that it is
predominately an intracellular CA II-like isozyme that is responsible, at least in part, for both
central pH/CO2 chemosensitivity and the CHC-induced augmentation of fictive breathing.
Further investigation is warranted in order to support the distinction between the involvement of
intracellular and extracellular CA. The use of extracellular CA inhibitors (i.e., relatively cell
impermeable inhibitors such as benzolamide) to the superfusate of in vitro brainstem-spinal cord
preparations of leopard frogs would provide further insight into this issue. In addition,
115
application of phosphatidylinositol-specific phospholipase C to the superfusate, which will
cleave the glycosyl-phosphatidyl-inositol anchor of existing extracellular CA isozymes (Zhu and
Sly, 1990; Tong et al., 2000; Sharom and Lehto, 2002; Svichar et al., 2006) would also be useful
in distinguishing between CA II-like and CA IV-like isozymes present in frog neural tissue. The
following chapter presents the results of a histochemical analysis of the location of active CA
(whether it is intracellular or extracellular).
116
CHAPTER 4
HISTOCHEMICAL ANALYSIS OF ACTIVE CARBONIC ANHYDRASE IN BRAINSTEMS
TAKEN FROM CONTROL, CHRONICALLY HYPERCAPNIC AND CHRONICALLY
HYPOXIC LEOPARD FROGS
117
4.1 INTRODUCTION
The results of chapter 2, as well as a number of studies in the literature (Erlichman et al.,
1994; Wang et al., 2002; Taylor et al, 2003) indicate that carbonic anhydrase (CA) is important
in the putative mechanism underlying respiratory-related central pH/CO2 chemoreception. Given
this, the goal of the current chapter was to use a histochemical approach to try and identify active
CA within the leopard frog brainstem and to qualitatively assess whether exposure to chronic
hypercapnia (CHC) or chronic hypoxia (CH) alters the amount of CA present.
The first enzymatic histochemical method used to localize carbonic anhydrase in tissue
was performed by Häusler (in 1958) with further modifications by Hansson (1967) and
Ridderstråle (1976; 1980; Lönnerholm and Ridderstråle, 1974). This histochemical method is
referred to as the cobalt-phosphate method and involves chemical reactions that ultimately
produce a black precipitate in and around regions of active tissue CA. The reaction medium
contains cobalt sulphate (CoSO4), potassium dihydrogen phosphate (KH2PO4), sulphuric acid
(H2SO4) and sodium bicarbonate (NaHCO3). Although the intermediate complexes formed
during the reaction of the previously mentioned chemicals have not been clearly identified or
isolated, equations 3A and 3B express complexes mentioned by Maren (1980). Equations 3D and
3E are unconfirmed hypothetical intermediates that I have included in order for the reader to
better understand the role of CoSO4 in the medium and the formation of the final visible
precipitate.
Sodium bicarbonate reacts with sulphuric acid forming carbonic acid (H2CO3) and
sodium sulphate (Na2SO4); shown in equation 3A. Sodium bicarbonate also reacts with
potassium dihydrogen phosphate to form additional carbonic acid and potassium sodium
hydrogen phosphate (KNaHPO4; shown in equation 3B.
118
Equation 3A: 2NaCHO3 + H2SO4 → 2H2CO3 + Na2SO4
Equation 3B: NaCHO3 + KH2PO4 → H2CO3 + KNaHPO4
Equation 3C: H2CO3 → H+ + HCO3
- → H2O + CO2 (escapes)
Equation 3D: CoSO4 + KNaHPO4→ CoHPO4 + KNaSO4
Equation 3E: CoHPO4 + (NH4)2S → CoS + (NH4)2HPO4
Carbonic acid (formed in equations 3A and 3B) breaks down into bicarbonate (HCO3-) and a
proton (H+) in a nearly spontaneous reaction (first part of equation 3C). HCO3
- then undergoes
dehydration to form CO2 (the uncatalyzed reaction rate being rapid enough, i.e., rate constant of
14 s-1
at room temperature, for this to occur; equation 3C; Maren, 1980). The uncatalyzed
dehydration and hydration reactions reach equilibrium as the CO2 bubbles out of solution. The
equilibrium occurs at an approximate pH of 6.14 for the chemical concentrations used in
Hansson’s medium. However, since the incubation medium is open to the environment, further
loss of CO2 from the surface can occur, particularly with large surface areas and disruption of the
surface layer through stirring (Maren, 1980). The alkalinisation of the incubation medium
caused by the escape of CO2 from its surface, leads to the formation of an insoluble cobalt
precipitate which Maren (1980) reasons is a complex phosphate as shown equation 3D. Maren
(1980) further reports that in 3mm deep incubation medium, the critical pH for precipitate
formation (pH 6.8) is reached in about 12 minutes. The presence of the enzyme carbonic
anhydrase (CA) presumably speeds up the time required to reach this critical pH. Hence, an
incubation time of 8 to 10 minutes, which is not long enough to allow precipitate formation via
the uncatalyzed CO2 dehydration reaction, should be sufficient for precipitate formation
localized to tissue containing CA. The precipitate is then converted to a visible and less soluble,
119
black deposit (i.e., cobalt sulphide or CoS; equation 3E) via exposure to ammonium sulphide
((NH4)2S); a blackening agent).
The cobalt-phosphate method has been criticized over the years on the basis of specificity
and kinetics (Muther, 1972; 1977). However, these criticisms have been rebutted by several
studies such as Maren (1980), Rossen and Musser (1972) and Lönnerholm (1974; 1980). On
balance, it appears as if this technique is a valid method for histochemical identification of active
CA on the basis that a specific CA inhibitor prevents staining and the chemical kinetics of the
reactions fit the criteria for precipitate formation within the allotted incubation time.
The data in chapter 2 demonstrated that treatment with the permeant CA inhibitor,
acetazolamide, abolished the CHC-induced increase in fictive breathing. Given this, I
hypothesised that brainstem preparations taken from frogs exposed to CHC would contain a
greater amount/activity of CA than preparations taken from control frogs or frogs exposed to
CH. If this hypothesis is correct, then histochemical analysis for active CA should reveal a
greater intensity of staining, using the cobalt-phosphate method, in preparations taken from frogs
exposed to CHC compared to chronically hypoxic and control frogs.
120
4.2 MATERIALS & METHODS
Histochemical Localization of Active Carbonic Anhydrase
Active CA was detected using a modified version of the cobalt-phosphate histochemical
method (i.e., Hansson’s method; Hansson, 1967). Fifteen brainstem-spinal cords from each of
the three experimental groups (normoxic/normocapnia control, CHC and CH) were fixed for a
period of 4 hrs at 4ºC in a 3% glutaraldehyde in aqueous phosphate buffer containing sodium
phosphate dibasic and sodium phosphate monobasic monohydrate (Ricca Chemical Company,
Arlington, Texas; pH 7.0). The brainstem preparations were kept frozen at minus 80ºC and then
sectioned into 20µm slices on a cryostat (Leica CM 3040S) at minus 20ºC. The sections were
then transferred to gelatin-chrome-alum coated slides.
Histochemical localisation of CA was performed on brain slices taken from three
different locations (Taylor, et al., 2003): 1) at the level of the hypoglossal nerve root (cnXII), 2)
at the level of the vagus nerve (cnX) root and 3) at the level of the trigeminal nerve (cnV) nerve
root. These sections were processed in groups such that slices from the same regions in the
brains taken from control, CH and CHC animals were exposed to the same staining conditions at
the same time.
The brain slice-mounted slides were incubated in an approximately 3 mm deep solution
containing (in mM) 2.90 CoSO4, 17.6 KH2PO4, 156 NaHCO3, and 15.9 H2SO4 (Taylor et al.,
2003) for 8 minutes. Following this, the slides were rinsed in distilled water, immersed in 0.5%
ammonium sulphide (blackening agent) and then rinsed again. The brain slices underwent
dehydration in alcohol and were then rinsed in xylene prior to placing a cover slip over the slide
(as per Taylor et al, 2003). 100 µM ACTZ was added to the incubation solution of 4 slides in
order to determine the specificity of the test (i.e., if the staining was specific for active CA then
121
addition of a potent CA inhibitor should reduce or abolish the staining). The stained slides were
then viewed under a light microscope (Axioplan 2) at three different magnifications (25X, 100X
and 400X). Two investigators, blinded to the source of the slices (i.e., control, CHC or CH
brains), independently analyzed the stain intensity of the ventrolateral surface of each section on
a scale of 0 (no stain), 1(least intense) to 3 (most intense).
122
4.3. RESULTS
Results obtained from this histochemical method were not consistent enough to confirm
our hypothesis that the stain intensity would be greater in brains taken from chronically
hypercapnic (CHC) frogs compared to brains from control and chronically hypoxic (CH) frogs.
The stained brain sections from this histochemical study did not appear to be drastically different
across the three groups (i.e., control, chronically hypercapnic and chronically hypoxic). Quite a
bit of variability was observed when ranking the intensity of the stain at the ventrolateral portion
of the slices. Average ranks given by investigator one for the control, CHC and CH groups were
1.6 ± 0.19, 2.3 ±0.19 and 1.8 ± 0.25, respectively. Investigator two produced similar average
ranks of 1.5 ± 0.29, 2.0 ± 0.58 and 2 ± 0.58 for the control, CHC and CH, respectively, brain
slices. When rounding these averages, the stain intensity for all three groups seems to rank
equally. In addition to this discrepancy, nonspecific staining made it difficult to distinguish
staining of tissue from precipitate formation on the slide in general.
Some slices (Fig. 27), showed slightly darker staining of tissue at 25X magnification in
the CHC and CH groups. Some sections appeared uniformly darker than their counterparts,
however, no specific aggregation of black CoS precipitate was noted.
123
Figure 27: Images from a histochemical analysis of active carbonic anhydrase (CA) localisation
using a modified version of Hansson’s method (or cobalt-phosphate method). All the brain slices
in this figure were taken from a region of the medulla near the level of the vagus nerve (cnX).
Panel A represents a control for the specificity of this stain and is a slice taken from a control
(i.e., normoxic, normocapnic) leopard frog brain that had been incubated in Hansson’s medium
containing 100µM acetazolamide (ACTZ). Panels B, C and D depict brain slices taken from
control (normoxic, normocapnic), chronically hypercapnic and chronically hypoxic frogs,
respectively. The red circle in panel C represents one example of a carbonic anhydrase positive
cell body.
124
125
4.4 Discussion
4.4.1 Goals and Predictions
The results of chapter 2 indicate that the CHC-induced increase in fictive breathing seen
in leopard frogs (and also cane toads; Gheshmy et al. 2006) was abolished by acetazolamide
(ACTZ) treatment. However, since ACTZ is a cell permeant inhibitor of CA, we could not
distinguish whether or not the effects seen in chapter 2 were due (in whole or in part) to
intracellular or extracellular CA function. Experiments described in chapter 3 were devised to
further explore this issue. Exogenous CA application did not alter the CHC-induced
augmentation of fictive breathing, suggesting that extracellular CA does not account for this in
vitro CHC response (chapter 3). In order to further support the conclusions of chapters 2 and 3, I
performed a histochemical analysis for active CA. The rationale here was that the cobalt-
phosphate staining method would allow for the visualization of active CA locations within the
frog brain tissue (i.e., whether it is located in the interstitial space/extracellular, neuronal cell
bodies/intracellular, etc.). Furthermore, I hypothesized that if CA is important for the mechanism
underlying the CHC-induced increase in fictive breathing then this may be due to: 1) an increase
in the amount of CA present within CO2 chemoreceptive cells of CHC, compared to control, frog
brains or 2) enhanced activity of already present CA. Assuming that an increase in the amount of
cellular CA occurs during CHC exposure, I expect to see a greater intensity of staining in CHC
frog brain tissue compared to controls using the cobalt-phosphate method.
The superficial region of the ventral medulla has been considered to be a central pH/CO2
chemoreceptive area since the 1960s (Mitchell et al., 1963). Therefore, the intensity of staining
for carbonic anhydrase (CA) at the ventrolateral surface of transverse medullary slices in brains
126
taken from CHC- and CH-exposed frogs was compared to the staining in brains taken from
control animals.
4.4.2 Summary of Results
The visual appearance of the CoS precipitate in stained tissues from previous studies
showed deposits of black precipitate at the location of active carbonic anhydrase. However,
Taylor and colleagues (2003) had observed darker staining of cell bodies in bullfrog brain slices
rather than black precipitate aggregations per se. The sections from this study, which were
stained based on the technique described Taylor et al., (2003), also tended to show somewhat
darker stained cell bodies in the chronic hypercapnic (CHC) compared to the control frog brain
slices. Some brain sections from CHC frogs appeared uniformly darker than their control
counterparts at 25X magnification; however, no specific deposits of dark CoS precipitate were
noted at higher magnification. This begs the question of whether the diffuse stain indicates
ubiquitous cytoplasmic CA or is rather non-specific staining. Some slides contained black
deposits around the periphery of the tissue whereas others did not. Hence, it is difficult to
determine the exact nature of this stain.
Significant variability was observed when ranking the intensity of the stain at the
ventrolateral portion of each slice. Overall average rankings were slightly higher (albeit by only
a few decimal points) for the CHC group compared to CH and controls. However, due to the
variability, I cannot conclusively say that the results support our hypothesis. Indeed, due to this
variability, I came to the conclusion that this method may not be as reliable as I initially thought
to determine changes in CA activity following CHC or CH.
127
4.4.3 Cobalt-Phosphate Histochemical Method
The cobalt-phosphate method devised by Hansson (1967) initially involved floating
freeze-dried tissue sections on the surface of the incubation medium and subsequent transfer to
slides. A couple of drawbacks of this method are that carbonic anhydrase (CA) from unfixed
sections may be lost to the incubation medium (Ridderstråle, 1991), tissue disintegration may
occur upon floating in the incubation medium and the impracticality of processing a large
number of sections (i.e., serial sections). The first issue was addressed by fixing the tissue prior
to staining. Tissue fixation generally reduces biologically measurable enzyme activity, however,
improved tissue morphology for microscopy makes it advantageous (Muther, 1972; Ridderstråle,
1991). Curiously, Muther (1972) reports that stain intensity (using the cobalt-phosphate method)
remains unaltered, and may even be better, in fixed tissue. In order to address tissue
disintegration and to expedite this procedure, sections were mounted onto slides prior to
incubation. A study by Loveridge (1978) and later studies (Coates et al., 1998; Taylor et al.,
2003) have shown that it is not necessary to float tissue in order for this technique to work. In
other words, cryostat-sectioned tissue mounted on slides can be incubated in the medium (as
done in this study) and yield similar results.
Although tissue fixation and transferral to slides prior to incubation in (a modified
version of) Hansson’s medium may explain the reduced, or lack of, staining seen in some tissue
in this study, earlier research reported distinguishable staining of tissue under similar conditions
(Loveridge, 1978; Coates et al., 1998; Taylor et al., 2003). Furthermore, the dehydration process
and clearing with xylene, which may also account for the weak staining obtained in this study,
128
did not seem to interfere with staining in earlier studies (Coates et al., 1998; Taylor et al., 2003).
Incubation time is another factor that may account for slight differences between studies. Earlier
studies have used incubation times ranging from 1 to 10 minutes (Hansson, 1967; Loveridge,
1978; Ridderstråle, 1991; Coates et al., 1998; Taylor et al., 2003). Eight minutes was chosen for
this study on the basis of preliminary results showing non-specific staining at higher incubation
times. However, perhaps a slightly longer incubation time may have lead to better tissue staining.
A final point for consideration is the fact that tissue slices were not hydrated prior to incubation
in the cobalt-phosphate medium since a preset protocol (i.e., that of Taylor et al., 2003) was
followed. However, on hindsight, perhaps tissue hydration would have allowed greater tissue
penetration of the stain.
4.5 Conclusion
The results from this chapter do not provide convincing support for the hypothesis that
the chronic hypercapnia (CHC)-induced augmentation of fictive breathing in leopard frogs (Rana
pipiens) is due (at least in part) to an increase in the amount of intracellular carbonic anhydrase
(CA). Further modifications (i.e., choice of different fixatives, change in incubation times,
hydration of tissue prior to incubation in Hansson’s medium) to this method may improve the
quality of staining obtained. Given the many permutations of change that may have been
necessary to obtain adequate staining, I decided not to pursue this particularly since an alternate,
in vivo assay for CA (i.e., using radiolabelled bicarbonate, HC14
O3-; Wood and Perry, 1991) may
yield more accurate and quantitatively comparable results.
129
CHAPTER 5
GENERAL DISCUSSION
130
5.1 Goals of the Thesis
Previous studies from this laboratory have shown that chronic exposure of cane toads
(Bufo marinus), a semi-terrestrial amphibian, to hypoxia (10% O2 for 10 days) and hypercapnia
(3.5% CO2 for 9 days) had opposing effects on an index of breathing (fictive breathing)
measured in vitro using brainstem-spinal cord preparations (Gheshmy et al., 2006; 2007;
McAneney and Reid, 2007). Chronic hypoxia (CH) blunted and chronic hypercapnia (CHC)
augmented pH/CO2-sensitive fictive breathing (Gheshmy et al., 2006; 2007; McAneney and
Reid, 2007). Further investigation revealed that the CHC-induced increase in fictive breathing
was due to altered afferent feedback from the CO2-sensitive olfactory chemoreceptors and
arterial O2/CO2 chemoreceptors (Gheshmy et al., 2006; 2007). Midbrain transection experiments
revealed that changes in central descending inhibitory inputs, from the midbrain to the medulla,
was the cause of the reduction in fictive breathing following CH exposure (McAneney and Reid,
2007). Taken together, these studies reveal that there is significant interaction and integration of
the various respiratory control systems during exposure to long-term respiratory challenges such
as CHC and CH in the semi-terrestrial amphibian, Bufo marinus. However, the putative
mechanism underlying the CHC-induced increase in fictive breathing, and that of the CH-
induced reduction in fictive breathing, at the level of the central pH/CO2 chemoreceptors
themselves, remained elusive.
Given that there is considerable support in the literature (Ritucci et al., 1997; 1998; Wang
et al., 2002; Putnam et al., 2004) for the notion that changes in intracellular pH are the stimulus
for central pH/CO2 chemoreceptor activation and that such pH changes ultimately depend on the
hydration of CO2 to HCO3- and H
+, I decided to examine the role of carbonic anhydrase (CA; the
enzyme that catalyzes the reversible hydration/dehydration of CO2) in the modulation of
131
pH/CO2-sensitive fictive breathing during exposure to CHC and CH. Previous studies have
shown that CA may be involved in both peripheral (Gray, 1971; Nurse, 1990; Iturriaga et al.,
1991; Iturriaga et al., 1993) and central (Erlichman et al., 1994; Taylor et al., 2003) pH/CO2
chemoreception.
In order to determine whether or not CA may be involved in the CHC- and CH-induced
changes in pH/CO2 chemosensitivity, giving rise to changes in fictive breathing, I first performed
a series of experiments (Chapter 2) in which the potent cell-permeant CA inhibitor,
acetazolamide (ACTZ), was applied to the brainstem-spinal cord preparations. Given the
importance of CA in the CHC-induced increase in fictive breathing (see Chapter 2), I then
attempted to determine whether the CA in question was intracellular or extracellular (Chapter 3).
Based on previous studies (Ritucci et al., 1997; 1998; Wang et al., 2002; Putnam et al, 2004), I
hypothesised that the effects observed in Chapter 2 (ACTZ abolished the CHC-induced increase
in fictive breathing) were due to effects on intracellular CA. To test this hypothesis, I added
exogenous CA to the medium bathing in vitro brainstem spinal cord preparations from leopard
frogs. Finally, in Chapter 4, I performed a histochemical analysis of the location of CA in brain
tissue from frogs exposed to control conditions, CHC and CH. The cobalt-phosphate technique
used in these experiments was designed to indicate both the location and amount (perceived via
stain intensity) of CA within these tissues.
5.2 Critique of In Vitro Brainstem-Spinal Cord Preparation
The previous studies from this laboratory, that served as the impetus for this thesis, as
well as the current studies enclosed in this thesis, all made use of the in vitro brainstem-spinal
cord preparation. This preparation is devoid of any peripheral input that is otherwise present in
132
the intact animal. For this reason, the in vitro brainstem-spinal cord preparation serves as a
means to study the central control of breathing. Central respiratory rhythm and pattern
generation, central pH/CO2 chemoreceptor function and midbrain influences on breathing are all
aspects that can be analyzed using the in vitro brainstem-spinal cord preparation.
However, one must not ignore the potential drawbacks inherent in this isolated in vitro
preparation. The lack of afferent input previously mentioned as an advantage also serves as a
disadvantage. In vivo, feedback from both peripheral and central control systems are integrated,
resulting in an overall level of respiratory drive. Respiratory stressors (i.e., hypoxia and
hypercapnia) that cause a certain response in vivo, due to stimulation of various different control
systems, may not result in as profound a response when administered in vitro to a reduced
preparation (Reid, 2006).
One such example is the hypercapnic ventilatory response seen in amphibian in vitro
brainstem-spinal cord preparations. In preparations taken from both cane toads and leopard frogs,
the hypercapnic response, or in other words, the response of fictive breathing to acute changes in
pH of the superfusate, were not as pronounced as the changes in breathing observed in vivo in
response to similar changes in arterial pH. Interaction between peripheral and central
chemoreceptors gives rise to the overall respiratory response to hypercapnia (Smatresk, 1990;
Smatresk and Smits, 1991; Kinkead and Milsom, 1994; Reid, 2006). However, in the isolated, in
vitro brainstem-spinal cord preparation, acidification of the superfusate stimulates central
pH/CO2 chemoreceptors, the only subset of chemoreceptors that is present.
According to Branco (1992), central pH/CO2 chemosensitivity is responsible for
approximately 80% of the in vivo hypercapnic ventilatory response. Yet, in vitro, the
contribution of central pH/CO2 chemosensitivity to the rise in fictive breathing following
133
acidification of the superfusate was not large enough to represent 80% of the changes in
breathing observed in vivo. Nevertheless, the use of in vitro brainstem-spinal cord preparations to
study the effects of CHC and CH exposure on central pH/CO2 chemoreceptor function is valid
since spontaneous motor output in vitro has been shown to correspond to breathing in the intact
animal (Sakakibara, 1984).
One possible explanation for the discrepancy between in vivo and in vitro central pH/CO2
chemosensitivity arises when considering the findings of a study by Winmill et al. (2005).
Winmill and colleagues (2005) suggest that there may be a means for central O2 detection in the
bullfrog based upon the observed reversible cessation of fictive breathing following acute
hypoxia in bullfrog in vitro isolated brainstem-spinal cord preparations. If this were in fact true,
then it would be interesting to see whether O2 (hypoxia) plays a modulatory role in central CO2
chemosensitivity (a similar relationship to that seen in the peripheral system). In other words,
perhaps central CO2 sensitivity may be dependent on the level of PO2 in the cerebrospinal fluid
and vice versa such that high levels of PO2 result in lower PCO2 sensitivity. This could possibly
explain the low levels of fictive breathing/CO2-sensitivity seen in the control preparations since
O2 levels in the in vitro superfusate were maintained high (approximately 650 mmHg).
5.3 CO2-Sensitive Respiratory Control Systems
CO2 is a stronger respiratory stimulus than O2 for terrestrial air breathers. In fact,
Smatresk et al. (1991) postulated that the evolution of central CO2 chemoreceptors coincides
with an increase in sensitivity to hypercapnia. There are numerous CO2-sensitive receptors that
134
can affect breathing. These receptors will be stimulated by CO2 during both acute and chronic
hypercapnia.
During hypercapnia in vivo, high PCO2 levels sensed by the nasal olfactory CO2
chemoreceptors cause an inhibition (whether it be via inhibitory neurotransmitter release or
reduced stimulatory signals) of breathing (Sakakibara, 1978; Coates and Ballam, 1990; Coates,
2001). Pulmonary stretch receptors (PSR), located in the lung walls, are also CO2-sensitive,
reducing their firing rate as CO2 levels increase (Milsom and Jones, 1977; Kuhlman and Fedde,
1979). PSR input has complex effects on amphibian breathing. Phasic PSR feedback stimulates
breathing and promotes the clustering of breaths (lung inflation) while tonic PSR feedback
during periods of breath-holding stimulates small, continuous deflation breaths. High levels of
inspired PCO2 result in elevated arterial PCO2 and subsequent stimulation of arterial
chemoreceptors (i.e., peripheral chemoreceptors located in the carotid labyrinth and
pulmocutaneous arch). The stimulatory signals from these chemoreceptors are carried by the
glossopharyngeal nerve to central respiratory centres.
CO2 in arterial blood crosses the blood brain barrier and results in high PCO2 in the
cerebrospinal fluid. The CO2 in the cerebrospinal fluid traverses the central respiratory pH/CO2
chemoreceptor cell membranes. Once inside the cell, CO2 hydration, aided by the enzyme
carbonic anhydrase (CA), produces rapid intracellular acidification and subsequent activation of
ion exchangers (i.e., Na+/H
+; HCO3
-/Cl
-; Na
+/Ca
2+). Ultimately, Ca
2+ influx into these
chemoreceptor cells results in neurotransmitter release which sends signals to central/medullary
respiratory centres which then trigger an increase in breathing.
135
5.4 Role of Carbonic Anhydrase in CO2 Chemoreception Following Chronic Hypercapnia
The results of the experiments in Chapters 2 through 4 indicate a role for carbonic
anhydrase (CA), at the level of central pH/CO2 chemoreceptors, in the mechanism underlying the
CHC-induced increase in fictive breathing. In Chapter 2, the application of a potent, cell-
permeant inhibitor of CA (acetazolamide; ACTZ) at concentrations that were deemed sufficient
enough to inhibit most, but not all CA, caused a blunting of fictive breathing in brainstem-spinal
cord preparations taken from frogs exposed to control and chronically hypercapnic (CHC)
conditions. However, no such effect was observed in preparations taken from frogs exposed to
chronic hypoxia (CH). The effects seen in the control group suggest a role for CA in central
pH/CO2 chemosensitivity while the effects seen in the CHC group (i.e., the CHC-induced
increase in fictive breathing was abolished following the addition of ACTZ) indicate that
alterations in CA activity or amount appear to be responsible for the effects following CHC.
Following this series of ACTZ experiments, the experiments in Chapter 3 were
performed in order to determine whether the effects of CA revealed in Chapter 2 were the result
of intracellular or extracellular CA. Chapter 3 showed that addition of exogenous CA to the
superfusate bathing the brainstem-spinal cord preparations did not significantly alter fictive
breathing other than a blunting of fictive breathing frequency in control preparations. If
extracellular CA was responsible for the effects seen in Chapter 2 then I predicted that there
would be an increase in fictive breathing following addition of CA in the control group. In other
words, the effects of CHC on fictive breathing would be mimicked by the addition of CA.
However, given that this did not occur, the results from Chapter 3 support my hypothesis that
intracellular CA, by catalyzing the hydration of CO2 within the chemoreceptor cells and
subsequent changes in intracellular pH, underlies central pH/CO2 chemosensitivity.
136
Chapter 4 described a histochemical analysis of the location of active CA in brain slices,
predominately from the medulla, taken from frogs exposed to control conditions as well as CH
and CHC. I expected to see darker staining of neuronal cell bodies/somas (indicating the
presence of intracellular CA) and generally darker stain intensity (indicative of increased amount
of intracellular CA) in brains taken from animals exposed to CHC. However, although some
slices suggested that this occurred, the overall inconsistency of the stained tissues did not allow
for a definitive conclusion.
5.5 Different Effects of Chronic Hypoxia in Terrestrial Versus Aquatic Amphibians
McAneney and Reid (2007) demonstrated that exposure to chronic hypoxia caused a
blunting of fictive breathing in the cane toad (Bufo marinus). In contrast, the results from this
thesis show that an identical protocol of CH exposure of leopard frogs (Rana pipiens) did not
alter fictive breathing. One possible explanation for this difference is the differing conditions
under which these two species encounter hypoxia in their natural environments. Unlike the
terrestrial anuran species that experience bouts of hypoxia while overwintering underground,
leopard frogs, a semi-aquatic species that overwinters under ice-covered bodies of water, may be
more tolerant of hypoxia (Pinder et al., 1992; Hermes-Lima and Zenteno-Savín, 2002).
Unlike some species of freshwater turtles (such as the painted turtle; Chrysemys picta)
that are anoxia-tolerant, leopard frogs can only tolerate anoxia for a period of a few days at low
temperatures (30 hrs at 5ºC; Hermes-Lima and Storey, 1996) and only a few hours (4-5 hrs;
Knickerbocker and Lutz, 2001) at room temperature. This short-term “tolerance” of anoxia
situates anuran amphibians in the intermediary zone on the spectrum consisting of anoxia-
137
sensitive mammalian brains on one end and anoxia-tolerant species such as fresh water turtles
(Hutchison and Dady, 1964; Bickler and Buck, 2007) on the other. Contrary to anoxia-tolerant
turtles that can maintain brain ATP levels for extended periods and mammalian brains that
rapidly lose ATP (i.e., within minutes), frog brains experience a slow reduction in brain ATP and
subsequent brain death during anoxic conditions (Bickler and Buck, 2007). However, leopard
frogs are quite hypoxia-tolerant and can maintain brain ATP levels during hypoxia (water PO2 of
30-60 mmHg which corresponds to approximately 4-8% O2) via hypometabolism in cold
temperatures for up to 16 weeks (Knickerbocker and Lutz, 2001). It is possible that the level of
hypoxic exposure in the current study was not sufficient to induce changes and that exposure to a
more severe level of hypoxia, such as those encountered in their natural environments, may
produce a significant modulation of fictive breathing. Alternatively, the temperature at which
hypoxia was experienced could also have affected ventilatory responses since most studies show
that hypoxia tolerance is prolonged at lower temperatures that promote greater metabolic savings
(Hermes-Lima and Storey, 1996; Tattersall and Boutilier, 1997; Boutilier, 2001). In all studies
included in this thesis, chronic hypoxic acclimatisation was performed at room temperature.
In addition, the current studies exposed leopard frogs to terrestrial normobaric hypoxia.
Since aquatic species, to a great extent, rely on cutaneous gas exchange (Burggren and West,
1982), perhaps exposure to aquatic hypoxia and hypercapnia may have produced greater changes
in the fictive ventilatory responses. During periods of inactivity, cutaneous gas exchange can
account for approximately 20% of O2 exchange and the majority of CO2 exchange in (leopard
frogs in) well-aerated waters (Pinder and Burggren, 1986). One would assume that these values
would be greater in the poorly-oxygenated waters found during overwintering conditions. During
terrestrial CH, the minimal contribution of cutaneous gas exchange would result in greater
138
reliance on the lungs for O2 and CO2 exchange. Perhaps, a build-up of arterial CO2 (due to the
loss of one of the major routes for its removal) may trigger CO2 chemoreceptors (both central
and peripheral) which in turn stimulate brainstem respiratory centres. This would offset the
hypoxia-induced descending inhibitory input to brainstem respiratory centres, resulting in no
overall change in breathing (as seen in the studies presented in this thesis).
It is also possible that the central inhibitory influences that reduce fictive breathing
during CH in toads (McAneney and Reid, 2007) are absent in frogs. However, midbrain control
of episodic breathing is the same in the two groups of animals (Kinkead et al., 1997; Gargaglioni
and Branco, 2000; 2001; 2003; 2004; Reid et al. 2000a; Gargaglioni et al., 2002; McAneney and
Reid, 2007). On the other hand, perhaps control over central CO2 chemoreception is different
between the two groups, given the aquatic (frog) versus terrestrial (toad) nature of these animals.
An early study examining the viability of Rana pipiens and Bufo terrestris (a terrestrial anuran
species) during submergence underwater at varying temperatures showed that R. pipiens was
more adapted to prolonged submergence (Hutchison and Dady, 1964). Hence, this adaptation to
submergence may coincide with greater adaptation to hypoxia.
5.6 Perspectives
Although this thesis project satisfactorily attained the initial goal of determining the role
of CA in the chronic hypercapnia-induced increase in central respiratory-related pH/CO2
chemosensitivity (manifested as an increase in fictive breathing), more questions remain to be
answered with regards to the intracellular versus extracellular location of CA. In order to further
distinguish intracellular from extracellular CA, future investigations could make use of relatively
139
cell-impermeant CA inhibitors such as benzolamide. Assuming that intracellular CA is involved
in central respiratory-related pH/CO2 chemosensitivity, addition of benzolamide to the
superfusate of isolated, in vitro brainstem-spinal cord preparations should not alter fictive
breathing. Further studies could look into the effects of phosphatidylinositol-specific
phospholipase C (PIP-C), an enzyme that cleaves the glycosyl-phosphatidyl-inositol that anchors
extracellular CA isozymes IV to the plasma membrane. Again, assuming that intracellular CA is
responsible for central respiratory-related pH/CO2 chemosensitivity, I would not expect fictive
breathing to be altered following addition of PIP-C to the superfusate of isolated, in vitro
brainstem-spinal cord preparations. As an alternative to the in vitro histochemical analysis of CA
location, in vivo assays for activity of CA (using radioactively labelled HCO3-) may produce
more reliable and conclusive results. Other isozymes have different kinetics and properties so
other suitable inhibitors may be used to verify the role of other forms of CA within the brain
(i.e., CAIII which is relatively insensitive to acetazolamide).
Having shown that CA is important not only in central pH/CO2 chemoreception but that it
also underlies the CHC-induced augmentation in fictive breathing in leopard frogs, it would be
worthwhile to explore the potential stimuli for this CA up-regulation. Central respiratory centres
process various stimulatory and inhibitory signals (originating from both afferent and central
sources) and modify breathing accordingly via efferent input to respiratory muscles (i.e.,
primarily via vagal, trigeminal or hypoglossal nerves which innerve the primary respiratory
muscles; McLean and Remmers, 1997). Given this, previous studies from this laboratory have
already explored the effects of eliminating olfactory chemoreceptor input, peripheral/arterial
chemoreceptor input and descending input from higher brain centres upon the CHC-induced
augmentation in fictive breathing in a semi-terrestrial amphibian (Bufo marinus, the cane toad).
140
The effects of pulmonary stretch receptor feedback on the CHC-induced increase in fictive
breathing also remains unknown and needs to be investigated.
141
REFERENCES
Bagust, J., Kelly, M.E. and Kerkut, G.A. (1985). An isolated mammalian brainstem-
spinal cord preparation suitable for the investigation of descending control of motor activity.
Brain Res. 327 (1-2), 370-374.
Bickler, P. E. (1992). Cerebral anoxia tolerance in turtles: regulation of intracellular
calcium and pH. Am. J. Physiol. Regul. Integr. Comp. Physiol. 263, R1298-R1302.
Bickler, P.E. and Buck, L.T. (2007). Hypoxia tolerance in reptiles, amphibians, and
fishes: life with variable oxygen availability. Annu. Rev. Physiol. 69, 145-170.
Black, A.M.S., McCloskey, D.I. and Torrance, R.W. (1966). The responses of peripheral
chemoreceptors to sudden changes of hypercapnic and hypoxic stimuli. J. Physiol. (London) 185,
67-68P.
Boggs, D.F., Kilgore, Jr., D.L. and Birchard, G.F. (1984). Respiratory physiology of
burrowing mammals and birds. Comp. Biochem. Physiol. 77A, 1-7.
Bongianni, F., Mutolo, D., Nardone, F. and Pantaleo, T. (2006). GABAergic and
glycinergic inhibitory mechanisms in the lamprey respiratory control. Brain Res. 1090 (1), 134-
145.
142
Boutilier, R.G., Randall, D.J., Shelton, G. and Toews, D.P. (1979). Acid-base
relationships in the blood of the toad, Bufo marinus. J. Exp. Biol. 82, 331-344.
Boutilier, R.G. and Heisler, N. (1988). Acid-base regulation and blood gases in the
anuran amphibian, Bufo marinus, during environmental hypercapnia. J. Exp. Biol. 134, 79-98.
Boutilier, R.G. (2001). Mechanisms of metabolic defense against hypoxia in hibernating
frogs. Respir. Physiol. 128 (3), 365-377.
Branco, L., Glass, M. and Hoffmann, A. (1992). Central chemoreceptor drive to
breathing in unanesthetized toads, Bufo paracnemis. Respir. Physiol. 87 (2), 195–204.
Brown, D. (1980). Carbonic anhydrase localization in mounted cryostat sections. Stain
Technol. 55 (2), 115-118.
Brown, D. and Waneck, G.L. (1992). Glycosyl-phosphatidylinositol-anchored membrane
proteins. J. Am. Soc. Nephrol., 3 (4), 895-906.
Buckler, K.J. and Vaughan-Jones, R.D. (1993). Effects of acidic stimuli on intracellular
calcium in isolated type I cells of the neonatal rat carotid body. Pflugers Arch. 425 (1-2), 22-27.
143
Bundy, H.F. and Cheng, B. (1976). Amphibian carbonic anhydrase: purification and
partial characterization of the enzyme from erythrocytes of Rana catesbeiana. Comp. Biochem.
Physiol. 55B (2), 265-271.
Burggren, W.W. and West, N.H. (1982). Changing respiratory importance of gills, lungs
and skin during metamorphosis in the bullfrog Rana catesbeiana. Respir. Physiol. 47 (2), 151-
164.
Burggren, W. and Doyle, M. (1986). Ontogeny of regulation of gill and lung ventilation
in the bullfrog, Rana catesbeiana. Respir. Physiol. 66 (3), 279-291.
Burggren, W. W. and Infantino, R. L. JR. (1994). The respiratory transition from water to
air breathing during amphibian metamorphosis. Am. Zool. 34 (2), 238-246.
Cammer, W. (1991). Carbonic anhydrase in myelin and glial cells in the mammalian
central nervous system. In: Dodgson, S.J., Tashian, R.E., Gros, G. and Carter, N. D. (Eds), The
Carbonic Anhydrases: Cellular Physiology and Molecular Genetics. Plenum Press, New York,
pp. 325-332.
Chegwidden, W.R. (1991). Purification of the carbonic anhydrases. In: Dodgson, S.J.,
Tashian, R.E., Gros, G. and Carter, N. D. (Eds), The Carbonic Anhydrases: Cellular Physiology
and Molecular Genetics. Plenum Press, New York, pp. 101-118.
144
Chegwidden, W.R. and Carter, N. (2000). Introduction to the carbonic anhydrases. In:
Chegwidden, W.R., Edwards, Y. and Carter, N. (Eds), The Carbonic Anhydrases: New Horizons.
Birkhäuser Verlag, Basel, pp. 14-28.
Coates, E.L. and Ballam, G.O. (1990). Olfactory receptor response to CO2 in bullfrogs.
Am. J. Physiol. 27, R1207–R1212.
Coates, E.L., Li, A.H. and Nattie, E.E. (1991). Acetazolamide on the ventral medulla of
the cat increases phrenic output and delays the ventilatory response to CO2. J. Physiol. 441, 433-
451.
Coates, E.L., Li, A., and Nattie, E.E. (1993). Widespread sites of brain stem ventilatory
chemoreceptors. J. Appl. Physiol. 75 (1), 5-14.
Coates, E.L., Wells, C.M.Q. and Smith, R.P. (1998). Identification of carbonic anhydrase
activity in bullfrog olfactory receptor neurons: histochemical localization and role in CO2
chemoreception. J. Comp.Physiol. 182A, 163-174.
Coates, E.L. (2001). Olfactory CO2 chemoreceptors. Respir. Physiol. 129,
219–229.
De la Torre, J.C. and Rowed, D.W. (1974). DMSO: a new respiratory stimulant? J. Clin.
Pharmacol. 14 (7), 345-353.
145
Delvolvé, I., Branchereau, P., Dubuc, R. and Cabelguen, J.M. (1999). Fictive rhythmic
motor patterns induced by NMDA in an in vitro brain stem-spinal cord preparation from an adult
urodele. J. Neurophysiol. 82 (2), 1074-1077.
Dodgson, S. (1991). The Carbonic Anydrases: Overview of their importance in cellular
physiology and in molecular genetics. In: Dodgson, S.J., Tashian, R.E., Gros, G. and Carter, N.
D. (Eds), The Carbonic Anhydrases: Cellular Physiology and Molecular Genetics. Plenum Press,
New York, pp. 3-14.
Douse, M.A. and Mitchell, G.S. (1990). Episodic respiratory related discharge in turtle
cranial motorneurons: in vivo and in vitro studies. Brain Res. 536 (1-2), 297-300.
Duffin, J. (2005). Role of acid-base balance in the chemoreflex control of breathing. J.
Appl. Physiol. 99, 2255-2265.
Emery, A. R., Berst, A. H. and Kodaira, K. (1972). Under-ice observations of wintering
sites of leopard frogs. Copeia 1972 (1), 123-126.
Eriksson, E. and Liljas, A. (1991). X-ray crystallographic studies of carbonic anydrase
isozymes I, II and III. In: Dodgson, S.J., Tashian, R.E., Gros, G. and Carter, N. D. (Eds), The
Carbonic Anhydrases: Cellular Physiology and Molecular Genetics. Plenum Press, New York,
pp. 33-48.
146
Erlichman, J.S., Coates, E.L. and Leiter, J.C. (1994). Carbonic anhydrase and CO2
chemoreception in the pulmonate snail Helix aspersa. Respir. Physiol. 98, 27-41.
Feldman, J.L. and Del Negro, C.A. (2006). Looking for inspiration: new perspectives on
respiratory rhythm. Nat. Rev. Neurosci. 7 (3), 232-242.
Fernley, R.T. (1990). Secreted carbonic anhydrases. In: Botre,F., Gros, G. and Storey,
B.T. (Eds.), Carbonic Anhydrase: From Biochemistry and Genetics to Physiology and Clinical
Medicine. VCH Publishers, New York, pp 178-185.
Forster, R.E. (2000). Remarks on the discovery of carbonic anhydrase. In: Chegwidden,
W.R., Carter, N.D. and Edwards, Y.H. (Eds.), The Carbonic Anhydrases: New Horizons.
Birkhäuser Verlag, Basel, Switzerland, pp. 1-11.
Fournier, S., Allard, M., Roussin, S. and Kinkead, R. (2007). Developmental changes in
central O2 chemoreflex in Rana catesbeiana: the role of noradrenergic modulation. J. Exp. Biol.
210, 3015-3026.
Galante, R.J., Kubin, L., Fishman, A. P. and Pack, A. I. (1996). Role of chloride-
mediated inhibition in respiratory rhythmogenesis in an in vitro brainstem of tadpole, Rana
catesbeiana. J Physiol. 492 (2), 545-558.
147
Gans, C., De Jongh, H.J. and Farber, J. (1969). Bullfrogs (Rana catesbeiana) ventilation:
how does the frog breathe? Science, 163 (872), 1223-1225.
Gans, C. and Pyles, R. (1983). Narial closure in toads; which muscles? Respir. Physiol.
53 (2), 215-223.
Gargaglioni, L.H. and Branco, L.G. (2000). Role of nucleus isthmi in the ventilatory
response to hypoxia of Bufo paracnemis. Respir. Physiol. 119 (1), 31-39.
Gargaglioni, L.H. and Branco, L.G. (2001). Effect of nitric oxide in the nucleus isthmi on
the hypoxic and hypercarbic drive breathing of toads. Am. J. Physiol. Regul. Integr. Comp.
Physiol. 281 (1), R338-345.
Gargaglioni, L.H., Coimbra, N.C. and Branco, L.G. (2002). Chemical lesions of the
nucleus isthmi increase the hypoxic and hypercarbic drive to breathing of toads. Respir. Physiol.
Neurobiol. 132 (3), 289-299.
Gargaglioni, L.H. and Branco, L.G. (2003). The role of glutamate in the nucleus isthmi on
the hypoxia- and hypercarbia-induced hyperventilation of toads. Respir. Physiol. Neurobiol. 135
(1), 47-58.
Gargaglioni, L.H. and Branco, L.G. (2004). Nucleus isthmi and control of breathing in
amphibians. Respir. Physiol. Neurobiol. 143 (2-3), 177-186.
148
Gargaglioni, L.H. and Milson, W.K. (2007). Control of breathing in anuran amphibians.
Comp. Biochem. Physiol. 147A (3), 665-684.
Gdovin, M.J., Torgerson, C.S. and Remmers, J.E. (1999). The fictively breathing tadpole
brainstem preparation as a model for the development of respiratory pattern generation and
central chemoreception. Comp. Biochem. Physiol. 124A (3), 275-286.
Gheshmy, A., Vukelich, R., Noronha, A. and Reid, S.G. (2006). Chronic hypercapnia
modulates respiratory-related central pH/CO2 chemoreception in an amphibian. J. Exp. Biol. 209,
1135–1146.
Gheshmy, A., Anari, A., Besada, D. and Reid, S.G. (2007). Afferent input modulates
the chronic hypercapnia-induced increase in respiratory-related central pH/CO2 chemosensitivity
in the cane toad (Bufo marinus). J. Exp. Biol. 210, 227–237.
Giacobini, E. (1961). Localization of carbonic anhydrase in the nervous system. Science.
134, 1524-1525.
González, C., Almaraz, L., Obeso, A. and Rigual, R. (1992). Oxygen and acid
chemoreception in the carotid body chemoreceptors. Trends Neurosci. 15 (4), 146-153.
Gray, B.A. (1971). On the speed of the carotid chemoreceptor response in relation to the
kinetics of CO2 hydration. Respir. Physiol. 11 (2), 235-246.
149
Hansson, H.P.J. (1967). Histochemical demonstration of carbonic anhydrase activity.
Histochemie. 11 (2), 112-128.
Hanson, M.A., Nye, P.C. and Torrance, R.W. (1981). The location of carbonic anhydrase
in relation to the blood–brain barrier at the medullary chemoreceptors of the cat. J. Physiol.
London 320, 113–125.
Henry, R.P. and Swenson, E.R. (2000). The distribution and physiological significance of
carbonic anhydrase in vertebrate gas exchange organs. Respir. Physiol. 121, 1-12.
Hermes-Lima, M., and Storey, K.B. (1996). Relationship between anoxia exposure and
antioxidant status in the frog Rana pipiens. Am. J. Physiol. 271 (4-2), R918-925.
Hermes-Lima, M. and Storey, K.B. (1998). Role of antioxidant defenses in the tolerance
of severe dehydration by anurans. The case of the leopard frog Rana pipiens. Mol. Cell Biochem.
189 (1-2), 79-89.
Hermes-Lima, M. and Zenteno-Savín, T. (2002). Animal response to drastic changes in
oxygen availability and physiological oxidative stress. Comp. Biochem. Physiol. 133C (4), 537-
556.
Hilvo, M., Supuran, C.T. and Parkkila, S. (2007). Characterization and inhibition of the
recently discovered carbonic anydrase isoforms CA XIII, XIV and XV. Curr. Top. Med. Chem. 7
(9), 893-899.
150
Hitzig, B.M. and Jackson, D.C. (1978). Central chemical control of ventilation in the
unanesthetized turtle. Am. J. Physiol. 235 (5), R257-264.
Hoffmann, A. and DeSousa, M.B.C. (1982). Cardiovascular reflexes in conscious toads.
J. Auton. Nerv. Syst. 5, 345-355.
Huang, W., Smith, S.E. and Chesler, M. (1995). Addition of carbonic anhydrase
augments extracellular pH buffering in rat cerebral cortex. J. Neurophysiol. 74 (4), 1806-1809.
Hutchison, H. and Dady, M.J. (1964). The viability of Rana pipiens and Bufo terrestris
submerged at different temperatures. Herpetologica. 20 (3), 149-162.
Iturriaga, R., Lahiri, S. and Mokashi, A. (1991). Carbonic anhydrase and chemoreception
in the cat carotid body. Am J Physiol. 261 (4-1), C565-573.
Iturriaga, R., Mokashi, A. and Lahiri, S. (1993). Dynamics of carotid body responses in
vitro in the presence of CO2-HCO3-: role of carbonic anhydrase. J. Appl. Physiol. 75 (4), 1587-
1594.
Jiang, C., Rojas, A., Wang, R. and Wang X. (2005). CO2 central chemosensitivity: why
are there so many sensing molecules? Respir. Physiol. Neurobiol. 145 (2-3), 115-126.
151
Johnson, S.M. and Mitchell, G.S. (1998). N-methyl-D-aspartate-mediated bulbospinal
respiratory drive is pH/PCO2-insensitive in turtle brainstem-spinal cord. Respir Physiol. 113 (3),
201-212.
Johnson, S.M. and Mitchell, G.S. (2000). Activity-dependent plasticity of descending
synaptic inputs to spinal motorneurons in an in vitro turtle brainstem-spinal cord preparation. J.
Neurosci. 20 (9), 3487-3495.
Kaur, IP., Smitha, R., Aggarwal, D. and Kapil, M. (2002). Acetazolamide: future
perspective in topical glaucoma therapeutics. Int. J. Pharm. 248 (1-2), 1-14.
Keifer, J. and Kalil, K. (1989). Modulation of spinal reflexes by pyramidal tract
stimulation in an in vitro brainstem-spinal cord preparation from the hamster. Exp. Brain Res. 78
(3), 633-640.
Kinkead, R. and Milsom, W.K. (1994). Chemoreceptors and control of episodic
breathing in the bullfrog (Rana catesbeiana). Respir. Physiol. 95 (1), 81–98.
Kinkead, R., Filmyer, W.G., Mitchell, G.S. and Milsom, W.K. (1994). Vagal input
enhances responsiveness of respiratory discharge to central changes in pH/CO2 in bullfrogs. J.
Appl. Physiol. 77 (4), 2048-2051.
Kinkead, R. and Milsom,W.K. (1996). CO2-sensitive olfactory and pulmonary receptor
152
modulation of episodic breathing in bullfrogs. Am. J. Physiol. 270, R134–144.
Kinkead, R. and Milsom, W.K. (1997). Role of pulmonary stretch receptor feedback in
control of episodic breathing in the bullfrog. Am. J. Physiol. 272 (2-2), R497-508.
Kinkead, R., Harris, M.B. and Milsom, W.K. (1997). The role of the nucleus isthmi in
respiratory pattern formation in bullfrogs. J. Exp. Biol. 200 (12), 1781-1793.
Knickerbocker, D.L. and Lutz, P.L. (2001). Slow ATP loss and the defense of ion
homeostasis in the anoxic frog brain. J. Exp. Biol. 204 (20), 3547-3551.
Kogo, N., Perry, S.F. and Remmers, J.E. (1994). Neural organization of the ventilatory
activity in the frog, Rana catesbeiana. I. J. Neurobiol. 25 (9), 1067-1079.
Kogo, N., Perry, S.F. and Remmers, J.E. (1997). Laryngeal motor control in frogs: role of
vagal and laryngeal feedback. J. Neurobiol. 33 (3), 213-222.
Kuhlman, W.D. and Fedde, M.R. (1979). Intrapulmonary receptors in the bullfrog:
Sensitivity to CO2. J. Comp. Physiol. 132, 69–75.
Lahiri, S. (1991). Carbonic anhydrase and chemoreception in carotid and aortic bodies. .
In: Dodgson, S.J., Tashian, R.E., Gros, G. and Carter, N. D. (Eds), The Carbonic Anhydrases:
Cellular Physiology and Molecular Genetics. Plenum Press, New York, pp. 341-344.
153
Lahiri, S. and Forster, R.E. II. (2003). CO2/H+ sensing: peripheral and central
chemoreception. Int. J. Biochem. Cell Biol. 35, 1413–1435.
Leusen, I. (1972). Regulation of cerebrospinal fluid composition with reference to
breathing. Physiol. Rev. 52 (1), 1-56.
Liljas, A., Håkansson, K., Jonsson, B.H. and Xue, Y. (1994). Inhibition and catalysis of
carbonic anhydrase. Recent crystallographic analyses. Eur. J. Biochem. 219 (1-2), 1-10.
Lindskog, S., Engberg, P., Forsman, C., Ibrahim, S.A., Jonsson, B.H., Simonsson, I. And
Tibell, L. (1984). Kinetics and mechanism of carbonic anhydrase isoenzymes. Ann. N. Y. Acad.
Sci. 429, 61-75.
Lindskog, S. and Silverman, D.N. (2000). The catalytic mechanism of mammalian
carbonic anhydrases. In: Chegwidden, W.R., Carter, N.D. and Edwards, Y.H. (Eds.), The
Carbonic Anhydrases: New Horizons. Birkhäuser Verlag, Basel, Switzerland, pp. 175-195.
Lönnerholm, G. (1974). Carbonic anhydrase histochemistry, a critical study of Hansson’s
cobalt-phosphate method. Acta Physiol. Scand. Suppl. 418, 1-43.
Lönnerholm, G. and Ridderstråle, Y. (1974). Distribution of carbonic anhydrase in the
frog nephron. Acta Physiol. Scand. 90 (4), 764-778.
154
Lönnerholm, G. (1980). Carbonic anhydrase in rat liver and rabbit skeletal muscle:
further evidence for the specificity of the histochemical cobalt-phosphate method of Hansson. J.
Histochem. Cytochem. 28 (5), 427-433.
Loveridge, N. (1978). A quantitative cytochemical method for measuring carbonic
anhydrase activity. Histochem J. 10 (3), 361-372.
Maren, T.H. (1967). Carbonic anhydrase: chemistry, physiology and inhibition. Physiol.
Rev. 47 (4), 595-781.
Maren, T.H. (1980). Kinetics, equilibrium and inhibition in the Hansson histochemical
procedure for carbonic anhydrase: a validation of the method. Histochem. J. 12 (2), 183-190.
McAneney, J., Gheshmy, A., Uthayalingam, S., and Reid, S.G. (2006). Chronic hypoxia
modulates NMDA-mediated regulation of the hypoxic ventilatory response in an amphibian,
Bufo marinus. Respir. Physiol. Neurobiol. 153, 23-38.
McAneney, J. and Reid, S.G. (2007). Chronic hypoxia attenuates central respiratory-related
pH/CO2 chemosensitivity in the cane toad. Respir. Physiol. Neurobiol. 156 (3), 266–275.
McClellan, A.D. (1984). Descending control and sensory gating of “fictive” swimming
and turning responses elicited in an in vitro preparation of the lamprey brainstem/spinal cord.
Brain Res. 302 (1), 151-162.
155
McLean, H.A., Kimura, N., Kogo, N., Perry, S.F. and Remmers, J.E. (1995a). Fictive
respiratory rhythm in the isolated brainstem of frogs. J. Comp. Physiol. 176A (5), 703-713.
McLean, H.A., Perry, S.F. and Remmers, J.E. (1995b). Two regions in the isolated
brainstem of the frog that modulate respiratory-related activity. J. Comp. Physiol. 177 (2), 135-
144.
Mclean, H.A. and Remmers, J.E. (1997). Characterization of respiratory-related neurons
in the isolated brainstem of the frog. J. Comp. Physiol. 181A (2), 153-159.
Meldrum, N.U. and Roughton, F.J. (1933). Carbonic anhydrase. Its preparation and
properties. J. Physiol. 80 (2), 113-142.
Milsom, W.K. and Jones, D.R. (1977). Carbon dioxide sensitivity of pulmonary
receptors in the frog. Experientia 33, 1167–1168.
Milsom, W.K. (1991). Intermittent breathing in vertebrates. Annu. Rev. Physiol.
53, 87–105.
Milsom, W.K. (2002). Phylogeny of CO2/H+
chemoreception in vertebrates. Respir.
Physiol. Neurobiol. 131 (1-2), 29-41.
Milsom, W.K. and Burleson, M.L. (2007). Peripheral arterial chemoreceptors and the
evolution of the carotid body. Respir. Phsyiol. Neurobiol. 157 (1), 4-11.
156
Mitchell, R.A., Loeschke, H.H., Massion, W.H. and Severinghaus, J.W. (1963).
Respiratory responses mediated through superficial chemosensitive areas on the medulla. J.
Appl. Physiol. 18, 523-533.
Morales, R.D. and Hedrick, M.S. (2002). Temperature and pH/CO2 modulate respiratory
activity in the isolated brainstem of the bullfrog (Rana catesbeiana). Comp. Biochem. Physiol.
132A, 477–487.
Muther, T.F. (1972). A critical evaluation of the histochemical methods for carbonic
anhydrase. J. Histochem. Cytochem. 20 (5), 319-330.
Muther, T.F. (1977). On the lack of specificity of the cobalt-bicarbonate method for
carbonic anhydrase. J. Histochem. Cytochem. 25 (9), 1043-1050.
Nasrallah, F.A., Garner, B., Ball, G.E. and Rae, C. (2008). Modulation of brain
metabolism by very low concentrations of the commonly used drug delivery vehicle dimethyl
sulfoxide (DMSO). J. Neurosci Res. 86 (1), 208-214.
Nattie, E.E. and Li, A. (1996). Central chemoreception in the region of the ventral
respiratory group in the rat. J. Appl. Physiol. 81 (5), 1987-1995.
Nattie, E. (1999). CO2, brainstem chemoreceptors and breathing. Prog. Neurobiol. 59 (4),
299-331.
157
Nattie, E. and Li, A. (2006). Central chemoreception 2005: a brief review. Auton.
Neurosci. 126-127, 332-338.
Nattie, E.E. and Li, A. (2008). Central chemoreception is a complex system function that
involves multiple brainstem sites. J. Appl. Physiol. [Epub ahead of print].
Necakov, A., Peever, J.H., Shen, L. and Duffin, J. (2002). Acetazolamide and respiratory
chemosensitivity to CO2 in the neonatal rat transverse medullary slice. Respir. Physiol.
Neurobiol. 132 (3), 279-287.
Neubauer, J. A. (1991). Carbonic anhydrase and sensory function in the central nervous
system. In: Dodgson, S.J., Tashian, R.E., Gros, G. and Carter, N. D. (Eds), The Carbonic
Anhydrases: Cellular Physiology and Molecular Genetics. Plenum Press, New York, pp. 319-
323.
Noronha-de-Souza, C.R., Bicego, K.C., Michel, G., Glass, M.L., Branco, L.G. and
Gargaglioni, L.H. (2006). Locus coeruleus is a central chemoreceptive site in toads. Am. J.
Physiol. 291 (4), R997-1006.
Nurse, C.A. (1990). Carbonic anhydrase and neuronal enzymes in cultured glomus cells
of the carotid body of the rat. Cell Tissue Res. 261 (1), 65-71.
Oka, K. (1958a). The influence of the transection of the brain upon the respiratory
movement of the frog. J. Physiol. Soc. Jpn. 20, 513-519.
158
Oka, K. (1958b). Further studies on the localization of the respiratory centres of the frog.
J. Physiol. Soc. Jpn. 20, 520-524.
Onimaru, H. and Homma, I. (2003). A novel functional neuron group for respiratory
rhythm generation in the ventral medulla. J. Neurosci. 23 (4), 1478-1486.
Parasrampuria, J. and Gupta, V.D. (1989). Preformulation studies of acetazolamide:
effect of pH, two buffer species, ionic strength and temperature on its stability. J. Pharm. Sci. 78,
855-557.
Parthe, V. (1981). Histochemical localization of carbonic anhydrase in vertebrate nervous
tissue. J. Neurosci. Res. 6 (1), 119-131.
Peers, C. and Buckler, K.J. (1995). Transduction of chemostimuli by the type I carotid
body cell. J. Membr. Biol. 144 (1), 1-9.
Peers, C. (2004). Interactions of chemostimuli at the single cell level: studies in a model
system. Exp. Physiol. 89 (1), 60-65.
Pinder, A.W. and Burggren, W.W. (1986). Ventilation and partitioning of oxygen uptake
in the frog Rana pipiens: effects of hypoxia and activity. J. Exp. Biol. 126, 453-68.
Pinder, A.W. (1987). Cutaneous diffusing capacity increases during hypoxia in cold
submerged bullfrogs (Rana catesbeiana). Respir. Physiol. 70, 85-95.
159
Pinder, A.W., Storey, K.B. and Ultsch, G.R. (1992). Estivation and hibernation. In:
Feder, M.E., Burggren, W.W. (Eds.), Environmental Physiology of the Amphibians. University
of Chicago Press, Chicago, pp. 250–276.
Putnam, R.W. (2001). Intracellular pH regulation of neurons in chemosensitive and
nonchemosensitive areas of brain slices. Respir. Physiol. 129 (1-2), 37-56.
Putnam, R.W., Filosa, J.A. and Ritucci, N.A. (2004). Cellular mechanisms involved in
CO2 and acid signalling in chemosensitive neurons. Am. J. Physiol. Cell Physiol. 287 (6), C1493-
1526.
Rand, A.S. (1950). Leopard frogs in caves in winter. Copeia 1950 (4), 324.
Reid, S.G. and Milsom, W.K. (1998). Respiratory pattern formation in the isolated
bullfrog (Rana catesbeiana) brainstem–spinal cord. Respir. Physiol. 114, 239–255.
Reid, S.G., Meier, J.T. and Milsom,W.K. (2000a). The influence of descending inputs
on breathing pattern formation in the isolated bullfrog brainstem–spinal cord. Respir. Physiol.
120, 197–211.
Reid, S. G., Milsom,W.K., Meier, J.T., Munns, S. andWest, N.H. (2000b). Pulmonary
vagal modulation of ventilation in toads (Bufo marinus). Respir. Physiol. 120, 213–230.
160
Reid, S.G. and West, N.H. (2004). Modulation of breathing by phasic pulmonary stretch
receptor feedback in an amphibian, Bufo marinus. Respir. Physiol. Neurobiol. 142 (2-3), 165-
183.
Reid, S.G. (2006). Chemoreceptor and pulmonary stretch receptor interactions within
amphibian respiratory control systems. Respir. Physiol. Neurobiol. 154, 153–164.
Rickli, E.E. and Edsall, J.T. (1962). Zinc binding and the sulfhydryl group of human
carbonic anhydrase. J. Biol. Chem. 237, PC258-260.
Ridderstråle, Y. (1976). Intracellular localization of carbonic anhydrase in the frog
nephron. Acta Physiol. Scand. 98 (4), 465-469.
Ridderstråle, Y. (1980). Intracellular localization of carbonic anhydrase in some
vertebrate nephrons. Acta Physiol. Scand.Suppl. 488, 1-22.
Ridderstråle, Y. and Hanson, M. (1985). Histochemical study of the distribution of
carbonic anhydrase in the cat brain. Acta Physiol. Scand. 124, 557–564.
Ritucci, N.A., Dean, J.B. and Putnam, R.W. (1997). Intracellular pH response to
hypercapnia in neurons from chemosensitive areas of the medulla. Am. J. Physiol. 273 (1-2),
R433-441.
161
Ritucci, N.A., Chambers-Kersh, L., Dean, J.B. and Putnam, R.W. (1998). Intracellular
pH regulation in neurons from chemosensitive and nonchemosensitive areas of the medulla. Am.
J. Physiol. 275 (4-2), R1152-1163.
Rocher, A., Obeso, A., Gonzalez, C. and Herreros, B. (1991). Ionic mechanisms for the
transduction of acidic stimuli in rabbit carotid body glomus cells. J. Physiol. 433, 533-548.
Rosen, S. and Musser, G.L. (1972). Observations on the specificity of newer
histochemical methods for the demonstration of carbonic anhydrase activity. J. Histochem.
Cytochem. 20 (11), 951-954.
Rosen, S. and Friedley, N.J. (1973). Carbonic anhydrase activity in Rana pipiens skin:
biochemical and histochemical analysis. Histochemie. 36 (1), 1-4.
Sakakibara, Y. (1978). Localization of CO2 sensor related to the inhibition of the
bullfrog respiration. Jpn. J. Physiol. 28 (6), 721–735.
Sakakibara, Y. (1984a). The pattern of respiratory nerve activity in the bullfrog. Jpn. J.
Physiol. 34, 269–282.
Sakakibara, Y. (1984b). Trigeminal nerve activity and buccal pressure as an index of total
inspiratory activity in the bullfrog. Jpn. J. Physiol. 34, 827-838.
162
Schoenwald, R.D., Garabedian, M.E. and Yakatan, G.J. (1978). Decreased bioavailability
of sustained release acetazolamide dosage forms. Drug Dev. Ind. Pharm. 4 (6), 599-609.
Scott, W.N. and Skipski, I. (1979). Toad carbonic anhydrase: purification of the enzyme
from erythrocytes of Bufo marinus and comparison with the enzyme activity in the urinary
bladder. Comp. Biochem. Physiol. 63B (3), 429-435.
Sharom, F.J.and Lehto, M.T. (2002). Glycosylphosphatidylinositol-anchored proteins:
structure, function, and cleavage by phosphatidylinositol-specific phospholipase C. Biochem.
Cell Biol. 80 (5), 535-549.
Sherrington, C.S. (1891). Note on Cheyne-Stokes breathing in the frog. J. Physiol. 12 (3),
292-298.
Simkiss, K. (1968). Calcium and carbonate metabolism in the frog (Rana temporaria)
during respiratory acidosis. Am. J. Physiol. 214 (3), 627-634.
Sly, W.S. and Hu, P.Y. (1995). Human carbonic anhydrases and carbonic anhydrase
deficiencies. Annu. Rev. Biochem. 64, 375-401.
Smatresk, N.J. (1990). Chemoreceptor modulation of endogenous respiratory rhythms in
vertebrates. Am. J. Physiol. 259 (5), R887-897.
163
Smatresk, N.J. and Smits, A.W. (1991). Effects of central and peripheral chemoreceptor
stimulation on ventilation in the marine toad, Bufo marinus. Respir. Physiol. 83 (2), 223–238.
Smith, J.C., Ellenberger, H.H., Ballanyi, K., Richter, D.W. and Feldman, J.L. (1991).
Pre-Bötzinger complex: a brainstem region that may generate rhythm in mammals. Science. 254
(5032), 726-729.
Srivaratharajah, K., Cui, A., McAneney, J. and Reid, S.G. (2008). Chronic hypoxic
hypercapnia modifies in vivo and in vitro ventilatory chemoreflexes in the cane toad. Respir.
Physiol. Neurobiol. 160 (3), 249-258.
Stewart, R., Reese, S. and Ultsch, G. (2004). The physiology of hibernation in Canadian
leopard frogs (Rana pipiens) and bullfrogs (Rana catesbeiana). Physiol. Biochem. Zool. 77 (1),
65-73.
Storey, K. B. (2007). Anoxia tolerance in turtles: metabolic regulation and gene
expression. Comp. Biochem. Physiol. 147A (2), 263-276.
Suzue, T. (1984). Respiratory rhythm generation in the in vitro brain stem-spinal cord
preparation of the neonatal rat. J. Physiol. 354, 173-183.
Svichar, N., Esquenazi, S., Waheed, A., Sly, W.S. and Chesler, M. (2006). Functional
demonstration of surface carbonic anhydrase IV activity on rat astrocytes. Glia. 53 (4), 241-247.
164
Takeda, S., Eriksson, L.I., Yamamoto, Y., Joensen, H., Onimaru, H. and Lindahl, S.G.
(2001). Opioid action on respiratory neuron activity of the isolated respiratory network in
newborn rats. Anesthesiology. 95 (3), 740-749.
Tattersall, G. J. and Boutilier, R. G. (1997). Balancing hypoxia and hypothermia in cold-
submerged frogs. J. Exp. Biol. 200 (6), 1031-1038.
Taylor, B.E., Harris, M.B., Coates, E.L., Gdovin, M.J. and Leiter, J.C. (2003). Central
CO2 chemoreception in developing bullfrogs: anomalous response to acetazolamide. J. Appl.
Physiol. 94, 1204–1212.
Toews, D. P. and Macintyre, D. (1978). Respiration and circulation in an apodan
amphibian. Can. J. Zool. 56, 998-1004.
Toews, D.P. and Heisler, N. (1982). The effects of hypercapnia on intracellular and
extracellular acid-base status in the toad Bufo marinus. J. Exp. Biol. 97, 79-86.
Toews, D. P. and Boutilier, R. G. (1986). Acid-base regulation in the Amphibia. In:
Heisler, N. (Ed), Acid-Base Regulation in Animals. Elsevier Science Publishers, Amsterdam, pp.
265-308.
Toews, D.P. and Stiffler, D.F. (1990). Compensation of progressive hypercapnia in the
toad (Bufo marinus) and the bullfrog (Rana catesbeiana). J. Exp. Biol. 148, 293-302.
165
Tong, C.K., Brion, L.P., Suarez, C. and Chesler, M. (2000). Interstitial carbonic
anhydrase (CA) activity in brain is attributable to membrane-bound CA type IV. J. Neurosci. 20
(22), 8247-8253.
Torrance, R.W. (1993). Carbonic anhydrase near central chemoreceptors. Adv. Exp. Med.
Biol. 337, 235-239.
Torgerson, C.S., Gdovin, M.J. and Remmers, J.E. (1998). Fictive gill and lung ventilation
in the pre- and postmetamorphic tadpole brain stem. J. Neurophysiol. 80 (4), 2015-2022.
Travis, D.M. (1971). Molecular CO2 is inert on carotid chemoreceptor: demonstration by
inhibition of carbonic anhydrase. J. Pharmacol. Exp. Ther. 178 (3), 529-540.
Vasilakos, K., Kimura, N., Wilson, R.J. and Remmers, J.E. (2006). Lung and buccal
ventilation in the frog: uncoupling coupled oscillators. Physiol. Biochem. Zool. 79 (6), 1010-
1018.
Vince, J.W. and Reithmeier, R.A. (2000). Identification of the carbonic anhydrase II
binding site in the Cl-/HCO3
- anion exchanger AE1. Biochemistry. 39 (18), 5527-5533.
West, N. H. and Jones, D. R. (1975). Breathing movements in the frog Rana pipiens. I.
The mechanical events associated with lung and buccal ventilation. Can. J. Zool. 53, 332-344.
166
West, N. H., Topor, Z. L. and Van Vliet, B. N. (1987). Hypoxemic threshold for lung
ventilation in the toad. Respir. Physiol. 70 (3), 377–390.
Wilson, R.J., Harris, M.B., Remmers, J.E. and Perry, S.F. (2000). Evolution of air-
breathing and central CO2/H+ respiratory chemosensitivity: new insights from an old fish? J Exp
Biol. 203 (22), 3505-3512.
Wilson, R.J., Vasilakos, K., Harris, M.B., Straus, C. and Remmers, J.E. (2002). Evidence
that ventilator rhythmogenesis in the frog involves two distinct neuronal oscillators. J. Physiol.
540 (2), 557-570.
Wilson, R., Vasilakos, K. and Remmers, J.E. (2006). Phylogeny of vertebrate respiratory
rhythm generators: the oscillator homology hypothesis. Respir. Physiol. Neurobiol. 154, 47-60.
Winmill, R.E., Chen, A.K. and Hedrick, S.A. (2005). Development of the respiratory
response to hypoxia in the isolated brainstem of the bullfrog Rana catesbeiana. J. Exp. Biol. 208,
213–222.
Wang, W., Bradley, S.R. and Richerson, G.B. (2002). Quantification of the response of
rat medullary raphe neurones to independent chages in pHo and PCO2. J. Physiol. 540, 951-970.
Wong, V., Barrett, C.P., Donati, E.J., Eng, L.F. and Guth, L. (1983). Carbonic anhydrase
activity in first-order sensory neurons of the rat. J. Histochem. Cytochem. 31 (2), 293-300.
167
Wong, V., Barrett, C.P., Donati, E.J. and Guth, L. (1987). Distribution of carbonic
anhydrase activity in neurons of the rat. J. Comp. Neurol. 257 (1), 122-129.
Wood, C.M. and Perry, S.F. (1991). A new in vitro assay for carbon dioxide excretion by
trout red blood cells: effects of catecholamines. J. Exp. Biol. 157, 349-366.
Xu, F., Zhang, Z., and Frazier, D.T. (2001). Microinjection of acetazolamide into the
fastigial nucleus augments respiratory output in the rat. J. Appl. Physiol. 91, 2342-2350.
Yu, Z.W. and Quinn, P.J. (1998). The modulation of membrane structure and stability by
dimethyl sulphoxide. Mol. Membr. Biol. 15 (2), 59-68.
Zhu, X.L. and Sly, W.S. (1990). Carbonic anhydrase IV from human lung. Purification,
characterization, and comparison with membrane carbonic anhydrase from human kidney. J.
Biol. Chem. 265 (15), 8795-8801.
Ziegler, T.W., Ludens, J.H. and Fanestil, D.D. (1974). Role of carbonic anhydrase in
urinary acidification by the toad bladder. Am. J. Physiol. 227 (5), 1132-1138.
Zimmer, B. and Milsom, W.K. (2004). Effect of hypothermia on respiratory rhythm
generation in hamster brainstem-spinal cord preparations. Respir. Physiol. Neurobiol. 142 (2-3),
237-249.
