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The hypoxic metabolic response : how time and PO₂ shape the way fishes combine aerobic, anaerobic and… Regan, Matthew D. 2017

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    THE HYPOXIC METABOLIC RESPONSE: HOW TIME AND PO2 SHAPE THE WAY FISHES COMBINE AEROBIC, ANAEROBIC AND DEPRESSED METABOLISM IN HYPOXIC ENVIRONMENTS   by   MATTHEW D. REGAN  B.Sc.Hon., Queen’s University, 2005 M.Sc., The University of British Columbia, 2008    A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF  THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY   in   The Faculty of Graduate and Postdoctoral Studies   (Zoology)   THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)   June 2017    © Matthew D. Regan, 2017  ii  ABSTRACT      Animals rely on O2 to balance cellular ATP supply and demand. In O2-limited hypoxic environments, survival depends on the maintenance of this balance and is accomplished through some combination of aerobic metabolism, anaerobic metabolism and metabolic rate depression (MRD). My thesis studied how fishes combine these three metabolic strategies as a total hypoxic metabolic response (HMR) to survive hypoxic environments that vary in O2 level (PwO2) and duration. Calorimetry is required to accurately measure the metabolic rates (MR) of hypoxia (or anoxia)-exposed fishes that are partially reliant on anaerobic glycolysis and/or MRD. Thus, I started by building a novel calorespirometer that simultaneously measures indices of aerobic metabolism, anaerobic metabolism and MRD, and used it for the remainder of my thesis projects.  Using goldfish, I found that time influences how PwO2 affects HMR. Under acute and continually decreasing PwO2 conditions, goldfish maintained routine O2 uptake rates (ṀO2) to ~3.0 kPa PwO2 (i.e., Pcrit), but sustained routine MR to 0.5 kPa by up-regulating anaerobic glycolysis. Under constant hypoxia (1 or 4 h) at a variety of PwO2s, however, goldfish maintained routine ṀO2 to ~0.7 kPa and consequently reduced their reliance on anaerobic glycolysis. I confirmed this rapidly enhanced O2 uptake ability in subsequent experiments by using different rates of hypoxia induction (RHI) to vary the amount of time goldfish spent at hypoxic PwO2s. Gradual RHIs yielded greater lamellar surface areas, haemoglobin-O2 binding affinities, and subsequently, lower Pcrits than rapid RHIs. However, goldfish only induced MRD below 0.7 kPa. To test the idea that MRD is reserved for extreme hypoxia, I compared two threespine stickleback populations from two isolated lakes: one that experiences deep, long-term hypoxia due to winterfreeze (Alta Lake), and the other that does not (Trout Lake). The two populations did not differ in Pcrit or capacities for anaerobic metabolism, but Alta Lake  iii sticklebacks, which were 2-fold more hypoxia-tolerant than Trout Lake sticklebacks, employed hypoxia-induced MRD while Trout Lake sticklebacks did not. My results reveal that the HMR varies with an animal’s biology and the abiotic aspects of its natural hypoxic environment in a way that may optimize hypoxic survival.                             iv LAY SUMMARY    Animals rely on O2 to maintain the balance of energy supply and demand. O2-limited hypoxic environments threaten this balance, and so a hypoxia-exposed animal’s survival depends on its ability to use aerobic metabolism, anaerobic metabolism and/or metabolic depression to maintain this balance. My thesis investigated how fishes combine these three metabolic strategies in a total hypoxic metabolic response (HMR), a crucial yet unknown dimension of hypoxic survival. Using goldfish and threespine stickleback as study species, I found that the HMR is plastic, influenced by the biology of the animal and the abiotic aspects of its hypoxic environment. Furthermore, the HMRs of goldfish and different stickleback populations appear finely tuned to the animals’ respective natural hypoxic environments, which vary from one another considerably. A species’ HMR may therefore be predictable based on its particular natural hypoxic environment. This knowledge may benefit conservation efforts as the world’s aquatic environments become increasingly hypoxic.                 v PREFACE      A version of Chapter 2 has been published as: Regan, M.D., Gosline, J.M. & Richards, J.G. (2013). A simple and affordable calorespirometer for measuring the metabolic rates of fishes. Journal of Experimental Biology 216: 4507-4513. I co-designed the calorespirometer with the help of Drs. John M. Gosline and Jeffrey G. Richards. I designed and carried out all experiments and analyzed their data. I wrote the manuscript with editorial input from Drs. Gosline and Richards.  A version of Chapter 3 has been published as: Regan, M.D., Gill, I.S. & Richards, J.G. (2017). Calorespirometry reveals that goldfish prioritize aerobic metabolism over metabolic rate depression in all but near-anoxic environments. Journal of Experimental Biology 220: 564-572. I conceived the study and designed the experiments with input from Dr. Jeffrey G. Richards. I carried out all experiments and analyzed their data. Ivan S. Gill assisted with sample preparation for the metabolite assays. I wrote the manuscript with editorial input from Dr. Richards. A version of Chapter 4 has been published as: Regan, M.D. and Richards, J.G. (2017). Rates of hypoxia induction alter mechanisms of O2 uptake and the critical O2 tension of goldfish. Journal of Experimental Biology (doi: 10.1242/jeb.154948). I conceived the study and designed the experiments with input from Dr. Jeffrey G. Richards. I carried out all experiments and analyzed their data. I wrote the manuscript with editorial input from Dr. Richards. All experiments in this thesis were approved by UBC’s Animal Care Committee (Protocol A13-0309). vi TABLE OF CONTENTS  ABSTRACT ............................................................................................................................. ii LAY SUMMARY ................................................................................................................... iv PREFACE ................................................................................................................................ v TABLE OF CONTENTS ...................................................................................................... vi LIST OF TABLES .................................................................................................................. x LIST OF FIGURES ............................................................................................................... xi ACKNOWLEDGEMENTS ................................................................................................ xiii DEDICATION ....................................................................................................................... xv  CHAPTER 1: General introduction ...................................................................................... 1 1.1  Environmental hypoxia ............................................................................................................... 2 1.2  The metabolic responses of fishes to hypoxia ............................................................................. 3 1.2.1  Aerobic metabolism ............................................................................................................. 3 1.2.2  Anaerobic metabolism ......................................................................................................... 5 1.2.3  Metabolic rate depression .................................................................................................... 6 1.3  The hypoxic metabolic response: the concurrent use of aerobic metabolism, anaerobic metabolism and metabolic rate depression in hypoxic environments ................................................ 8 1.4  Thesis objectives and hypotheses ................................................................................................ 9  CHAPTER 2: A simple and affordable calorespirometer for measuring the metabolic rates of fishes ..................................................................................................................... 13 2.1  Summary ................................................................................................................................... 13 2.2  Introduction ............................................................................................................................... 14 2.3  Materials and methods .............................................................................................................. 15 2.3.1  Theory and overview ......................................................................................................... 15 2.3.2  Calorimeter ........................................................................................................................ 16  vii 2.3.4  Fish and reference chambers .............................................................................................. 16 2.3.5  Respirometer ...................................................................................................................... 17 2.3.6  Setup and optimization ...................................................................................................... 17 2.3.7  Water supply and gas mixing ............................................................................................. 18 2.3.8  Heat calibration and measurements ................................................................................... 18 2.3.9  Oxygen uptake measurements ........................................................................................... 19 2.3.10  Experimental animals ....................................................................................................... 19 2.3.11  Experimental protocols .................................................................................................... 20 2.3.12  Data and Statistical Analysis ........................................................................................... 21 2.4  Results and discussion ............................................................................................................... 21 2.4.1  Metabolic heat .................................................................................................................... 21 2.4.2  Oxygen uptake rate ............................................................................................................ 22 2.4.3  Tips on effective calorespirometry .................................................................................... 23 2.4.4  Concluding remarks ........................................................................................................... 24  CHAPTER 3: Calorespirometry reveals that goldfish prioritize aerobic metabolism over metabolic rate depression in all but near-anoxic environments .......................... 32 3.1  Summary ................................................................................................................................... 32 3.2  Introduction ............................................................................................................................... 33 3.3  Materials and methods .............................................................................................................. 35 3.3.1  Study organisms ................................................................................................................. 35 3.3.2  Calorespirometer ................................................................................................................ 35 3.3.3  Hypoxic exposures ............................................................................................................. 36 3.3.4  Comparison of closed-chamber and flow-through calorespirometry ................................ 37 3.3.5  Terminal sampling experiments ......................................................................................... 38 3.3.6  Lactate and ethanol analyses .............................................................................................. 38 3.3.7  O2 equilibrium curves ........................................................................................................ 39 3.3.8  Pcrit calculation ................................................................................................................... 39 3.3.9  Data analysis and statistics ................................................................................................. 40 3.4  Results ....................................................................................................................................... 40 3.4.1  Closed-chamber calorespirometry experiments ................................................................. 40 3.4.2  Flow-through calorespirometry experiments ..................................................................... 40 3.4.3  Whole body anaerobic end-product concentrations ........................................................... 41  viii 3.4.4  Closed-chamber versus flow-through calorespirometry .................................................... 41 3.4.5  Hb-O2 equilibrium curves .................................................................................................. 41 3.5 Discussion .................................................................................................................................. 41 3.5.1  Metabolic responses to hypoxia ......................................................................................... 42 3.5.2  Hb-O2 affinity and initiation of MRD ................................................................................ 45 3.5.3  Ecological implications of MRD ....................................................................................... 45 3.5.4  Conclusions ........................................................................................................................ 46  CHAPTER 4: Rates of hypoxia induction alter mechanisms of O2 uptake and the critical O2 tension of goldfish ........................................................................................... 53 4.1  Summary ................................................................................................................................... 53 4.2  Introduction ............................................................................................................................... 54 4.3  Materials and methods .............................................................................................................. 56 4.3.1  Animals .............................................................................................................................. 56 4.3.2  Respirometry ...................................................................................................................... 57 4.3.3  Parallel hypoxic exposures for physiological measurements ............................................ 60 4.3.4  Gill morphometrics ............................................................................................................ 61 4.3.5  Blood analyses ................................................................................................................... 62 4.3.6  CO2 and nitrogenous end-product measurements .............................................................. 63 4.3.7  Pcrit calculation ................................................................................................................... 63 4.3.8  Data analysis and statistics ................................................................................................. 64 4.4  Results ....................................................................................................................................... 64 4.4.1  Respirometry ...................................................................................................................... 64 4.4.2  Effect of RHI on gill morphology ...................................................................................... 65 4.4.3  Effect of RHI on Hb-O2 affinity, [Hb] and RBC [NTP] .................................................... 65 4.4.4  Effect of RHI on plasma lactate ......................................................................................... 65 4.4.5  Metabolic end-product accumulation ................................................................................ 65 4.5  Discussion ................................................................................................................................. 66 4.5.1  Effects of time on the physiology of O2 uptake ................................................................. 66 4.5.2  Respirometric technique and waste accumulation ............................................................. 69 4.5.3  Implications and recommendations ................................................................................... 71 4.5.4  Conclusions ........................................................................................................................ 72  ix CHAPTER 5: Metabolic depression and the rapid evolution of hypoxia tolerance in threespine sticklebacks, Gasterosteus aculeatus ............................................................. 79 5.1  Summary ................................................................................................................................... 79 5.2  Introduction ............................................................................................................................... 80 5.3  Materials and methods .............................................................................................................. 81 5.3.1  Lakes .................................................................................................................................. 81 5.3.2  Field collection and husbandry .......................................................................................... 82 5.3.3  Hypoxic exposures, calorespirometry and time-to-LOE ................................................... 82 5.3.4  Metabolite assays ............................................................................................................... 83 5.3.5  Statistical analyses ............................................................................................................. 83 5.4  Results ....................................................................................................................................... 84 5.5  Discussion ................................................................................................................................. 85 5.5.1  Metabolic rate depression and hypoxia tolerance .............................................................. 85 5.5.2  Intraspecific variation in hypoxia tolerance ....................................................................... 88 5.5.3  Conclusions ........................................................................................................................ 88  CHAPTER 6: General discussion ........................................................................................ 95 6.1  Thesis highlights and main contributions .................................................................................. 96 6.1.1  Calorespirometry: easier than might be expected .............................................................. 96 6.1.2  Hypoxia-induced MRD: a response reserved for extreme environments .......................... 97 6.1.3  Pcrit: musings on biology and methodology ....................................................................... 98 6.1.4  Threespine stickleback: a potentially powerful model for hypoxia research .................... 98 6.2  The hypoxic environment as sculptor of the HMR ................................................................... 99 6.2.1 Q2: Severe PwO2 + short duration ..................................................................................... 100 6.2.2 Q4: Severe PwO2 + long duration ...................................................................................... 104 6.2.3 Q3: Moderate PwO2 + long duration ................................................................................. 107 6.3  Summary and perspectives ...................................................................................................... 109 REFERENCES .................................................................................................................... 114    x LIST OF TABLES  Table 2-1. Components for calorespirometer construction .................................................... 30	   Table 2-2. Effects of FCCP on metabolic heat of goldfish .................................................... 31	   Table 3-2. Whole body concentrations of lactate and ethanol in goldfish ............................. 52	   Table 5-1. Characteristics of Alta Lake and Trout Lake ........................................................ 93	   Table 5-2. Whole body concentrations of glycogen and lactate in sticklebacks native to             Alta Lake and Trout Lake ..................................................................................... 94	   xi LIST OF FIGURES  Figure 1-1. The aerobic metabolic responses of a typical fish to decreasing water PO2 at three different levels of aerobic metabolism ................................................................... 12	   Figure 2-1. A wiring diagram and schematics for the functional component of the calorespirometer and the PO2 optode chambers ............................................................. 25	   Figure 2-2. A two-dimensional schematic detailing the calorespirometric setup .................. 26	   Figure 2-3. Average millivoltage measured by calorespirometer in response to known quantities of heat liberated within the calorespirometer’s fish chamber ........................ 27	   Figure 2-4. A representative trace showing a 28 h calorespirometry experiment on a single goldfish ........................................................................................................................... 28	   Figure 2-5. Mean measurements of metabolic heat and O2 consumption rate in goldfish held at 40 kPa PO2 pre- and post-exposure to severe hypoxia (0.25 kPa) ............................. 28	   Figure 3-1. Closed-chamber calorespirometry measurements of ṀO2 and metabolic heat in goldfish ........................................................................................................................... 48	   Figure 3-2. Flow-through calorespirometry measurements of ṀO2, metabolic heat, and glycolytic end-products in goldfish held at different PwO2s for 1 and 4 h ..................... 49	   Figure 3-3. A comparison of ṀO2 measurements of goldfish made using closed-chamber and flow-through calorespirometry ....................................................................................... 50	   Figure 3-4. O2 equilibrium curve for the whole blood of goldfish ........................................ 51	    xii Figure 4-1. The effect of water PO2 on the ṀO2 of goldfish exposed to rapid, typical and gradual rates of hypoxic induction .................................................................................. 73	   Figure 4-2. The effect of Pcrit trial duration on the average Pcrit and normoxic ṀO2 values of goldfish ........................................................................................................................... 74	   Figure 4-3. The effect of the rate of hypoxia induction on the total lamellar surface area of goldfish ........................................................................................................................... 75	   Figure 4-4. The effect of the rate of hypoxia induction on the O2 equilibrium curve and Hb P50 value of goldfish ....................................................................................................... 76	   Figure 4-5. The effect of the rate of hypoxia induction on blood parameters of goldfish ..... 77	   Figure 4-6. The effect of closed-circuit respirometry on water chemistry and the buildup of metabolic wastes ............................................................................................................. 78	   Figure 5-1. The time taken for two populations of threespine sticklebacks to lose dorsal-ventral equilibrium when exposed to severely hypoxic water (1.3 kPa PwO2) ............... 89	   Figure 5-2. Closed-chamber ṀO2 measurements as a function of PwO2, normoxic ṀO2, and Pcrits for two populations of threespine sticklebacks ...................................................... 91	   Figure 5-3. Flow-through calorespirometric measurements of metabolic heat and ṀO2 as a function of time in severe hypoxia for two populations of threespine sticklebacks ....... 92	   Figure 6-1. A matrix of environmental hypoxia portraying various natural hypoxic environments according to their PwO2 and hypoxic durations ..................................... 112	   Figure 6-2. The hypoxic metabolic responses of species adapted to natural aquatic hypoxic environments listed in Fig. 6-1 and detailed in the text ................................................ 113	   xiii ACKNOWLEDGEMENTS      The completion of this thesis will see only me acquire three new post-name letters, but many people have played roles in its achievement. These are those people and some of the roles they have played.  Firstly, Jeffrey Richards, devoted pragmatist and gentleman scientist with a low tolerance for conjecture and speculation. I learned many things from Jeff as he guided me through my PhD’s peaks and valleys, but perhaps most influential were his lessons on the necessity of empiricism when making decisions, any decisions, ranging from drawing scientific conclusions to deciding how to invest one’s time. This mode of thinking has permeated all aspects of my life, and each has benefited from it. Less consequential but no less embraced, “compared to” is dead to me. It was a slow death, but it’s death nonetheless.  I also appreciate the guidance of my committee members. Trish Schulte has played a significant role in my development as a scientist throughout both MSc and PhD degrees. She remains my ultimate appraiser of work quality. If she says it’s good, it’s good; if she says it’s crap, it’s definitely crap. On the topic of crap, Tony Farrell was initially unconvinced by our calorespirometer’s function and performance, and my consequent attempts to convince him of its accuracy resulted in a Methods & Techniques paper that otherwise would not have been. And Bill Milsom clarified for me what a hypothesis does and does not comprise, an important bit of knowledge for any experimental scientist. Also, he’s a decent companion for scooter trips around remote communist islands.  John Gosline was absolutely essential to the completion of this thesis. His ingenuity made the calorespirometer possible, and his unshakeable conviction during the early phases that “it will work” gave me the confidence to invest time and energy in a somewhat risky project. As usual, he was right. I am truly indebted to him and miss our conversations about life, science and other things.   Colin “C.J.” Brauner has very much influenced my development as a scientist and a human these past 12 years. The longer I know him, the more astounded I am by the depth to  xiv which he thinks about pretty much everything. And for what it’s worth, also not a bad companion for island scooter adventures should the reader ever need one.  Agnes Lacombe has been a mentor in all sorts of ways, cultivating my love for teaching, encouraging my highest-flying ambitions, and even helping me grow as a husband and son. I don’t know which is most important, but I do know that TAing with Agnes has made my time at UBC considerably richer and more enjoyable.  Ivan Gill, undergraduate student, functioned as my work partner for a portion of my PhD. Ivan taught me something over that time. I don’t know what, exactly, but I think it was important.  I’d be remiss not to acknowledge UBC’s Zoology Department as a whole. From its Head (the delightful Bob Shadwick), to its administrative arm, to its student and faculty bodies, to its Workshop technicians (Bruce, Vinnie, Pak) located somewhere in its nether regions, the Zoology Department has been a wonderful place to spend such a large chunk of my life. I’ll miss the hell outta this place.  I am fortunate to count many fellow graduate students as close friends. (Alphabetically) Rush Dhillon, Milica Mandic, Mike Sackville, Ben Speers-Roesch and Dave Toews go back to pre-PhD days, each playing a significant role throughout both of my graduate degrees. Perhaps one of these days I’ll get around to sharing with each of them just what these roles were, but for now, I’ll just say thank you. And to my other friends who have become dear to me, including (alphabetically) Georgie Cox, Anne Dalziel, Yvonne Dzal, Till Harter, Gigi Lau, Phil Morrison and Ryan Shartau, thank you.  Next, my mom and dad. It’s hard to imagine two more unconditionally supportive people. Their sincere efforts to better understand the physiology of fishes pretty well says it all (and this is coming from a fish physiologist). They are truly wonderful people, and I am lucky to have been born into their lives.  Finally, Emmanuelle. This may be the only part of my thesis you read, but pretty well every page of it has been influenced by you in one way or other. You encouraged me to pursue a PhD, and you encouraged me to devote myself to it. I am forever grateful for your support. I’d ask how I am to ever repay you, but I’m sure you’ll think of something.  xv DEDICATION        For Emmanuelle, my rudder, my sail, and my keel.    1 Chapter 1   General introduction     The maintenance of stable cellular conditions and proper cellular function requires a high turnover of energy in the form of ATP. This ATP can be supplied aerobically via oxidative phosphorylation or anaerobically via substrate-level phosphorylation (e.g., anaerobic glycolysis). Aerobic pathways yield up to 18-fold more ATP per unit of substrate than anaerobic pathways, and perhaps because of this, almost all animals on earth have evolved an ultimate reliance on O2 to supply their cells with sufficient quantities of ATP to match cellular ATP demand. But many environments on earth are low in O2 (i.e., hypoxic), and despite the critical importance of O2 in supplying cellular ATP, a range of anatomically and physiologically specialized animals can be found living in many of them (Bickler and Buck, 2007; Ramirez et al., 2007). These environments include the subterranean, the high-altitude and the aquatic, and the animals living here risk their abilities to supply ATP aerobically, and consequently, risk upsetting the balance of energy supply and demand.  The fact that these animals are living in these environments suggests they routinely maintain their cellular ATP balance (Boutilier, 2001). There are three general mechanisms used to do this at low levels of environmental O2: sustained aerobic metabolism, activation of anaerobic metabolism and metabolic rate depression (MRD). Some combination of these mechanisms allows an animal to maintain energy balance and survive a hypoxic exposure. Much work has been put towards understanding each of these mechanisms in isolation, but very little work has focused on how they are used simultaneously and in combination in what I will refer to as a hypoxic metabolic response (HMR). My thesis will investigate the HMR of fishes, and how it varies with exposure to different hypoxic environments. But before delving into the HMR and the mechanisms it comprises, it would be prudent to first clarify what is meant by ‘hypoxia’.  2  1.1  Environmental hypoxia Environmental hypoxia is most simply defined as a partial pressure of O2 (PO2, PwO2 for water) lower than full air saturation (i.e., less than 157 mmHg or 21 kPa at sea level and 20°C, less any vapour pressure for terrestrial animals). Such environments are naturally prevalent in aquatic systems owing to the density stratification of the water column, O2 consumption on the part of decomposing organic matter, ice cover, algal blooms, tidal cycles, and water body isolation (Diaz and Breitburg, 2009). Furthermore, anthropogenic practices resulting in rising global temperatures and increased eutrophication events have increased the prevalence, severity and duration of these hypoxic events (Boesch, 2002; Diaz and Breitburg, 2009; Diaz and Rosenberg, 1995; Smith et al., 2006). From a general perspective, defining hypoxia in this absolute sense is useful when comparing the responses and/or tolerances of different species to a particular PO2. From an individual’s perspective, however, defining hypoxia involves determining the PO2 at which physiological function is first compromised (Farrell and Richards, 2009), and this requires accounting for the various hypoxic responses that contribute to the individual’s HMR.   Hypoxic environments vary in their hypoxic severities (PO2), hypoxic durations, and rates of hypoxic induction (RHIs). Among aquatic systems, there is high variation in hypoxic environments. For example, oceanic O2 minimum zones are stable hypoxic environments that are characterized by chronic and severe PwO2s (≤4.2 kPa; Seibel, 2011). At the other end of the spectrum, tidepools located high in the intertidal zone are in a constant state of flux, oscillating daily between ~80 kPa and anoxia, and rarely holding stable at any PwO2 (Richards, 2011). And between these on the spectrum of hypoxic environments are winterfreeze lakes, which gradually descend towards anoxia over ~2 months and then hold there until spring thaw because O2 cannot move into the water until the ice melts (Vornanen, 2004). With such variation in PO2, RHI and duration, there is probably no such thing as a ‘grand unified’ strategy of hypoxia tolerance. Rather, the hypoxia defense responses (including HMR) of a species are probably shaped by the particular hypoxic environment to which the species is adapted; because these environments differ, so too will the hypoxic survival strategies of the species that inhabit them. A broad survey of the literature reveals that the hypoxia defense strategies employed by fishes do indeed vary, and this may be the  3 result of adaptation to different types of hypoxic environments. But the inconsistency of the hypoxic exposure protocols across studies, and the fact that the majority of hypoxia studies focus on hypoxic severity (fewer on duration, and almost non on RHI) make such broad comparisons difficult.  1.2  The metabolic responses of fishes to hypoxia 1.2.1  Aerobic metabolism Aerobic metabolism centers on oxidative phosphorylation, the O2-dependent process by which ATP is produced in the mitochondria. Because fishes rely predominantly on aerobic metabolism under normoxic, steady state conditions, O2 uptake rate from the environment (ṀO2) is widely used as a proxy for metabolic rate (Nelson, 2016). There are different levels of metabolic rate that can be supported aerobically (see Fig. 1-1). Standard metabolic rate (SMR) is the ṀO2 of an awake, post-absorptive, and entirely inactive ectothermic animal in its thermoneutral zone (Chabot et al., 2016). It is equivalent to the basal metabolic rates of endothermic animals, and essentially represents the minimal cost of living at a particular temperature. At the other end of the spectrum, maximum metabolic rate (MMR) is the highest ṀO2 an animal can attain in a given environment (Norin and Clark, 2016). The difference between SMR and MMR represents an animal’s aerobic scope, an ṀO2 range that supports higher level functions such as growth, digestion, locomotion and reproduction (Claireaux and Chabot, 2016). Within the aerobic scope lies routine metabolic rate (RMR). Similar to SMR, RMR is the ṀO2 of an awake, post-absorptive, and entirely inactive ectothermic animal in its thermoneutral zone, but RMR also accounts for small movements that are typical of fish under experimental conditions (Chabot et al., 2016). The costs of these movements are typically low, so RMR tends to be far closer to SMR than to MMR (Chabot et al., 2016; Fig. 1-1). Maintaining aerobic metabolism at any rate requires transporting O2 from its origin in the environment to its terminus at the mitochondria. In water-breathing fishes, this process occurs via a five-step cascade: 1, breathing, which brings O2 into contact with the respiratory (gill) surface; 2, branchial diffusion across the water-blood barrier; 3, circulation throughout the body in the bloodstream; 4, tissue diffusion across the blood-mitochondria interface; and 5, use in the mitochondria as an electron acceptor, ultimately producing ATP (Weibel, 1984).  4 Optimizing this cascade to more efficiently move O2 from the environment to the mitochondria can aid hypoxic survival, and hypoxia-adapted animals have evolved traits at each step to do so. Most animals possess mechanisms that enhance O2 extraction and delivery to tissues as environmental PO2 is reduced. For fishes, these mechanisms include increases to gill surface area (Sollid et al., 2003), haemoglobin (Hb) synthesis (Gracey et al., 2001) and concentration in the blood (Affonso et al., 2002), hematocrit (Lai et al., 2006; Turko et al., 2014), Hb-O2 binding affinity (Turko et al., 2014), and ventilation frequency and amplitude (Holeton and Randall, 1967; Itazawa and Takeda, 1978; Tzaneva et al., 2011; Vulesevic and Perry, 2006), as well as a redistribution of blood supply to critical tissues (Sundin et al., 1995). Together, these mechanisms allow fishes to maintain stable ṀO2s across a range of PwO2s (called oxyregulation). Should environmental O2 levels continue to decrease, however, a PwO2 will reached at which the fish’s compensatory mechanisms of O2 uptake can no longer maintain stable ṀO2. This PwO2 is called the critical O2 level (Pcrit), and at PwO2s below this, the fish’s ṀO2 becomes dependent on the environmental O2 level (called oxyconformation; Claireaux and Chabot, 2016). In the context of the aerobic hierarchy shown in Fig. 1-1, a fish has three corresponding Pcrit values: Pcrit-Std, Pcrit-Max and Pcrit-Rtn, the lowest PwO2s at which ṀO2 can support SMR, MMR and RMR, respectively (Fig. 1-1). Investigators typically use Pcrit-Std (Claireaux and Chabot, 2016) or Pcrit-Rtn (McBryan et al., 2013) when discussing Pcrit in the context of hypoxia tolerance. Unless otherwise indicated, I will use Pcrit-Rtn (referred to simply as ‘Pcrit’) throughout this thesis owing to the difficulty in achieving true SMR under experimental conditions (but see Chabot et al. 2016). At Pcrit, the fish’s aerobic scope is either at or near zero (for SMR- and RMR-based estimates, respectively), and at PwO2s below Pcrit, the fish’s ability to supply ATP aerobically is limited (Farrell and Richards, 2009). Pcrit therefore reflects a fish’s ability to acquire and use environmental O2 at stable rates as a function of PwO2, with a lower Pcrit value indicating a greater ability to do so in hypoxic environments. Among species, Pcrit values tend to be lower in hypoxia-tolerant species than in non-tolerant species (Mandic et al., 2009b; Nilsson and Östlund-Nilsson, 2008; Speers-Roesch et al., 2012a), and this is often the result of variation in traits related to O2 extraction (Mandic et al., 2009b; Nikinmaa, 2001; Perry and Reid, 1992; Petersen and Gamperl, 2011; Sollid et al., 2003; Takeda, 1990). Even within an  5 individual, Pcrit can be reduced as a result of hypoxic acclimation (Borowiec et al., 2015; Sollid et al., 2003). This relationship of Pcrit and hypoxia tolerance has led some to use Pcrit as a metric of hypoxia tolerance per se, but the completeness of this connection is (rightly) debated because Pcrit does not incorporate the contributions of anaerobic metabolism and MRD to hypoxia tolerance (Claireaux and Chabot, 2016; Rogers et al., 2016; Speers-Roesch et al., 2013; Urbina and Glover, 2013). In any case, a low Pcrit is beneficial because it allows the animal to maintain RMR (or SMR) in more hypoxic environments while avoiding a reliance on anaerobic glycolysis and/or MRD. But for any fish, should environmental PwO2 decrease below its Pcrit, ṀO2 will decrease, and with it, the fish’s ability to produce ATP aerobically. The balance of cellular ATP supply and demand therefore becomes more reliant on anaerobic glycolysis and/or MRD, and solely reliant on these should PwO2 fall to 0 kPa.  1.2.2  Anaerobic metabolism Fishes have two primary methods of generating ATP anaerobically: substrate-level phosphorylation via creatine phosphate (CrP) and anaerobic glycolysis (hereafter referred to as glycolysis). Substrate-level phosphorylation via CrP occurs in the cytosol of certain tissues (e.g., muscle) and involves the rapid, direct transfer of phosphate from CrP to ADP. Because the cell’s CrP reserves are small, CrP can be quickly depleted, forcing it to rely on glycolysis for the anaerobic supply of ATP should aerobic process remain constrained (Wang and Richards, 2011).  Glycolysis, defined as the splitting of sugars, occurs in the cytosol. The carbohydrate source for anaerobic glycolysis is glycogen, an endogenously stored branched polysaccharide from which glucose monomers are cleaved by glycogen phosphorylase for entry into the glycolysis pathway. Because hypoxia exposure suppresses the appetite and digestive functions of fishes (Wang et al., 2009), species with larger glycogen stores (e.g., goldfish, carp, tilapia) can fuel glycolysis for longer time periods than species with smaller glycogen stores (e.g., rainbow trout), and this prolongs their hypoxic/anoxic survival time (Richards, 2009). When O2 is present in the mitochondria, the two products of glycolysis are transported into the mitochondria, where pyruvate is used as the starting compound for the  6 TCA cycle and a reducing equivalent (from NADH) is used for the electron transport chain. The resulting NAD+ remains in the cytosol where it is again available for further glycolytic activity. When O2 levels in the mitochondria are low, however, the electron transport system becomes unable (or less able) to accept reducing equivalents and NADH accumulates in the cytosol, reducing NAD+ and potentially halting glycolysis altogether. To avoid this, NADH is recycled back into NAD+ in a lactate dehydrogenase-catalyzed reaction that simultaneously converts pyruvate into lactate. When lactate formation rate exceeds its removal rate (as during hypoxia), it accumulates in the tissue. The level of lactate in a tissue is therefore indicative of its O2-independent glycolytic activity and may be used to estimate its reliance on anaerobic ATP production. The formation of lactate results in a parallel and equimolar production of protons from ATP hydrolysis (Hochachka and Somero, 2002). These protons may negatively impact the health and, consequently, hypoxia tolerance of the fish (Driedzic and Gesser, 1994; Nilsson and Östlund-Nilsson, 2008). Most species rely on traditional modes of acid-base regulation (e.g., intrinsic buffering) to cope with this acidosis, but a few especially hypoxia/anoxia tolerant fish species (crucian carp, goldfish, bitterling, and possibly lanternfishes and desert pupfishes) have evolved the ability to produce ethanol, not lactate and protons, as the ultimate end-product of glycolysis (Heuton et al., 2015; Nilsson, 1988; Shoubridge and Hochachka, 1980; Torres et al., 2012; Wissing and Zebe, 1988). This ability reduces the risk of a metabolic acidosis when these species are exposed to long bouts of hypoxia, and goes a long way to explaining their exceptional hypoxia tolerances (Vornanen et al., 2009). Glycolysis is beneficial in O2-limited environments because it allows for an O2-independent supply of ATP. But the relative inefficiency of glycolysis and fact that glycogen, the required fuel for glycolysis, is of finite supply will invariably lead to a problem: should the hypoxic exposure last too long, even the large glycogen stores of a species like goldfish risk being exhausted. To compensate, energetic demands need to decrease.  1.2.3  Metabolic rate depression An effective way to extend the lifetime of a finite fuel supply is to reduce the rate at which it is used. For an animal to survive a particularly long and/or severe bout of hypoxia,  7 this means reducing its ATP consumption rate so as to extend the time period over which its finite glycogen stores can be used. This is accomplished through MRD, which is defined as a reduction in metabolic rate below standard metabolic rate (SMR; Hochachka and Somero, 2002; Richards, 2009). MRD is widely used among species spanning the phyla (Guppy and Withers, 1999), but this discussion will focus on how ectothermic vertebrates use hypoxia-induced MRD. MRD is made manifest through adjustments at the behavioural, physiological, and biochemical levels. Behaviourally, metabolic rate can be reduced through reductions in locomotor activity, feeding, mating and courtship behaviour, and reductions in these behaviours are typical of hypoxia-exposed fishes (Brett and Groves, 1979; McKenzie et al., 1995; Nilsson et al., 1993; Pedersen, 1987; Schurmann and Steffensen, 1994; Wang et al., 2009; Wu, 2009). Physiologically, metabolic rate can be reduced through reductions in growth, digestion, specific dynamic action, gonad development and gametogenesis, and ventilatory effort once PwO2 has dropped below Pcrit (Fitzgibbon et al., 2007; Wang et al., 2009). And biochemically, metabolic rate can be further depressed below SMR through reductions in the major cellular ATP-consuming pathways, including gluconeogenesis, protein turnover and the maintenance of membrane ion gradients. These processes respectively account for 17%, 29% and 53% of the total cellular energy demand of isolated turtle hepatocytes under normoxic conditions (Hochachka et al., 1996). However, when these hepatocytes are exposed to anoxia, these demands are reduced by 100%, 75% and 93%, respectively, resulting in a 90% reduction in total cellular energy demand (Hochachka et al., 1996). The contributions these processes make to total cellular energy demand, and the degree to which they are down-regulated, likely vary across tissues and species (see Bickler and Buck, 2007; Hylland et al., 1997; Vornanen et al., 2009). But so long as they are down-regulated, the result will be a depressed metabolic rate and a prolonged survival time in hypoxia/anoxia.  Despite MRD’s effectiveness of maintaining ATP balance in hypoxic conditions, not all fish species are capable of employing it. For example, zebrafish (Danio rerio), a species native to stagnant, shallow water bodies that regularly become hypoxic (Spence et al., 2008), do not depress metabolic rate when exposed to at least two different sub-Pcrit PwO2s (Stangl and Wegener, 1996; Regan and Richards, unpublished). The reason they have not evolved (or  8 alternatively, secondarily lost) an ability to induce MRD is not known, but it may relate to MRD’s inherent physiological and ecological costs. For torpid mammals, these include oxidative damage (Carey et al., 2000), reduced growth, repair and immunocompetence (Burton and Reichman, 1999), cognitive impairments stemming from neuronal damage (Popov et al., 1992), ceased reproduction (Humphries et al., 2003), and increased susceptibility to predation by aquatic and aerial predators stemming from significantly reduced motor activity (Humphries et al., 2003). These costs may be more or less relevant to a given species depending on its ecological environment and its ability to mitigate them. For a species like zebrafish that is unable to employ MRD, it may be that the costs of MRD outweigh its benefits, especially if there are other ways to maintain ATP balance in their particular hypoxic environment. But almost no work has been done to quantify the costs and benefits that come with hypoxia-induced MRD in fishes.  1.3  The hypoxic metabolic response: the concurrent use of aerobic metabolism, anaerobic metabolism and metabolic rate depression in hypoxic environments  Surprisingly few studies have measured how hypoxia-exposed fishes concurrently use aerobic metabolism, anaerobic metabolism and MRD as a total HMR. The few studies that have suggest that different species have different HMRs. Common sole (Solea solea) appear to induce anaerobic glycolysis at the same PwO2 as their Pcrit for ṀO2 (2.5 kPa), entering a progressively more depressed metabolic state as PwO2 is further reduced (Dalla Via et al., 1994; van den Thillart et al., 1994). On the other hand, Amazonian oscars (Astronotus ocellatus) only become reliant on anaerobic glycolysis at 1.3 kPa PwO2, despite their Pcrit of ṀO2 being at 4.2 kPa (Muusze et al., 1998).  Together, these results suggest that the two species have different glycolytic control mechanisms (i.e., a typical Pasteur effect for sole; a reverse Pasteur effect for oscar), but the different hypoxia exposure protocols employed by each study make this difficult to interpret. Specifically, the common sole studies exposed single fish to 12 h at a single hypoxic PwO2, while the oscar study exposed single fish to progressively deepening hypoxia over ~24 h, holding them at each of seven hypoxic PwO2s for ~3 h each. With only a single end-point for the lactate sample, the dynamics of lactate accumulation (i.e., anaerobic reliance) throughout the exposures is unclear. Also, in the case of oscar, the representative trace of ṀO2 makes it  9 clear that aerobic reliance is affected by time at each hypoxic PwO2. Finally, MRD was not directly measured in these studies, but rather deduced from estimated ATP turnover rates that were calculated from the ṀO2 and lactate data. Real-time measurements of hypoxic metabolic rates, including depressed metabolic rates, require the use of direct calorimetry (discussed in section 1.4).  Few studies have employed calorimetry to measure the metabolic rates of fishes, and of those that have, only two have made some attempt to measure HMR as a function of PwO2 (non have done so as a function of RHI or hypoxic duration). Tilapia (Oreochromis mossambicus) appear to have a Pcrit of ṀO2 between 3.2 and 1 kPa and induce MRD (as indicated by metabolic heat) in the same PwO2 range (van Ginneken et al., 1997). No indices of anaerobic glycolysis were measured in this study, but estimates based on oxycaloric equivalents suggest the fish become more reliant on glycolysis in this same PwO2 range. On the other hand, goldfish (Carassius auratus) appear to have a Pcrit of ṀO2 of ~4.2 kPa while MRD is only induced at ~2.1 kPa (van Ginneken et al., 2004). The authors do not speculate on anaerobic reliance, but it is probably the case that the maintained metabolic heat between 4.2 and 2.1 kPa is owing to an increased reliance on anaerobic glycolysis. These results suggest that while goldfish and tilapia maintain SMR to similar PwO2s, potentially different HMRs are used; goldfish employ anaerobic glycolysis to maintain SMR once O2 becomes limiting, while tilapia arrest metabolic rate once O2 becomes limiting and enter MRD. However, the lack of reliable anaerobic metabolite measurements makes this conclusion speculative, especially with regard to the effect of duration. In fairness, these studies’ objectives were to determine the effects of light and PwO2 on activity and total metabolic rate (tilapia) and to characterize the relationship of heart rate, metabolic rate and PwO2 (goldfish). They are nevertheless informative with respect to HMR, but a better experimental design for the study of HMR would be to simultaneously measure ṀO2, metabolic heat and glycolytic end-products in the same species as a function of PwO2, RHI and hypoxic duration.   1.4  Thesis objectives and hypotheses The objective of my thesis was to determine how fishes combine their use of aerobic metabolism, anaerobic metabolism and MRD in hypoxic environments, and how this use is  10 affected by variation in hypoxic severity, RHI and duration. I used goldfish as my model fish species because they are exceptionally tolerant of hypoxia, are capable of MRD (Addink et al., 1991; van Ginneken 2004), and are well studied with many literature values for comparison (Chapters 3, 4). To further explore the effect of hypoxic duration on HMR (MRD in particular), I used two isolated populations of threespine sticklebacks (Gasterosteus aculeatus) that are native to lakes with different O2 regimes (Chapter 5). But because most of this work required the use of calorimetry, my thesis began with the construction of a novel calorespirometer (Chapter 2). Chapter 2 describes the design and construction of a novel calorespirometer I built in collaboration with Dr. John Gosline. This apparatus was necessary for the accurate measurement of hypoxic metabolic rate (including MRD) because, under severe hypoxia when anaerobic pathways are contributing to ATP production, respirometric measurements using ṀO2 alone will inevitably underestimate metabolic rate. Separating ATP turnover from ṀO2 is therefore necessary when measuring the metabolic rates of animals in hypoxic environments, and the most effective way to do so is by measuring metabolic heat. The catabolic processes—aerobic and anaerobic—that produce cellular ATP are inefficient and generate heat as a byproduct. The same is true for the many energy-consuming processes of the cell. The amount of heat lost by an organism is therefore directly related to its total ATP turnover rate (Kaiyala and Ramsay, 2011; McLean and Tobin, 1987; Mendelsohn et al., 1964), and measuring this heat via calorimetry provides the most accurate measurement of the organism’s metabolic rate. In Chapter 3, I explored the effect of hypoxic severity on HMR, hypothesizing that MRD is employed by goldfish at hypoxic PwO2s (not just anoxia) and initiated at PwO2 just below Pcrit for ṀO2, where the negative impacts of reduced aerobic capacity and increased anaerobic reliance begin to accrue. I tested this hypothesis using a combination of closed-chamber and flow-through calorespirometry to measure ṀO2 (i.e., aerobic metabolism) and metabolic heat (i.e., MRD) at different PwO2s, as well as terminal sampling experiments to measure concentrations of accumulated and excreted glycolytic end-products (i.e., anaerobic metabolism) at these same PwO2s. In Chapter 4, I explored the effect of RHI on HMR, hypothesizing that, compared with rapid RHIs, gradual RHIs will afford an organism more time to alter plastic phenotypes  11 associated with O2 uptake and subsequently reduce their Pcrit of ṀO2. I tested this hypothesis by determining Pcrit values for goldfish exposed to short, typical and long duration Pcrit trials to represent different RHIs, and then ran parallel hypoxic exposures of different RHIs to investigate morphological and physiological traits of goldfish that might play causal roles in a RHI-related shift in Pcrit. In Chapters 3 and 5, I explored the effect of hypoxic duration using two different approaches. In Chapter 3, I used the methods described above to characterize the HMR of goldfish at a variety of PwO2s over acute, 1 h and 4 h timescales. In Chapter 5, I used the threespine stickleback model system to investigate how HMR varies in two isolated stickleback populations that are native to different lakes with different hypoxic regimes, one that experiences winterfreeze-induced long-term hypoxia (Alta Lake), and another that does not experience long-term hypoxia (Trout Lake). Specifically, I hypothesized that the Alta Lake sticklebacks would employ MRD to achieve a greater hypoxia tolerance than the Trout Lake sticklebacks, and I tested this hypothesis by measuring indices of aerobic metabolism, anaerobic metabolism and MRD as described above, and running time-to-loss of equilibrium experiments to quantify the two populations’ hypoxia tolerances. The general outcome of my thesis is a comprehensive understanding of how goldfish simultaneously use aerobic metabolism, anaerobic metabolism and MRD to survive hypoxic environments that vary in severity, RHI and duration. In particular, it advances our understanding of how time influences the use of aerobic metabolism in hypoxia (Chapter 4), and what type of hypoxic environments might favour the least-studied of the three metabolic mechanisms of hypoxia defense, MRD (Chapters 3 and 5). My thesis also makes methodological contributions to the field in the form of a novel calorespirometer, the demonstration of RHI as a significant contributor to the outcome of Pcrit experiments, and the identification of the threespine stickleback model as a powerful system with which to investigate the mechanisms and evolution of hypoxia tolerance.   Fig 1   12    Figure 1-1. The aerobic metabolic responses of a typical fish to decreasing water PO2 at three different levels of aerobic metabolism. Solid curves represent the required ṀO2s to support maximal, routine and standard metabolic rates. Each level of metabolism has its own critical O2 tension (Pcrit), the water PO2 at which the fish’s compensatory mechanisms of O2 extraction and use become insufficient to support maximal, routine or standard ṀO2, respectively. Supporting each respective level of metabolism at water PO2s below Pcrit requires increased reliance on anaerobic ATP supply pathways. 13 Chapter 2   A simple and affordable calorespirometer for measuring the metabolic rates of fishes      2.1  Summary Calorimetry is the measurement of the heat liberated during energy transformations in chemical reactions. When applied to living organisms, it measures the heat released due to the energy transformations associated with metabolism under both aerobic and anaerobic conditions. This is in contrast to the often-used respirometric techniques for assessing energy turnover, which can only make precise measurements under fully aerobic conditions. Accordingly, calorimetry is considered the “gold standard” for quantifying metabolic rate, yet despite this, it remains a seldom-used technique among comparative physiologists. The reasons for this are related to the expense and perceived difficulty of the technique. I have designed and constructed an inexpensive flow-through calorespirometer capable of detecting rates of metabolic heat loss and oxygen uptake (ṀO2) in fishes under a variety of environmental conditions over long-term experiments. The metabolic heat of the fish is detected as a (micro)voltage by a collection of Peltier units wired in series, while oxygen optodes placed on the inflowing and outflowing water lines are used for the calculation of ṀO2. The apparatus is constructed in a differential fashion to account for ambient temperature fluctuations. This paper describes the design and construction of the calorespirometer for ~$1,300 CAD. Using the goldfish (Carassius auratus auratus), I show that the calorespirometer is sensitive to changes in metabolic rate brought about by pharmacological manipulation and severe hypoxia exposures.   14 2.2  Introduction The accurate measurement of metabolic rate has tremendous value across many disciplines in the life sciences. The rate at which an organism consumes and utilizes energy provides insight into its biology from the level of its cells to its ecology (Hochachka and Somero, 2002; Brown et al., 2004). The most widely used method for assessing metabolic rate is through the measurement of oxygen (O2) uptake rate (ṀO2), which, under aerobic conditions, provides a reasonably good estimate of metabolic rate. However, under circumstances like hypoxia, where the metabolic rate of an organism cannot be solely supported by aerobic metabolism and anaerobic processes are utilized to buffer ATP turnover, measurements of ṀO2 could drastically underestimate metabolic rate. This is most evident in cases of anoxia-tolerant organisms like the painted turtle (Chrysemya picta), crucian carp (Carassius carassius), and goldfish (Carassius auratus auratus), where attempts to quantify metabolic rate via respirometry in anoxia are futile due to the organism’s complete reliance on anaerobic processes to support energy turnover. Like aerobic pathways, however, these pathways yield heat as a byproduct, and the total amount of heat lost by an animal to its environment is proportional to its total energy turnover (minus that conserved in carbon bonds) (Mendelsohn, 1964; Mclean and Tobin, 1987; Kaiyala and Ramsay, 2011). Measuring this heat via calorimetry is therefore an effective way of estimating an animal’s metabolic rate in situations where aerobic metabolism may be compromised. Despite direct animal calorimetry being the “gold standard for quantifying the fire of life” (Kaiyala and Ramsay, 2011), it is a seldom-used technique owing to its reputed difficulty and expense when compared with respirometry. These difficulties are especially true when working with ectothermic animals like fishes, whose lower metabolic rates produce less heat compared with similarly sized endotherms. Measuring these low levels of heat requires an especially sensitive calorimeter, and to date, these have been extremely expensive to purchase. Efforts have been made over the years to produce inexpensive systems to measure heat in fishes (Davies, 1966; Stevens and Fry, 1970), but they have not been described in sufficient detail to facilitate their reconstruction. With high-density thermocouple Peltier units being widely available, it should be possible to construct a relatively simple and inexpensive calorimeter of high sensitivity. This chapter describes the construction and testing of such an apparatus, capable of converting a fish’s metabolic heat to  15 a voltage through use of Peltier units and the thermoelectric effect (more specifically, the Seebeck effect). Furthermore, the fish chamber is designed to operate under flow-through conditions to enable environmental manipulations and the simultaneous measurement of inflowing and outflowing partial pressure of O2 (PO2), which can be used to calculate ṀO2 (hence, calorespirometer). I tested the calorespirometer’s function using goldfish, a species that is well known to undergo metabolic rate depression in response to hypoxia/anoxia exposure (van Waversveld et al., 1988; Addink et al., 1991; Stangl and Wegener, 1996; Richards, 2009).  2.3  Materials and methods 2.3.1  Theory and overview The Seebeck effect allows a heat flux to be converted to a voltage as it passes through a thermally conductive element such as a thermocouple. Peltier units are composed of a number of antimony telluride thermocouples connected in series that, with a Seebeck coefficient of 213 µV Kelvin-1 (K), are highly sensitive to temperature change. In building my calorespirometer, I placed a collection of Peltier units between a small fish/reference chamber and a large mass of aluminum so that the metabolic heat produced by the fish would flow through the Peltier units and into the mass of aluminum. The calorespirometer was assembled in a differential fashion with identical fish and reference chambers attached on either side of the aluminum mass. To measure metabolic heat loss from the fish, both chambers were treated identically apart from the presence of a fish (or resistor; see heat calibration and measurements section below) in one side, and I monitored the net voltage between the two chambers. This differential configuration accounted for any fluctuation in ambient temperature. Below, I detail the construction of the apparatus and its major components, and explain how it was assembled to optimize performance. A complete list of its essential and accessorizing components and their costs is shown in Table 2-1.      16 2.3.2  Calorimeter For this section, “calorimeter” will refer exclusively to the component of the calorespirometer responsible for the detection of heat and its conversion to a voltage. This component, shown in Fig. 2-1A, B, was assembled symmetrically with two identical sides centered on a block of aluminum (98 x 48 x 48 mm). Two Peltier units (approx. 40 x 40 x 4.7 mm; 127 couples; Custom Thermoeletric Peltier module 12711-5L31-03CQ) were affixed to each side of this block using an ultra thin layer of silver conductive epoxy (MG Chemical #8331) and connected in series so as to maximize the voltage reading (Fig. 2-1A). A brass block (approx. 78 x 26 x 26 mm) was affixed to the opposite side of each group of Peltier units using the silver conductive epoxy. Brass was ideal for this component as its hardness and machinability allowed for especially thin walls and its high thermal conductivity optimized heat flow. The brass blocks had a 25 mm diameter bore into which a fish or reference chamber could be inserted. Together with the Peltier units and the brass blocks, the central block of aluminum was bolted to another aluminum block (approx. 98 x 152 x 48 mm) into which two cylinders were bored (approx. 25 mm) and through which the fish chamber and reference chamber could be inserted into and removed from the calorimeter’s brass blocks (Fig. 2-1B).  2.3.4  Fish and reference chambers Identical 32 mL flow-through chambers were constructed to serve as the fish chamber and the reference chamber (Fig. 2-1B). These chambers were constructed of stainless steel tubing (approx. 77 mm length, 25 mm OD, 24 mm ID, 0.5 mm wall thickness; McMaster-Carr #6622K152), with a stainless steel cap of 0.5 mm thickness permanently welded to the upstream end of the chamber. Inserted through this cap were stainless steel inflow and outflow water lines (1 mm OD, 0.8 mm ID, 0.1 mm wall thickness), the inflow water running along the chamber’s bottom all the way to the downstream end, and the outflow water line situated at the chamber’s top and mounted flush with the stainless steel cap at the upstream end of the chamber. The water lines were oriented this way to optimize mixing within the chamber and to allow for an easy path of exit for any gas bubbles that may enter the chamber. Finally, a removable Plexiglas cap equipped with a rubber gasket was placed at the downstream end of the chamber through which the fish could be inserted and removed. This  17 cap could also accommodate a PO2 optode (Ocean Optics OR125) that was used to measure the water PO2 (PwO2) within the chamber. Apart from this optode, the fish chamber and reference chamber were identical.  2.3.5  Respirometer For this section, “respirometer” will refer exclusively to the component of the calorespirometer responsible for the measurement of PO2 and determination of ṀO2. This component was built in a flow-through fashion and incorporated exclusively on the fish chamber side of the calorimeter. Small stainless steel chambers of 1 mL (Fig. 2-1C) were built to accommodate PO2 optodes (Ocean Optics OR125) on both the inflow and outflow water lines immediately adjacent the bored-out aluminum block (i.e., as close to the fish chamber as possible), and the difference between the PwO2 values measured by these optodes allowed for the calculation of the fish’s ṀO2. No heat or electrical signals from the activated PO2 optodes could be detected by the calorimeter, thus their use did not affect measurements of metabolic heat loss.    2.3.6  Setup and optimization To provide a thermally stable environment for the calorespirometer, it was placed within an insulated ice chest (Coleman 6-Day XtremeTM) inside an additional enclosure (foam insulation, 5.08 cm in thickness), located within a temperature controlled (20 ± 0.1oC) environmental chamber measuring 3 x 3 x 2.5 m. The insulated ice chest was lined with aluminum blocks totaling approximately 40 kg, and the calorespirometer was placed in the centre of the chest. The aluminum was used as a heat sink, drawing heat from the fish and reference chambers through the Peltier units. The aluminum’s high thermal inertia, a function of its mass, thermal conductivity (237 W m-1 K-1) and (molar) heat capacity (24.2 J mol-1 K-1), made it an especially effective heat sink and ensured the Peltier units accounted for as much of the fish’s metabolic heat as possible. As heat from the chambers flowed through the Peltier units, the net voltage was measured using a Keithley Model 147 nanovoltmeter. The leads from the Peltier units were soldered to the pure copper lead from the voltmeter and this junction was affixed to the aluminum mass using electrical tape to minimize its possible (albeit small) effect on the  18 measured voltage. The amplified signal was then digitally converted using a DATAQ DI-148 data acquisition system and recorded on a Dell Precision M4300 laptop computer using DATAQ WinDaq software.  2.3.7  Water supply and gas mixing The water supplying the fish and reference chambers was sourced from a common 2 L recirculating volume. This volume was held in an insulated ice chest identical to the one housing the calorespirometer (minus the aluminum) and placed adjacently. Water was drawn out of the beaker by a peristaltic pump (Gilson Minipuls 3), pushed into a gas equilibration chamber (see below), and then into the stainless steel tubing supplying the fish and reference chambers. Water flowed into and out of the chambers as described previously, and was returned to the original 2 L beaker for recirculation. To manipulate the gas tension in the fish and reference chambers, gas mixing was done using a precision gas mixer (Corning 192) and the mixed gas was equilibrated with the water supply in two ways. First, the mixed gas was bubbled into the 2 L recirculating supply volume, and second, the mixed gas flowed into a 1.5 L glass gas equilibration chamber within which the supply water flowed through Silastic tubing before flowing into the stainless steel tubing supplying the fish and reference chambers (Fig. 2-2).  2.3.8  Heat calibration and measurements To calibrate the calorimeter, I used three different resistors (9890 Ω, 19 980 Ω and 39 560 Ω; 5% tolerance; resistances measured using a Fluke 73 multimeter whose accuracy was calibrated using 16 different resistors of 1% tolerance) and the following equation:  Q = V2 / R  where Q is heat flow (in watts), V is voltage applied to a resistor (in volts), and R is the resistance of the resistor (in ohms). The resistors were each embedded in an epoxy-filled glass test tube to protect them from the water. During calibration, one of the glass/epoxy-embedded resistors was placed into the fish/reference chamber, the whole unit inserted into the calorespirometer and held under experimental conditions (water flow rate of 22 mL h-1;  19 normoxia; inflow, chamber, and outflow PO2 optodes running) in order to account for any lost metabolic heat owing to the flow-through design. A voltage (measured at 4.98 V and produced by a C-TON Industries PW2-5 model power supply via a two wire system of negligible resistance [< 0.35 Ω] when compared with the resistances of the resistors) was applied to the resistor yielding heat flows of 2.528 mW, 1.251 mW and 0.632 mW for the 9890 Ω, 19 980 Ω and 39 560 Ω resistors, respectively. Once the measured voltage from the calorespirometer stabilized, it was recorded. Calibration with each resistor was performed three times and a calibration curve relating applied heat (in watts) to measured voltage was constructed and used to convert the metabolic heat of fishes, measured in millivolts, to milliwatts.   2.3.9  Oxygen uptake measurements  Oxygen uptake rates (moles of O2 consumed per hour per gram of tissue) were calculated from measurements of inflow and outflow PwO2, water flow rate, and animal weight according to the Fick principle:  ṀO2 = [ΔPwO2⋅αO2⋅flow] / [animal weight]  where ΔPwO2 is the difference in PwO2 between inflowing and outflowing water (in mmHg), αO2 is the solubility coefficient of O2 in water at the experimental temperature (1.8230 µmol mmHg-1 l-1 at 20oC), flow is measured in L h-1, and animal weight is measured in grams. Partial pressure measurements of mmHg were later converted to kPa.  2.3.10  Experimental animals Goldfish (Carassius auratus auratus) of 0.756 ± 0.087 g (mean ± SEM, n = 5) wet body weight were acquired from a commercial fish dealer and held at the Department of Zoology’s Aquatic Facility at The University of British Columbia, Vancouver, BC, Canada. Fish were held at a stocking density of <0.4 g L-1 in a 76 L recirculating system and maintained in well-aerated, dechlorinated City of Vancouver tap water at 20oC under a 12:12 h light:dark cycle. Water in the recirculating system was replaced every 7 to 10 days. Fish were fed to satiation daily (Nutrifin Max Goldfish Flakes) except 24 h before being  20 transferred to the calorespirometer, during which period feeding was suspended. The University of British Columbia Animal Care Committee approved all procedures involving fish (protocol A13-0309).  2.3.11  Experimental protocols Before each experiment, the PO2 optodes were calibrated in air and 100% nitrogen. A single goldfish was then inserted into the fish chamber via the removable cap, and the chamber was sealed and slid into place within the calorespirometer’s brass block. The peristaltic pump was turned on, supplying both fish and reference chambers with oxygenated water at a rate of 22 mL h-1. Water temperature was maintained at 20oC throughout all experimental trials. The fish was allowed to habituate to the chamber for 16 to 18 hours, which was sufficient time to allow for both the thermal equilibration of the calorespirometer and the recovery of the fish from handling stress. After the habituation period, I conducted several experiments designed to test the calorimeter’s function. To ensure it was capable of detecting variation in metabolic heat produced by fishes, I used carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP; Sigma-Aldrich C2920) in an attempt to increase metabolic rate via mitochondrial uncoupling, and benzocaine (Sigma-Aldrich E1501), an anaesthetic, to decrease metabolic rate. These experiments were repeated three times. The next set of experiments were carried out to ensure the apparatus was capable of detecting the previously observed O2 dependent changes in metabolic heat produced by goldfish (van Waversveld et al., 1988; van Waversveld et al., 1989; Addink et al., 1991). Water PO2 was decreased from approximately 40 kPa to 0-0.25 kPa, where it was held for 1.5 h before being returned to normoxia. This experiment was repeated five times. Although the baseline heat signal remained stable over the duration of each run, it fluctuated between runs by ±0.03 mV. In order to accurately determine the baseline heat signal for each experiment, I introduced an overdose of anesthetic in the fish chamber (final concentration ~300 µmol l-1 benzocaine) to euthanize the fish in the chamber at the end of the experiment. The fish’s metabolic heat quickly subsided after the addition of the anesthetic, stabilizing at a baseline value within ~25 min (preliminary experiments showed no further decrease in heat loss rate over 3 hours). After ~1 h of stable baseline reading, the fish was removed from the calorespirometer and the experiment concluded.  21  2.3.12  Data and Statistical Analysis Statistical analyses consisted of one-way analysis of variance that was performed using SigmaStat version 4.0.  2.4  Results and discussion 2.4.1  Metabolic heat The calorespirometer was both stable and sensitive. Under flow-through conditions of 22 mL h-1 and 20oC but without a fish present, heat flow measurements showed very small oscillations (±0.35 mW) around the baseline and there was no net drift in baseline heat detected over 72 hours (data not shown). Changes in PO2 of inflowing water (between 0 and 40 kPa) and turning the PO2 probes on and off had no effect on heat flow (data not shown).    The heat calibration generated a linear relationship between applied wattage and measured voltage (Fig. 2-3; equation of the line mV = 0.1371⋅mW) that could be used to convert the metabolic heat of a fish, measured in millivolts, to milliwatts. The heat pulses also revealed the calorespirometer’s sensitivity to be 141.15 µV mW-1 at a water flow rate of 22 mL h-1, a sensitivity in close agreement with that of the only known commercially available calorespirometer that can accommodate a fish (as used in Addink et al., 1991). The next step involved inserting a fish into the fish chamber to determine if the apparatus was capable of measuring its metabolic heat under fully oxygenated conditions. The representative trace in Fig. 2-4 shows that approximately 15 h were required for the fish to habituate to its new environment (Fig. 2-4) and for the calorespirometer to thermally equilibrate after insertion of the fish (time zero on Fig. 2-4). During these preliminary trials, water PO2 was maintained at ~40 kPa to ensure adequate oxygen delivery and compensate for the low water flow rate (22 mL h-1). The fish’s rate of metabolic heat loss stabilized by 15 hours and remained relatively constant at ~1.5 mW g-1 (Fig. 2-4), with sporadic increases in heat likely due to episodes of activity. To ensure I could detect variation in metabolic heat loss, pharmacological agents with known effects on metabolism were introduced to the fish and reference chambers by briefly transferring the inflow lines from the 2 L water supply beaker to a vessel holding the pharmacological agent. FCCP is an uncoupling agent that increases the permeability of the  22 mitochondrial inner membrane, dissipating the proton gradient used to drive ATP production via oxidative phosphorylation. I predicted this would increase the fish’s metabolic heat, and in fact, sequential additions of 4 µmol L-1 FCCP (up to 12 µmol L-1 FCCP) resulted in incremental increases in metabolic heat, up to a 60.5±10.3% increase compared with controls (P<0.001; Table 2-2). Similarly, benzocaine, a widely used anaesthetic, was administered in the same way with the prediction that it would decrease metabolic heat. A single dose of ~100 µmol L-1 benzocaine resulted in a 68.6±5.1% decrease in heat loss compared with controls (P<0.001). Goldfish have been shown to reversibly depress their metabolic rate by 70 to 80% when exposed to anoxia (van Waversveld et al., 1988; van Waversveld et al., 1989; Addink et al., 1991; Stangl and Wegener, 1996), and my results are consistent with these previous findings (Figs. 2-4, 2-5A). At the end of the 15 hour habituation period, water PO2 was decreased over 90 min to between 0 and 0.25 kPa. The fish was then held at this PO2 for 1.5 hours during which metabolic heat loss decreased and stabilized at an average value that was ~30% that of the average resting level (Fig. 2-5A). When PO2 was returned to ~40 kPa, metabolic heat returned to levels that were equal to pre-hypoxia levels. Following a 2 h recovery period, the fish was euthanized with an overdose of anaesthetic to determine the baseline heat signal as described previously.  2.4.2  Oxygen uptake rate The trends for ṀO2 measurements paralleled those for the metabolic heat measurements discussed above. Specifically, high ṀO2 values were measured over the initial five hours after the fish was introduced to the calorespiromenter, gradually decreasing to stable levels after 12 to 15 hours in the calorespirometer (Fig. 2-4). my mass-specific routine ṀO2 values are higher than those reported elsewhere for goldfish (van Waversveld et al., 1988), but this variance is likely accounted for by differences in size (our fish are 12 times smaller than those used by van Waversveld et al., 1988), fasting regime, and habituation time and conditions between the studies. Upon exposure to anoxia/hypoxia, ṀO2 fell to near-zero levels, returning to routine levels upon the reintroduction of O2 (Fig. 2-5B). In order to maximize the sensitivity of my calorespirometer for heat detection, I used a low rate of water flow through the fish and reference chambers, which affected the time  23 domain over which ṀO2 could be measured. After a change in inflowing PO2, about 60 minutes were required for the PO2 in the outflowing water to stabilize, and thus, during this equilibration period, calculations of ṀO2 were inaccurate. Apart from this period, the fish’s O2 consumption could be accurately and constantly measured in real time in parallel with its rate of metabolic heat loss. It is important to note that this ~60 min equilibration period was not needed for the measurement of metabolic heat; the calorimeter responded instantly to changes in heat and stabilized within ~25 min (Fig. 2-3 inset).  2.4.3  Tips on effective calorespirometry Despite the calorespirometer’s straightforward design and assembly, much attention was needed when preparing the apparatus for experimental use. Central to most of this was the extreme thermal sensitivity of the Peltier units. The differential design of my calorespirometer should theoretically account for fluctuations in ambient temperature, but effort was still required to ensure all heat produced by electrical equipment in the environmental chamber (e.g., computer, voltmeter, peristaltic pump, etc.) was evenly distributed across the enclosed, insulated ice chest. Fans and heat funnels were used for this, and any vulnerable parts on the insulated ice chest (especially drilled holes for the passage of water lines and electrical cables) were patched with form-fitting foam insulation. This was particularly important for holes in close proximity to the Peltier units. Although it was not a problem with my setup, care should also be taken to ensure the voltage reading is not being affected by electrical activity on the circuit into which the voltmeter is plugged.  The accurate and precise determination of a fish’s metabolic rate demands a baseline heat signal that is known and stable. It is possible that with a highly controlled environment and a faithful duplication of experimental setup procedures and the orientation of all components, an identical inter-experiment baseline heat signal can be generated. However, despite my efforts, I noticed an inter-experiment fluctuation in baseline heat signal by ± 0.03 mV (although mean intra-experimental baseline drift was negligible). This required the fish to be euthanized via an overdose of anaesthetic at the end of each experiment as described previously. Although this is not ideal, the accurate and precise determination of the fish’s metabolic rate required it. It is possible that this approach may be needed in other calorespirometers built from this design.  24  Finally, a calorespirometer like the one described here will inevitably come with a few limitations that need addressing. First, the flow-through design that allows for environmental manipulation and long experimental durations means some of the metabolic heat produced by the fish will be washed downstream, resulting in a possible underestimation of its metabolic rate. This effect will be minimized through the use of a relatively low flow rate, and all but eliminated by performing the heat calibration process at the experimental flow rate (See Materials and Methods). Second, the use of a low water flow rate could result in the accumulation of metabolic end products (e.g. CO2) in the fish chamber that could have their own effects on the organism. In my hands, measured PCO2 values never exceeded 1.1 kPa in a typical 24 hour experiment and thus did not likely have a negative effect on the fish’s metabolic rate (Fry et al., 1947). Should the outflowing water contain high PCO2 or metabolic waste, a higher flow rate is recommended, though this will decrease the calorimeter’s sensitivity. Third, as discussed above, there are different time delays for the measurements of ṀO2 and heat loss that must be taken into account when assessing metabolic rate. In general, if both measurements are required, the time resolution for measurements will be approximately 1 to 2 h. And finally, due to the long habituation time required for accurate measurements of ṀO2 and metabolic heat, the animals are in a fasted state. Duration of fasting has been shown to influence metabolic rate (Davies, 1966), so care must be taken to ensure that all animals are treated similarly.   2.4.4  Concluding remarks I have constructed a calorespirometer that is capable of simultaneously measuring the ṀO2 and metabolic heat of fishes, making it possible to measure metabolic rate in environments that compromise aerobic ATP supply pathways. Combined with its low cost of construction and simple, modifiable design, this apparatus is obtainable to most researchers and has the potential to shed light on the metabolic responses of a broad range of species in any number of environments.   25 Figure 1  Figure 2-1. A wiring diagram (A) and schematics for the functional component of the calorespirometer (B) and the PO2 optode chambers (C). Wiring diagram of the four Peltier units (fish chamber side on right, reference chamber side on left). Dashed lines represent positive wires, thick black lines (towards the center) represent negative wires, and white circles represent the soldered junctions between wires (each affixed to the aluminum mass; see Materials and Methods for details). a, Peltier units (Custom Thermoelectric Peltier module 12711-5L31-03CQ); b, central aluminum block; c, pure copper lead to voltmeter. (B) Schematic of the functional component of the calorespirometer. a, Peltier units; b, PO2 optodes (simplified; see (C) below); c, bored out brass blocks; d, 32 mL fish chamber; e, inflowing stainless steel water line; f, outflowing stainless steel water line; g, 32 mL reference chamber; h, aluminum block. This portion of the calorespirometer is embedded within a 40 kg mass of aluminum located within a highly insulated ice chest. (C) Detailed schematic of the PO2 optode chamber. a, 1 mL stainless steel water chamber; b, inflow water line; c, Plexiglas base; d, affixed rubber stopper to tilt chamber and promote the exit of any gas bubbles; e, outflow water line; f, rubber gasket; g, PO2 optode tip. !"#"$"%"&!'"("&!)"("!"#"'"&"(" #"$"%"!" $"'" '"&"(" 26 Figure 2   Figure 2-2. A two-dimensional schematic detailing the calorespirometric setup. a, precision gas mixer; b, 2 L water volume supplying the fish and reference chamber; c, inflowing water lines; d, outflowing water lines; e, peristaltic pump; f, PO2 equilibration chamber; g, PO2 optodes; h, fish chamber; i, Peltier units; j, reference chamber; k, nanovoltmeter; l, data acquisition system; m, data acquisition computer; n, insulated ice chests. The calorespirometer (including PO2 optodes) and water supply are housed within foam insulation-enshrouded ice chests, themselves within a thermally regulated environmental chamber with ambient fluctuations of not more than 0.1oC. All heat-producing electrical equipment is housed outside of the insulated ice chests.       !"# $"#%#&#'#'#(#(#)#*#+#+#,#-#.#/#0#1#2# 2#Fig. 2  27 Figure 3   Figure 2-3. Average millivoltage measured by calorespirometer in response to known quantities of heat liberated within the calorespirometer’s fish chamber. Power supply was measured at 4.98 V, and resistors were measured at 9890 Ω, 19 980 Ω and 39 560 Ω, resulting in milliwattages of 2.528 mW, 1.251 mW and 0.632 mW, respectively. Sample size of 3 for each average value, with error bars representing s.e.m. Trendline is forced through zero. Inset, a rectangular heat pulse of 2.528 mW switched on at “+” and switched off at “≠”.        y = 0.1371x r2 = 0.997 +!Fig. 3  28 Figure 4   Figure 2-4. A representative trace showing a 28 h calorespirometry experiment on a single goldfish of 0.628 g. The fish chamber containing the fish was inserted into the calorespirometer at time zero. This particular run saw a habituation period of ~14 h, a more stable normoxic period of ~10 h, an anoxic exposure of ~1.5 h, and a recovery of ~2.5 h. Water temperature of 20oC throughout. PO2 reading is from the PO2 optode located within the fish chamber (see text for details).        Figure 5  Habituation Normoxia Hypoxia/anoxia Recovery Fig. 4  29   Figure 2-5. Mean measurements of metabolic heat loss (A) and O2 consumption rate (B) in goldfish held at 40 kPa PO2 before (pre) and after (post) exposure to severe hypoxia (0.25 kPa). Average wet body weight of fish was 0.756 ± 0.087 g, and water temperature was 20oC. Sample size of 5 for each, with error bars representing s.e.m. Asterisks denote statistically significant differences (P<0.001).        01234*AMetabolic heat loss (mW g-1)0246810*Pre-exposure(~40 kPa)Exposure(~0.25 kPa)Post-exposure(~40 kPa)BMO2 (µmol h-1 g-1) 30 Table 2-1. Required components and costs for construction of the calorespirometer described in the Materials and Methods section. Calorespirometer-specific components Total cost Aluminum blocks (8) $648  Brass rod (1) $125  Peltier units (4) $78  Ice chest (2) $140  Stainless steel tubing (for chambers) $47  Stainless steel tubing (for water lines) $60  Materials for PO2 optode chambers (2) $75  Plexiglas caps (2) $8  Silver conductive epoxy $5  Styrofoam insulation $80   $1,266  Additional components   Voltmeter and copper lead (1)           - Computer and data acquisition system           - PO2 optodes, hardware, software (3)           - Peristaltic pump and tubing (1)           - Gas mixer (1)           - Temperature controlled environment chamber           - Machining costs           - Table 1-0-1 Prices in Canadian dollars; taxes not included.             31 Table 2-2. Percent increase in heat lost by goldfish when exposed to increasing concentrations of FCCP.      Sham 4.0 µmol l-1 8.0 µmol l-1 12.0 µmol l-1 FCCP 1.0 ± 4.8 25.2 ± 10.8* 47.9 ± 3.5* 60.5 ± 10.3* Table 0-2 All percent increases are relative to resting levels of metabolic heat loss. Water temperature of 20oC. Sample size of 3 for each, with values representing mean ± s.e.m. Asterisks denote statistically significant difference from resting.   32 Chapter 3  Calorespirometry reveals that goldfish prioritize aerobic metabolism over metabolic rate depression in all but  near-anoxic environments      3.1  Summary Metabolic rate depression (MRD) has long been proposed as the key metabolic strategy of hypoxic survival, but surprisingly, the effects of changes in hypoxic O2 tensions (PwO2) on MRD are largely unexplored. I simultaneously measured the O2 uptake rate ( ̇ṀO2) and metabolic heat of goldfish using calorespirometry to test the hypothesis that MRD is employed at hypoxic PwO2s and initiated just below Pcrit, the PwO2 below which ṀO2 is forced to progressively decline as the fish oxyconforms to decreasing PwO2. Specifically, I used closed-chamber and flow-through calorespirometry together with terminal sampling experiments to examine the effects of PwO2 and time on ṀO2, metabolic heat and anaerobic metabolism (lactate and ethanol production). The closed-chamber and flow-through experiments yielded slightly different results. Under closed-chamber conditions with a continually decreasing PwO2, goldfish showed a Pcrit of 3.0±0.3 kPa and metabolic heat production was only depressed at PwO2 between 0 and 0.67 kPa. Under flow-through conditions with PwO2 held at a variety of oxygen tensions for 1 and 4 h, goldfish also initiated MRD between 0 and 0.67 kPa, but maintained ṀO2 to 0.67 kPa, indicating that Pcrit is at or below this PwO2. Anaerobic metabolism was strongly activated at PwO2 ≤1.3 kPa, but only used within the first hour at 1.3 and 0.67 kPa as anaerobic end-products did not accumulate between 1 and 4 h exposure. Taken together, it appears that goldfish reserve MRD for near-anoxia, supporting routine metabolic rate at sub-Pcrit PwO2s with the help of  33 anaerobic glycolysis in the closed-chamber experiments, and aerobically after an initial (<1 h) activation of anaerobic glycolysis in the flow-through experiments, even at 0.67 kPa PwO2.  3.2  Introduction Aerobic pathways of ATP production yield ~18-times more ATP than anaerobic pathways (Hochachka and Somero, 2002). Consequently, environmental hypoxia and the corresponding shift to anaerobic metabolism seriously threaten energy balance by reducing an animal’s ability to generate sufficient ATP to meet metabolic demands. Despite the critical importance of aerobic respiration to the maintenance of metabolic function, many organisms inhabit and thrive in various hypoxic and even anoxic environments (Bickler and Buck, 2007; Ramirez et al., 2007). Fishes are particularly adept at surviving low-oxygen environments, having independently evolved hypoxia tolerance numerous times (Hochachka and Lutz, 2001) due to the relatively high prevalence of hypoxia among aquatic habitats (Boesch, 2002; Diaz and Breitburg, 2009; Diaz and Rosenberg, 1995; Smith et al., 2006). Metabolic rate depression (MRD) has been proposed as the hallmark response enabling hypoxic survival in hypoxia-tolerant animals (e.g., Boutilier and St-Pierre, 2000; Hochachka et al., 1996). MRD is achieved through reductions in whole animal (e.g., locomotion, reproduction, feeding) and cellular (e.g., growth, repair, protein synthesis) processes (Guppy and Withers, 1999; Richards, 2010), reducing ATP demand and rates of anaerobic fuel depletion (glycogen) and waste accumulation (lactate and protons). Although MRD is a well described response to anoxia exposure in a range of animals including fruit flies (Callier et al., 2015), goldfish (Addink et al., 1991 van Waversveld et al., 1989) and turtles (Jackson, 1968), it has been suggested that MRD would also enhance hypoxic survival (Boutilier and St-Pierre, 2000; Hochachka et al., 1996). Indirect (i.e., non-calorimetric) measurements on common frogs (Donohoe and Boutilier, 1998) and direct (i.e., calorimetric) measurements on goldfish (van Ginneken et al., 1994, 2004) and tilapia (van Ginneken et al., 1997) suggest that MRD may be employed at hypoxic O2 tensions (PwO2 for water). Indeed, goldfish reduced metabolic heat by ~31% at ~2.1 kPa PwO2 compared with normoxia (~21 kPa) (van Ginneken et al., 2004), and tilapia reduced metabolic heat by ~40% at ~1.1 kPa PwO2 (van Ginneken et al., 1997). However, it is still unknown how these changes in  34 metabolic heat correspond with changes in aerobic and anaerobic metabolism, and how metabolic heat may be affected by PwO2. MRD would be particularly important at PwO2s below an animal’s critical PwO2 for O2 uptake rate (ṀO2), referred to as Pcrit, which is the PwO2 at which ṀO2 becomes dependent on environmental PO2. Pcrit is largely determined by the O2 binding affinity of hemoglobin (Hb) (Mandic et al., 2009b), and at PwO2s below Pcrit the ability to extract environmental O2 to saturate Hb is constrained and thus unable to support routine metabolic rate (MR) aerobically. The animal can attempt to sustain ATP production at routine levels through an up-regulation of anaerobic glycolysis, but this comes with the depletion of carbohydrate reserves and the accumulation of deleterious anaerobic end-products (Richards, 2009), ultimately limiting hypoxic survival time (Lague et al., 2012; Speers-Roesch et al., 2013). However, if the animal is capable of reducing its energy-consuming processes through a controlled, hypoxia-induced MRD, then it could simultaneously mitigate the negative consequences of reduced ATP production and increased rates of fuel depletion and waste accumulation. I therefore hypothesized that MRD is employed at hypoxic PwO2s and is initiated just below Pcrit, where the negative impacts of reduced aerobic capacity and increased anaerobic reliance begin to accrue. I tested this hypothesis using closed-chamber and flow-through calorespirometry to simultaneously measure ṀO2, MRD (via metabolic heat) and anaerobic glycolysis (via excretion rates of the anaerobic end-product ethanol) in goldfish held at PwO2s ranging from normoxia to anoxia. I also performed terminal sampling experiments on goldfish exposed to the same PwO2s as used in the calorespirometry experiments to fully quantify whole-body anaerobic metabolism. Goldfish were chosen owing to their exceptional hypoxia tolerance and well documented ability to induce MRD (e.g., Addink et al., 1991; van Ginneken et al., 2004; van Waversveld et al., 1989), something not all fish species are capable of (Stangl and Wegener, 1996), and I used calorespirometry because it is the “gold standard” of MR measurements and the only way to accurately do so on hypoxemic animals in real time (see Kaiyala and Ramsay, 2011; Nelson, 2016). Despite the superiority of calorespirometry, only a few studies have measured the metabolic heat of fishes (Chapter 2; Addink et al., 1991; Stangl and Wegener, 1996; van Ginneken et al., 1994, 1997, 2004; van Waversveld et al., 1989) and only three of these (van Ginneken et al., 1994, 1997 2004) have measured  35 metabolic heat at PwO2 other than normoxia and anoxia. While the data from these studies suggest MRD is employed at hypoxic PwO2s, the authors exposed their organisms to progressive hypoxia over sometimes prolonged periods of time and they did not relate their measurements to Pcrit nor directly assess the contributions of anaerobic metabolism at various hypoxic PwO2s. Furthermore, other studies that have attempted to examine the role of MRD and other metabolic and respiratory responses to hypoxia have not attempted, to my knowledge, to directly assess the relative contributions of MRD, aerobic respiration, and anaerobic metabolism at different hypoxic PwO2s over time. Consequently, we still do not have a comprehensive picture of the hypoxic survival strategies of fishes.   3.3  Materials and methods 3.3.1  Study organisms I obtained adult goldfish (Carassius auratus auratus; 2.06±0.39 g wet mass; n=264; sex unknown) from a commercial supplier (Delta Aquatics, Burnaby, BC, Canada) and held them under a 12 h:12 h light:dark cycle in a 76 L recirculating system of aerated, dechlorinated, 17°C water at the University of British Columbia (Vancouver, BC, Canada). Stocking density was <0.4 g L-1 and water in the recirculating system was replaced weekly. I fed the fish to satiation daily (Nutrafin Max Goldfish Flakes) except for 24 h before transfer to the experimental apparatus, when feeding ceased. The University of British Columbia’s Animal Care Committee approved all procedures (protocol A13-0309).  3.3.2  Calorespirometer I used a differential calorespirometer to simultaneously measure metabolic heat and ṀO2 under closed and flow-through conditions. Details on the design and operation of the calorespirometer are described in Chapter 2. Briefly, the metabolic heat of a fish is detected as a voltage by a collection of Peltier units (Custom Thermoelectric Peltier module 12711-5L31- 03CQ, Bishopville, MD, USA) via the Seebeck effect and converted to wattage using an empirically determined calibration coefficient (see Chapter 2). The design of the calorespirometer allows for the simultaneous measurements of metabolic heat and ṀO2 using PO2 optodes (Ocean Optics OR125, Dunedin, FL, USA) placed on the inflowing and outflowing water lines as well as in the fish chamber. To determine ṀO2 under closed- 36 chamber conditions, a PO2 optode within the fish chamber measured the change in PwO2 over sequential 5 min intervals and was then corrected for chamber volume and fish weight according to ṀO2 = (ΔCO2  ΔT-1  V) M-1  where CO2 is O2 content of the water converted to µmol L-1 from PwO2 using the solubility factor of 1.9312 µmol L-1 mmHg-1 (Boutilier et al., 1984), T is the time period over which the change in CO2 is calculated (5 min), V is the fish chamber volume (32 mL) minus the volume displaced by the fish itself, and M is the mass of the fish. To determine ṀO2 under flow-through conditions, the difference in PwO2 between inflowing and outflowing water lines supplying the fish chamber was measured using the same PO2 optode and corrected for flow rate and fish weight according to ṀO2 = ((CiO2 - CoO2)  FR) M-1  where CiO2 and CoO2 are O2 content of inflowing and outflowing water, respectively converted from PwO2 as described above, FR is water flow rate (22 mL h-1), and M is the mass of the fish. Under flow-through conditions, the chamber PwO2 could be held constant for extended time periods, allowing me to measure ṀO2 and metabolic heat at different time points at any desired PwO2.   3.3.3  Hypoxic exposures Individual fish were transferred to a flow-through calorespirometer held at 17°C and a flow rate of 22 mL h-1, and in this apparatus I performed both closed-chamber and flow-through calorespirometry experiments following a 16 h normoxic habituation period. For the closed-chamber experiments (n=8), the trial began by stopping water flow and allowing the fish to reduce PwO2 from normoxia to anoxia over 60-90 min. The experiment was ended when the chamber PwO2 reached anoxia, at which point I introduced a lethal dose of anaesthetic (buffered MS-222, final chamber concentration of 150 mg L-1) to determine the calorespirometer’s baseline heat signature. For the flow-through experiments, inflowing PwO2 was manually adjusted to yield one of four chamber PwO2s over a ~60 min period (20, 1.3, 0.67 or 0 kPa; n=3-6 for each) and the animals were maintained at one of these PwO2s for up to 4 h (referred to as the experimental period). I measured metabolic heat over the full 21 h period (16 h normoxia habituation, 1 h transition to exposure PwO2, 4 h experimental  37 period) and collected effluent water samples either before (time 0) or at 1 and 4 h during the experimental period for measurements of ethanol (a glycolytic end-product excreted across goldfish gills). Following the experiment, I introduced a lethal dose of anaesthetic (buffered MS-222, see above) to determine the calorespirometer’s baseline heat signature. At the end of each experiment, I recalibrated the PO2 optodes to determine any drift that had occurred over the course of the experiment (up to ~10%) for the purpose of later correction, and then washed the calorespirometer and its water lines with a 10% bleach solution. The flow-through and closed-chamber calorespirometry experiments were performed in fall 2014 and winter 2015, respectively.    3.3.4  Comparison of closed-chamber and flow-through calorespirometry In order to more directly compare the results of the closed-chamber and flow-through calorespirometry experiments, I conducted a back-to-back comparison of the two techniques using the same fish. This was required because my first experiments (presented in Figs. 3-1 and 3-2) using these techniques were conducted at different times of year and yielded different routine normoxic ṀO2 values, which could affect my determination of Pcrit. I measured routine ṀO2 using both techniques and determined Pcrit during closed-chamber respirometry (Pcrit was not determined via flow-through calorespirometry because it would require the fish to undergo multiple runs at different PwO2s). Briefly, fish were introduced to the calorespirometer and allowed to habituate under the same conditions as the calorespirometry experiments described above. Following the habituation period, I first measured the fish’s routine ṀO2 under normoxia using flow-through respirometry (as described above), then immediately closed off the respirometer chamber and measured ṀO2 using closed-chamber respirometry (as described above). I repeated this three times for each of six fish, and the PwO2 was not allowed to drop below 16 kPa during these closed-chamber measurements. Following the final closed-chamber measurement, I allowed the fish to deplete the chamber’s O2 content so as to determine its Pcrit. Metabolic heat was not measured during these back-to-back experiments.      38 3.3.5  Terminal sampling experiments To better estimate the effects of PwO2 on anaerobic metabolism, I ran parallel hypoxia exposures where I euthanized animals to measure whole-body concentrations of lactate and ethanol. For each PwO2, I exposed 24 goldfish spread across six 10 L tanks (four fish per tank) and sample two replicate tanks at each of 0, 1 and 4 h to match the experimental periods of the calorespirometry experiments (n=8 per time point). I sampled fish by inconspicuously introducing a lethal dose of anaesthetic (buffered MS-222, see above), weighing the individual fish, freezing them immediately in liquid N2, then storing them at -80°C for later metabolite analyses. I ensured the conditions between these experiments and the calorespirometry experiments were similar by including a 16 h habituation period followed by a 1 h transition period to the desired PwO2, conducting exposures in the dark, and by preventing the fish in the tank from accessing the air-water interface (which was not available to the calorespirometry fish). Thus, the main difference between this experiment and the calorespirometer experiment was vessel size (calorespirometer chamber was 32 mL and exposure tanks were 10 L), which could affect the ability of the fish to move during the hypoxia exposures and yield different levels of lactate and ethanol accumulation. I am however confident that fish movement is minimal in the calorespirometer based on the relatively smooth heat traces observed over the habituation and experimental periods and periodic visual inspection of the fish in the 10 L tanks revealed little to no movement, especially during the hypoxia exposures. Thus, despite the differences in exposure regimes, the fish from both the calorespirometry and the terminal sampling experiments likely responded to hypoxia in a similar manner.  3.3.6  Lactate and ethanol analyses In order to link my whole body calorespirometry measurements of MR to the activation of anaerobic metabolism, I measured whole body concentrations of lactate and ethanol. Entire goldfish from the 10 L tank exposures were ground into a fine powder using a liquid N2-chilled mortar and pestle. To extract the metabolites (lactate and ethanol) from the powder, an aliquot of powder was weighed and transferred to a 2 mL centrifuge tube containing 1 mL of ice cold 30% HClO4 and immediately homogenized at 0°C using a Polytron homogenizer set to the highest setting for 30 s. The resulting homogenate was then  39 centrifuged at 20 000 g for 5 min at 4°C and the supernatant was transferred to a new 1.5 mL centrifuge tube and neutralized using 3 M Tris base to avoid the volatilization of ethanol that occurs in association with vigorous CO2 production when HClO4 is neutralized with K2CO3. I confirmed that neutralization with Tris base does not affect my enzymatic analysis. I measured ethanol immediately following neutralization using a commercial kit designed for biological ethanol analysis (Diagnostic Chemical Ltd., PEI, Canada), and then froze the unused portion of the sample extract for later lactate analysis. Lactate concentration was measured using the LDH reaction according to the protocols outlined in Bergmeyer (1983).  3.3.7  O2 equilibrium curves To understand how goldfish’s Hb-O2 affinity related to Pcrit and MRD, I constructed O2 equilibrium curves for the whole blood of five normoxia-acclimated goldfish using the thin film spectrophotometric technique (Lilly et al., 2013). Blood was collected from the caudal artery of anaesthetized fish using 60 µl heparinized capillary tubes. I then centrifuged the tubes and resuspended the red blood cells in HEPES buffer (pH 7.8) to ensure a consistent blood pH across all samples. A Wostoff gas mixing pump (H. Wösthoff Messtechnik GmbH, Bochum, Germany) mixed compressed O2 and N2 to each of seven PO2s for the construction of the O2 equilibrium curves, and Hb P50 values (the PO2 at which the blood is 50% saturated with O2) were calculated using the equation of each sigmoidal curve as calculated by SigmaStat 11.0.  3.3.8  Pcrit calculation Pcrit is defined as the PwO2 at which an organism’s routine ṀO2 transitions from being independent of, to being dependent upon, PwO2. I determined Pcrit for each individual in the closed-chamber calorespirometry experiments using the BASIC program (Yeager and Ultsch, 1989), which uses a two-segment linear regression model to determine Pcrit as the PwO2 at which the two linear trend lines intersect on a graph plotting ṀO2 as a function of PwO2. Some individuals’ ṀO2s increased above routine ṀO2 levels at hypoxic PwO2s close to Pcrit, and including these ṀO2 values would overestimate Pcrit. To prevent this, I excluded from my routine ṀO2 estimation any ṀO2 value that exceeded 1.5 times the standard deviation of an individual’s average ṀO2 between 13 and 21 kPa PwO2.  40  3.3.9  Data analysis and statistics ṀO2 and ethanol production rates were calculated at each time point, while metabolic heat was represented by averaging the continual heat measurements made over the 20 min straddling the time point (e.g., 50-70 min for 1 h time point). All data are presented as means ± s.e.m. The effects of PwO2 on each variable were determined using one-way ANOVA (SigmaStat 11.0).  3.4  Results 3.4.1  Closed-chamber calorespirometry experiments  I used closed-chamber calorespirometry to measure Pcrit and to characterize the effects of a progressive reduction in PwO2 on ṀO2 and metabolic heat. PwO2 in the closed-chamber experiments was decreased from normoxia to anoxia by the fish’s own ṀO2 over 60-90 min (depending on the fish’s ṀO2). Pcrit was calculated to be 3.0±0.3 kPa (Fig. 3-1). At PwO2s above Pcrit, there were no significant effects of changes in PwO2 on the average routine ṀO2, while at PwO2s below Pcrit (at which the fish spent ~30 min), ṀO2 progressively fell to zero as the goldfish depleted the available oxygen. Metabolic heat was maintained at routine levels at all PwO2s between 20 and 0.5 kPa (Fig. 3-1) but was depressed upon reaching anoxia, eventually stabilizing at ~21% of routine normoxic values (a MRD of 79%; Fig. 3-1) after ~20 min.  3.4.2  Flow-through calorespirometry experiments I used flow-through calorespirometry to characterize the effects of PwO2 and time on ṀO2, metabolic heat and excreted ethanol. I held individuals at one of four PwO2s (20, 1.3, 0.67 or 0 kPa) for 1 and 4 h. For 1 h exposures, ṀO2 and metabolic heat were maintained at routine levels to PwO2 of 0.67 kPa, while at PwO2s below this, ṀO2 fell to zero and metabolic heat fell to ~32% of routine levels (a MRD of 68%; Fig. 3-2A). Similarly, for 4 h exposures, ṀO2 and metabolic heat were maintained at routine levels to PwO2 of 0.67 kPa, while at PwO2s below this, ṀO2 fell to zero and metabolic heat fell to ~20% of routine levels (a MRD of 80%; Fig. 3-2B).  41 Ethanol excretion rates were undetectable following 1 h exposure at all PwO2s (Fig. 3-2A). These rates increased following 4 h exposure, and higher rates were generally detected at lower PwO2s (Fig. 3-2B), but these increases were not statistically significant (Fig. 3-2B).  3.4.3  Whole body anaerobic end-product concentrations Whole body concentrations of lactate significantly increased over time 0 values following 1 and 4 h at 1.3 kPa, 0.67 kPa and anoxia (Table 3-1). Whole body concentrations of ethanol significantly increased over time 0 values following 1 and 4 h of anoxia exposure (Table 3-1). The total anaerobic end-product concentrations at 1.3 and 0.67 kPa following 4 h were similar to those following 1 h, suggesting the rate of anaerobic end-product accumulation fell to near-zero levels after 1 h (Fig. 3-2A,B). A similar result was observed for the anoxia-exposed fish though to a lesser extent, with anaerobic end-product concentrations being ~1.8-fold higher following 4 h exposure than following 1 h exposure (Fig. 3-2A,B).  3.4.4  Closed-chamber versus flow-through calorespirometry   Individual ṀO2 values determined in the same fish in a back-to-back comparison of closed-chamber and flow-through respirometry were positively correlated (Fig. 3-3A; n=18, r=0.925, P<0.0001) and yielded similar mean ṀO2 values (Fig. 3-3B; t=0.423, P=0.678). The closed-chamber portion of these experiments yielded a Pcrit of 2.7±0.2 kPa (n=6).  3.4.5  Hb-O2 equilibrium curves  The Hb of goldfish displayed a very high affinity for O2, resulting in a steep O2 equilibrium curve and an average whole blood P50 of 0.49±0.12 kPa (Fig. 3-4).  3.5 Discussion I hypothesized that goldfish employ MRD at hypoxic PwO2s and initiate it at PwO2s just below Pcrit. This hypothesis predicted that metabolic heat would decrease from routine levels at a PwO2 below Pcrit, when the fish’s ability to take up environmental O2 to support a routine ṀO2 was compromised. The closed-chamber calorespirometry experiments yielded a Pcrit of 3.0±0.3 kPa (Fig. 3-1), consistent with the Pcrit values reported in other studies on  42 goldfish (Fry and Hart, 1948; Fu et al., 2011). However, contrary to my hypothesis that MRD is initiated at hypoxic PwO2s just below Pcrit, metabolic heat was maintained at routine normoxic levels to a PwO2 of 0.67 kPa and MRD was only evident in goldfish exposed to anoxia. The magnitude of the anoxia-induced MRD [79% depression in closed-chamber experiments; 68% (1 h) and 80% (4 h) in flow-through experiments; Figs. 3-1, 3-2] was very similar to what has been shown previously for anoxia-exposed goldfish using calorimetry (Addink et al., 1991; Stangl and Wegener, 1996; van Ginneken et al., 1994).  3.5.1  Metabolic responses to hypoxia Goldfish maintained routine metabolic rate (MR) at severely hypoxic PwO2s under both closed-chamber and flow-through conditions (Figs. 3-1, 3-2), but appear to have used different strategies to do so. In the closed-chamber experiments, metabolic heat was maintained at routine normoxic levels to 0.67 kPa despite a decrease in ṀO2 at 3.0 kPa (Fig. 3-1), suggesting anaerobic glycolysis was up-regulated to support MR (though lactate and ethanol could not be measured in closed-chamber experiments as a function of PwO2). In the flow-through experiments, metabolic heat was similarly maintained at routine normoxic levels to 0.67 kPa at both 1 and 4 h, but unlike the closed-chamber experiments, ṀO2 was maintained at near-routine levels at all hypoxic PwO2s tested. This suggests that MR was supported aerobically even at severely hypoxic PwO2s and that MRD is reserved for all but severely hypoxic (<0.67 kPa) or near-anoxic environments. This is different than the results of van Ginneken and colleagues (1994, 2004) who showed moderate 27% and 33% decreases in heat production along with lower ṀO2 in goldfish exposed to 3.5 kPa and 2.1 kPa, respectively. These incongruent results are likely due to differences in experimental design and study goals. van Ginneken et al. (1994, 2004) exposed each fish in their studies to progressive hypoxia over prolonged periods of time (e.g., 8.4, 4.2, 2.1 and finally 0.63 kPa over a 16 h period in van Ginneken et al., 2004), which does not allow the authors to disentangle the effects of PwO2 and time on metabolic heat and ṀO2. In contrast, my flow-through calorespirometry experiments exposed goldfish to only a single hypoxic PwO2 (after a 1 h adjustment period) for up to 4 h and I assessed the effects of varying hypoxic PwO2s using different individuals, allowing me to independently assess the effects of PwO2 and time on metabolic responses. Using this approach, I clearly show that within 1 h exposure,  43 goldfish are capable of maintaining oxygen uptake under severely hypoxic conditions (0.67 kPa), obviating the need for hypoxia-induced MRD.   Elevated levels of lactate and ethanol at PwO2 ≤ 1.3 kPa at 1 and 4 h indicate that anaerobic glycolysis also contributed to maintaining MR, though in slightly different ways in anoxia and hypoxia. In anoxia, lactate and ethanol levels continued to increase throughout the 4 h exposure but their rate of accumulation decreased from 5.81 µmol h-1 g-1 during the first hour to 1.73 µmol h-1 g-1 during the subsequent 3 h. These results are consistent with those observed in tissues from anoxia-exposed turtles (Trachemys scripta elegans), where lactate production rates were elevated during the first hour of anoxia exposure and subsequently decreased between 1 and 5 h anoxia in brain, liver and white muscle (Kelly and Storey, 1988). Combined, these results suggest that there is an initial reliance on anaerobic metabolism upon anoxia exposure that may compensate for the anoxia-induced limitations on aerobic ATP production while MRD is initiated. In hypoxia, the early reliance on anaerobic metabolism was temporally even more profound than in anoxia. Lactate and ethanol accumulation was confined entirely to the first hour of hypoxia exposure at 1.3 and 0.67 kPa, while ṀO2 was concurrently maintained at routine normoxic levels throughout the hypoxic exposures. Taken together, these data suggest that total ATP turnover is higher over the first hour of hypoxia exposure than in normoxia. Indeed, whole-body estimates of total ATP turnover during this period indicate that it increases from ~10 µmol h-1 g-1 in normoxia to ~17 and 15 µmol h-1 g-1 at 1.3 and 0.67 kPa, respectively (assuming P:O2 of 6 and ATP:lactate/ethanol of 1), while heat production does not change. These inconsistences are likely a consequence of not being able to temporally match my measurements of anaerobic metabolism (taken as the delta accumulation of lactate and ethanol over the entire hour plus the PwO2 adjustment period) and ṀO2 and heat, which were taken at the end of the 1 h (between 50 and 70 min exposure). As such, it is possible there are temporal shifts in fuel selection within the first hour of hypoxia exposure, with lactate and/or ethanol accumulating during the initial descent towards the target PwO2 as ṀO2-sustaining mechanisms are up-regulated. Finer scale studies are needed to confirm this idea. The Pcrit values derived from the closed-chamber and flow-through calorespirometry experiments differed substantially, with Pcrit shifting from 3.0 kPa in the closed-chamber experiments to somewhere between 0 and 0.67 kPa in the flow-through experiments (the  44 exact value cannot be determined). These technique-specific differences in Pcrit are consistent with a recent study comparing closed and intermittent-flow respirometry (Snyder et al., 2016), which attributed the higher Pcrit in closed respirometry to metabolic waste accumulation and a faster decline in PwO2. Similar factors may be at play in my closed-chamber calorespirometry experiments resulting in an overestimation Pcrit. Another possible explanation might be that the routine normoxic ṀO2 (and heat) in the closed-chamber experiments was ~2-fold higher than in the flow-through experiments (c.f. Figs. 3-1, 3-2). All else being equal, this would necessitate the fish from the closed-chamber experiments adopting an oxy-conforming strategy at a higher PwO2, yielding a higher Pcrit. The back-to-back comparison of the calorespirometry techniques suggests that time of year may influence measured ṀO2 values, causing upwards of a two-fold change in ṀO2 and heat production, but the differences in ṀO2 obtained in closed-chamber respirometry do not appear to affect Pcrit and therefore do not explain why Pcrit is higher in the closed-chamber experiment relative to the flow-through experiment.  Another factor possibly contributing to the lower Pcrit values obtained from the flow-through experiments versus those from the closed-chamber experiments is time. Fishes possess many mechanisms that enhance O2 uptake with decreasing PwO2, including increases to gill surface area (Sollid et al., 2003), Hb synthesis (Gracey et al., 2001) and concentration in the blood (Affonso et al., 2002), hematocrit (Lai et al., 2006; Turko et al., 2014), Hb-O2 affinity (Turko et al., 2014), ventilation frequency and amplitude (Holeton and Randall, 1967; Itazawa and Takeda, 1978; Tzaneva et al., 2011; Vulesevic and Perry, 2006), as well as redistributed blood supply to critical tissues (Sundin et al., 1995). While these mechanisms effectively enhance the uptake of environmental O2 and its distribution throughout the body, their induction takes time, varying from minutes to days depending on the physiological response examined. Because the fish in the flow-through experiments had spent 1 or 4 h at each PwO2 when their ṀO2 was measured (in addition to the ~1 h required to reduce the PwO2 from normoxia to the target PwO2), they may have had additional time to initiate some of these mechanisms of enhanced O2 uptake compared to the closed-chamber fish that saw only ~30 min of continually decreasing sub-Pcrit hypoxic conditions. If Pcrit is in fact influenced by the rate and duration of hypoxia induction over relatively short time scales, then it becomes important to apply similar methodologies and time courses both within and  45 between studies (something that is not currently done; see Rogers et al., 2016) to ensure Pcrit values are comparable. This is especially true when Pcrit is used as a reference point for models that, for example, predict how climate change will reshape the distribution of fishes around the world (Deutsch et al., 2015).  3.5.2  Hb-O2 affinity and initiation of MRD Our results show that MRD is initiated in goldfish at a PwO2 somewhere between 0 and 0.67 kPa. Interestingly, my analysis of whole blood Hb-O2 affinity reveals a Hb P50 value of 0.49±0.12 kPa (Fig. 3-4; consistent with Burggren, 1982), within the PwO2 range that goldfish appear to reduce ṀO2 in the flow-through experiments and initiate MRD. It is therefore tempting to think of a causal link between the supply of O2 to the tissues and the initiation of MRD. Considerable debate exists regarding the signal for MRD, with some data supporting signals residing on the energy production side of the cellular energy flux pathways (Bishop and Brand, 2000; Bishop et al., 2002; De Zwaan and Wijsman, 1976; Hochachka, 1982; Hochachka, 1985; Plaxton and Storey, 1984; Rees and Hand, 1991) and some data supporting signals on the energy consumption side (Caligiuri et al., 1981; Flanigan and Withers, 1991; Robin et al., 1979; Sick et al., 1982; see reviews by Guppy, 2004; Guppy and Withers, 1999; Storey and Storey, 1990). If Hb-O2 affinity were in fact a signal for MRD, this would place the signal on the energy production side, consistent with some of the more recent views in the field (see Guppy, 2004). Similarly, Coulson (1977) postulated that metabolic rate was directly proportional to the circulatory system’s ability to supply the tissues with O2, and this idea gained empirical support when van Ginneken et al. (2004) showed a correlation between hypoxia-induced decreases in metabolic rate and heart rate. All told, it is not unreasonable to speculate that a signal for hypoxia-induced MRD involves the supply of O2 to the tissue. The association between Hb P50 and the PwO2 of MRD initiation is therefore enticing and worth further investigation.    3.5.3  Ecological implications of MRD The fact that goldfish appear to initiate MRD only near anoxia and maintain ṀO2 without a long-term activation of anaerobic metabolism is well suited to the fish’s (and the closely related crucian carp’s, Carassius carassius) natural lake habitat. While these lakes  46 become ice-covered in winter and eventually anoxic, they are severely hypoxic (Vornanen, 2004) for most of the winter at PwO2s that my study reveals goldfish remain aerobic. Goldfish can therefore maintain routine MR for most of the winter without relying on anaerobic glycolysis and/or MRD until it is entirely necessary. This strategy conserves the goldfish’s finite anaerobic fuel stores (glycogen), reduces the accumulation of deleterious anaerobic end-products (lactate, protons and ethanol), and allows the goldfish to retain routine function and behaviour under most natural conditions.  Another benefit of a near-anoxic induced MRD is a delayed accumulation of MRD’s inherent physiological and ecological costs (Humphries et al., 2003). These include oxidative stress resulting from the production of reactive O2 species (Carey et al., 2000), impaired immunocompetence resulting from reduced lymphocyte production (Burton and Reichman, 1999), impaired cognitive and memory function resulting from reductions in synaptic contacts and dendritic branching (Popov et al., 1992), and significant reductions in sensory and motor activity (Choi et al., 1998) that increase predation susceptibility. Because the costs associated with each of these likely accumulate with time, a hypoxic survival strategy that involves an extended bout of MRD is likely to cause significant damage regardless of its effectiveness to balance cellular energy supply and demand. Goldfish’s predominant reliance on aerobic respiration is therefore the ideal strategy for surviving long-term hypoxic bouts because it minimizes the time the fish is forced to rely on MRD and the physiological costs that come with it. Taken together, goldfish’s overall hypoxia tolerance strategy appears finely tuned to its particular hypoxic environment characterized by long, protracted descents into eventual anoxia. This may be the case with other species too; because hypoxic environments vary greatly in their severity, duration and rate of hypoxic induction, the hypoxia tolerance strategies employed by organisms native to these different environments are likely to be just as variable.  3.5.4  Conclusions By demonstrating that goldfish prioritize O2 uptake over MRD in all but near-anoxic environments, my results suggest two things. First, the exceptional hypoxia tolerance of goldfish owes more to its O2 extraction abilities than to MRD. Second, MRD is not  47 necessarily a key mechanism of hypoxic survival as has been hypothesized (Hochachka et al., 1996), but of anoxic survival. While MRD is an effective means of balancing energy supply and demand, the potential costs associated with reducing cellular and whole-body processes may threaten organismal fitness and preclude its selection in all but the most extreme environments.         48  Figure 6   Figure 3-1. Closed-chamber calorespirometry measurements of ṀO2 and metabolic heat in goldfish. PwO2 was reduced from normoxia to anoxia over 60 to 90 min due to the fish’s O2 consumption. Data are mean±s.e.m, n=8. Data points sharing a letter are not significantly different (1-way ANOVA, P>0.05).               !"#$%&'()'0 1 2 3 4 5 6 200246801234   bab,cdddd d,e d,ed,ec,d,eb,c,eabbbbbbbbbbbWater PO2 (kPa)MO2 (µmol h-1 g-1)Metabolic heat (J g-1) 49 Figure 7   Figure 3-2. Flow-through calorespirometry measurements of ṀO2, metabolic heat, and glycolytic end-products in goldfish held at different PwO2s for 1 (A) and 4 (B) h. Data are mean±s.e.m, n=3-6, and data points are offset slightly on the X-axis for clarity. Data points sharing a letter are not significantly different (1-way ANOVA, P>0.05).    0.00.51.01.52.02.53.00.00.40.81.2036912150.00.20.40.60.81.00.00.51.01.52.02.53.00.00.40.81.2036912150.00.20.40.60.81.0Metabolic heat (J g-1) Ethanol excretion rate (µmol h-1 g-1) MO2 (µmol h-1 g-1) A B a b b c a a a a a b b a a a a a a a b b b b c a Water PO2 (kPa) 0 1 2 3 19 20 21 Whole body lactate + ethanol (µmol g-1) !"#$%&'()'0 1 2 3 19 20 21  50 Figure 8  Figure 3-3. A comparison of ṀO2 measurements of goldfish made using closed-chamber and flow-through calorespirometry. Both respirometric techniques were performed in the same apparatus following a ≥16 h habituation period, at the same time of day (~10:00am PST), and at PwO2 ≥16 kPa. In (A), ṀO2 values resulting from closed and flow-through techniques, with measurements made back-to-back on the same fish (n=18, r=0.925, P<0.0001). In (B), average ṀO2 measurements for each technique, with error bars representing s.e.m. (n=18; t=0.423, P=0.678).   !"#$%&'()'2 3 4 52345AClosed-chamber MO2 (µmol h-1 g-1)Flow-through MO2 (µmol h-1 g-1)01234Closed-chamber Flow-throughBRoutine normoxic MO2 (µmol h-1 g-1)  51 Figure 9   Figure 3-4. O2 equilibrium curve for the whole blood of 5 normoxia-acclimated goldfish. Red blood cells were separated from plasma and resuspended in HEPES buffer (pH 7.8). O2 levels were achieved using a Wösthoff gas mixing pump attached to cylinders of compressed O2 and N2.         !"#$%&'()'0 1 2 3 4 5 6 22020406080100PO2 (kPa)Hb P50 = 0.49±0.12 kPaHb-O2 saturation (%) 52 Table 3-1. Whole body concentrations (µmol g-1) of lactate and ethanol in goldfish exposed to different PwO2s for different periods of time. Different letters indicate significant differences between time points within a PwO2 exposure (P>0.05). [Ethanol] measurements at 20 kPa were not taken.     0 kPa 0.67 kPa 1.3 kPa 20 kPa [Lactate] 0 h 1.20±0.40a 0.22±0.18a 0.53±0.47a 0.21±0.20  1 h 6.42±0.54b 4.21±0.34b 3.11±0.48b 0.15±0.14  4 h 10.6±0.93c 4.89±0.86b 4.52±0.72b 0.11±0.15 [Ethanol] 0 h 0.61±0.11a 0.74±0.14 0.87±0.20 -  1 h 1.20±0.17b 1.02±0.10 0.77±0.21 -  4 h 2.24±0.20c 1.07±0.10 0.45±0.08 - Table 0-1   53 Chapter 4  Rates of hypoxia induction alter mechanisms of O2 uptake and the critical O2 tension of goldfish      4.1  Summary The rate of hypoxia induction (RHI) is an important but overlooked dimension of environmental hypoxia that may affect an organism’s survival. I hypothesized that, compared with rapid RHI, gradual RHI will afford an organism more time to alter plastic phenotypes associated with O2 uptake and subsequently reduce the critical O2 tension (Pcrit) of O2 uptake rate (ṀO2). I investigated this by determining Pcrit values for goldfish exposed to short (~24 min), typical (~84 min) and long (~480 min) duration Pcrit trials to represent different RHIs. Consistent with my predictions, long duration Pcrit trials yielded significantly lower Pcrit values (1.0-1.4 kPa) than short and typical duration trials, which did not differ (2.6±0.3 and 2.5±0.2 kPa, respectively). Parallel experiments revealed these time-related shifts in Pcrit were associated with changes in aspects of the O2 transport cascade: gill surface areas and haemoglobin-O2 binding affinities were significantly higher in fish exposed to gradual RHIs over 480 min than fish exposed to rapid RHIs over 60 min. My results also revealed that the choice of respirometric technique (i.e., closed versus intermittent) did not affect Pcrit or routine ṀO2, despite the significantly reduced water pH and elevated CO2 and ammonia levels associated with closed-circuit Pcrit trials of ~90 min. Together, these results demonstrate that gradual RHIs result in alterations to physiological parameters that enhance O2 uptake in hypoxic environments. I therefore recommend rapid RHIs (<90 min) when determining an organism’s innate Pcrit so as to avoid the confounding effects of hypoxic acclimation.   54 4.2  Introduction Environmental hypoxia is a common characteristic of many aquatic systems and is becoming increasingly prevalent, severe and long-lasting due to anthropogenic and climate change effects (Friedrich et al., 2014; IPCC, 2014; Schmidtko et al., 2017; Smith et al., 2006). Many studies have examined the physiological impacts of hypoxia exposure on a diverse array of fish species, but these studies have focused almost exclusively on either the severity of the hypoxic exposure (i.e., water PO2, PwO2) or its duration. However, a third dimension of hypoxic exposure, the rate of hypoxia induction (RHI), has received very little attention and is rarely even controlled for (or at least reported) when environmental hypoxia is experimentally induced (Rogers et al., 2016). This is unlike other abiotic variables such as temperature, which are typically altered at consistent rates across studies (e.g., 0.2-0.3°C min-1 for the determination of critical thermal maxima; CTmax) due to the effects these rates have on organismal responses (e.g., temperature tolerance in fishes; Mora and Maya, 2006). Similarly, RHIs may also influence the physiological responses of fishes to hypoxia, particularly time-dependent responses related to environmental O2 extraction. Most fishes possess mechanisms that enhance O2 extraction and delivery to tissues as environmental PO2 is reduced, such as increased haemoglobin (Hb) synthesis (Gracey et al., 2001) and concentration in the blood (Affonso et al., 2002), increased hematocrit (Lai et al., 2006; Turko et al., 2014), increased Hb-O2 binding affinity (Turko et al., 2014), increased ventilation frequency and amplitude (Holeton and Randall, 1967; Itazawa and Takeda, 1978; Tzaneva et al., 2011; Vulesevic and Perry, 2006), and a redistribution of blood supply to critical tissues (Sundin et al., 1995). Some fishes, including the goldfish (Carassius auratus) and other species (Anttila et al., 2015; Borowiec et al., 2015; Crispo and Chapman, 2010; Dhillon et al., 2013; Ong et al., 2007; Turko et al., 2012), also have the ability to dramatically increase lamellar surface area in response to hypoxia exposure through apoptotic reductions to inter-lamellar cell mass (ILCM; Sollid et al., 2003). While these modifications to different parts of the O2 transport cascade function to improve O2 uptake at low PwO2, the time-courses over which these modifications are enacted differ and my potentially impact the critical PO2 (Pcrit) of O2 uptake rate (ṀO2). Pcrit is defined as the PwO2 at which a fish’s ṀO2 transitions from being regulated at some stable level independent of PwO2 (i.e., oxyregulation) to being dependent upon PwO2  55 (i.e., oxyconformation). The stable, oxyregulated ṀO2 typically represents the ṀO2 required to maintain the fish’s standard metabolic rate (SMR) or routine metabolic rate (RMR). Standard metabolic rate is the ṀO2 of an awake, post-absorptive and entirely inactive ectothermic animal (Chabot et al., 2016), while RMR is ṀO2 under the same conditions, but also accounts for the small movements that are typical of fishes under experimental conditions (Chabot et al., 2016). At Pcrit, the fish’s aerobic scope is either at or near zero (for SMR- and RMR-based estimates, respectively), and at PwO2s below Pcrit, the fish’s ability to generate ATP aerobically is limited (Farrell and Richards, 2009). Pcrit therefore reflects a fish’s ability to acquire and use environmental O2 as a function of PwO2, with a lower Pcrit indicating a greater ability to extract O2 to maintain aerobic metabolism in hypoxic environments. A low Pcrit is beneficial because it allows the animal to maintain a routine level of function and activity in hypoxic environments while avoiding a reliance on anaerobic glycolysis and/or metabolic rate depression. Indeed, my results from Chapter 3 reveal that goldfish prioritize their use of aerobic metabolism in hypoxic environments over their exceptional ability to induce metabolic rate depression, which they reserve for anoxic environments. Goldfish also appear to enhance their ability to extract environmental O2 over relatively short time periods in hypoxia, which in theory should result in a lowering of their Pcrit value (Chapter 3). Because this ability is influenced by a suite of O2 extraction mechanisms that are both plastic and time-dependent, I hypothesized that gradual RHIs would allow fish to induce plastic mechanisms that enhance O2 extraction, resulting in lower Pcrit values than those of fish exposed to rapid RHIs. I tested this hypothesis by determining the Pcrit values of goldfish exposed to progressive reductions in PwO2 (from normoxia to near-anoxia; referred to as Pcrit trials) over different durations. Pcrit is typically measured using closed-chamber respirometry, whereby PwO2 is decreased from normoxia to some hypoxic PwO2 by the fish’s own respiration. The rate at which PwO2 decreases therefore depends on the fish’s ṀO2 and the chamber volume, and these trials typically take 60 to 90 minutes to complete (Fry and Hart, 1948; Mandic et al., 2009; Rogers et al., 2016; Sollid et al., 2003; Speers-Roesch et al., 2011). In this study, I used different respirometric techniques to vary the duration of Pcrit trials to determine the effects of rapid (~24 min), typical (~84 min) and gradual (~480 min) RHIs on the Pcrit of goldfish. I supplemented my respirometry experiments with parallel hypoxic exposures of  56 different RHIs to investigate morphological and physiological traits of goldfish that might play causal roles in a time-related shift in Pcrit. These analyses included: gill morphometrics to investigate changes in respiratory gas exchange surface area; whole blood [Hb] to investigate effects on O2 carrying capacity; O2 equilibrium curves (OECs) to investigate effects on Hb-O2 binding affinity; red blood cell (RBC) organic phosphate (NTP; nucleoside triphosphates) concentrations to investigate effects on allosteric modulation of Hb-O2 binding affinity; and lactate accumulation to investigate effects on anaerobic reliance. In addition to these biological assessments, my methods allowed me to address certain technical aspects of respirometry. Specifically, my use of different respirometric techniques allowed the effects of time and technique on the determination of Pcrit to be disentangled, thus addressing a longstanding concern over the use of closed-chamber respirometry and its associated metabolic waste accumulation for the determination of Pcrit (Keys, 1930; Rogers et al., 2016; Snyder et al., 2016; Steffensen, 1989). Finally, I chose goldfish as my study species because they have well-characterized responses to hypoxia exposure (Dhillon et al., 2013; Mitrovic et al., 2009), including a well-resolved Pcrit as determined by closed-chamber respirometry (Chapter 3; Dhillon et al., 2013; Fry and Hart, 1948; Fu et al., 2011), that could aid my analysis of how RHI might influence the underlying physiology of Pcrit.  4.3  Materials and methods 4.3.1  Animals Goldfish (Carassius auratus auratus; 2.87±0.14 g wet mass; N=84; sex unknown) were purchased from a commercial supplier (The Little Fish Company, Surrey, BC, Canada) and held under a 12 h:12 h light:dark cycle in a series of 100 L recirculating systems of well-aerated, dechlorinated, 17°C water at the University of British Columbia (Vancouver, BC, Canada). Stocking density was <0.3 g L-1 and water in the recirculating system was replaced weekly. Fish were fed to satiation daily (Nutrafin Max Goldfish Flakes) except for 24 h before transfer to the experimental apparatus, when feeding ceased. UBC’s Animal Care Committee approved all procedures (protocol A13-0309).   57 4.3.2  Respirometry I exposed goldfish to Pcrit trials of short (~24 min), typical (~84 min, which represents the duration of a typical closed-chamber Pcrit trial) and long (~480 min) durations to represent progressively reduced RHIs. These different RHIs were achieved  using different respirometric techniques (details below), while the respirometer chambers, animal transfer protocol, habituation period, and mean fish mass remained consistent across all trials. Fish were only used once. I used two 32 mL flow-through respirometer chambers made from stainless steel as described in Chapter 2. For each trial, I inserted a fish into the chamber and held them under flow-through conditions for at least 16 h prior to commencing the Pcrit trial. The fish chamber was supplied with flow-through water at a rate of 190 mL h-1 and maintained at 17°C. Inflowing water was drawn from a well-mixed reservoir held at ~26 kPa (manually controlled using compressed N2 and O2) and pumped to the respirometer chamber via a peristaltic pump (Gilson Minipuls 3, Middleton, WI, USA) through a combination of stainless steel tubing and gas-impermeable Tygon peristaltic tubing. The PwO2 of the inflowing water was maintained slightly hyperoxic to ensure that the outflowing water was always at or slightly above normoxic PwO2. Following the habituation period, I conducted my respirometry experiments. For the typical duration Pcrit trials (84±8 min), I used closed-circuit respirometry. To start the trial, the inflowing and outflowing water supply lines were short-circuited so as to create a closed loop, with water recirculating through the chamber by the peristaltic pump at the same rate (190 mL h-1) as during the habituation period to ensure minimal disturbance to the fish and good mixing of the chamber’s water volume. Chamber PwO2 was then allowed to decrease in proportion to the fish’s ṀO2. An O2 optode placed within the chamber (see Chapter 2) continuously measured PwO2, and ṀO2 was calculated according to ṀO2 = (ΔCO2  ΔT-1  V) M-1 where CO2 is O2 content of the water converted to µmol L-1 from PwO2 using the solubility factor of 14.485 µmol L-1 kPa-1 (Boutilier et al., 1984), T is the time period over which the change in CO2 is calculated (5 or 2 min; see below), V is the fish chamber volume (32 mL) plus the volume of the closed-circuit water lines minus the volume displaced by the fish itself, and M is the mass of the fish. The trials were ended when PwO2 reached 0 kPa, at which point  58 the short-circuit was dismantled and flow-through conditions were reestablished to return chamber PwO2 to conditions similar to the habituation period.  For the short duration Pcrit trials (24±2 min), I again used closed-circuit respirometry as described for the typical Pcrit trials. To shorten the trial and hasten the PwO2 decline, I made initial normoxic ṀO2 readings and then manually replaced the entire water volume of the respirometry chamber and its water supply lines over ~5 min with water equilibrated to 5.3 kPa PwO2 using a 60 mL syringe. Water PwO2 was therefore reduced from normoxia to ~5.3 kPa not by the fish’s own ṀO2, but by the active replacement of the water volume. At this point, I attached the water supply lines to the peristaltic tubing, turned the pump back on to 190 mL h-1, and allowed the fish to deplete the closed system’s O2 through its own respiration (typically over a ~20 min period). I chose 5.3 kPa as my replacement PwO2 for two reasons: first, it allowed reliable ṀO2 measurements to be made starting at ~4.8 kPa, which provided enough ṀO2 data points above Pcrit to allow me to construct robust Pcrit traces; second, the amount of time required for the fish to reduce PwO2 from 5.3 kPa to anoxia put the overall duration of these Pcrit trials within my targeted duration of between 20 and 30 min. Although these procedures reduced the overall duration of the Pcrit trial, I must point out that the RHI below 5.3 kPa was similar to that of the typical duration trials. If mechanisms of enhanced O2 extraction are only induced a PwO2 <5.3 kPa, then these two techniques could result in similar Pcrit values. Prior to actively replacing the water volume, I converted the system to closed-circuit and made a series of normoxic ṀO2 readings between 25 and 19 kPa to aid in my calculation of Pcrit (see below). Upon reaching 19 kPa, I converted the system back to flow-through, reestablished a normoxic PwO2 of ~21 kPa, and then commenced the active water volume replacement. For the long duration Pcrit trials, I used three different respirometric techniques to ensure the mean Pcrit values were the result of Pcrit trial duration and not of the respirometric technique per se. These trials varied in average duration from 434 to 562 min depending on the technique used. I chose a time duration of ~480 min because it was significantly longer than the typical trial durations, but in line with some of the longer Pcrit trial durations observed in the literature (see Rogers et al., 2016). For my first technique, I used closed-circuit respirometry where I added a 250 mL water reservoir to reduce the rate at which the  59 fish’s respiration depleted the system’s O2. This reservoir was a glass bottle placed immediately after the peristaltic pump. Water leaving the respirometer chamber was pumped into the reservoir directly over a stir bar that mixed the water volume and prevented any O2 stratification in the bottle. Water flowed out of the reservoir through a stainless steel line that punctured the bottle’s rubber stopper and went directly into the stainless steel line supplying the respirometer chamber. All materials used were gas-impermeable glass or stainless steel. Attaching this reservoir to the closed-circuit system took ~2 min, after which the peristaltic pump was turned back on and the Pcrit trial run according the closed-circuit technique described for the typical duration Pcrit trials. The average duration for these closed-circuit trials was 434±56 min. Second, I used flow-through respirometry where ṀO2 was calculated according to ṀO2 = ((CiO2 - CoO2)  FR) M-1 where CiO2 and CoO2 are respectively O2 content of inflowing and outflowing water converted from PwO2 as described above (I used a single PO2 optode for these measurements), FR is water flow rate (190 mL h-1), and M is the mass of the fish. I held each fish at 26, 16, 5.3, 2.7, 1.3, 0.7 and 0 kPa, each PwO2 in series, in that order and for 1 h, and at each PwO2 I measured ṀO2 at 10, 30 and 60 min (10 min was minimum time required to ensure PwO2 had equilibrated across the respirometer and the upstream and downstream PwO2 measurement chambers). Because the calculated ṀO2 at each PwO2 was nearly identical at each of the three time points, I averaged across the time points and calculated Pcrit from those averaged ṀO2 values for each individual. The average duration for these flow-through trials was 562±19 min, including the time required to reach target PwO2s.  Third, I used a variation on intermittent flow respirometry that combined flow-through and closed-circuit respirometry. I used flow-through conditions to manually reduce PwO2 from normoxia to ~2.8 kPa over ~430 min and then commenced a period of closed-circuit respirometry which took an additional ~15 min. I chose a target PwO2 of 2.8 kPa to start the closed-circuit portion of the trial based upon my earlier short-term Pcrit trials (which used the same combined respirometric technique) that suggested I could reliably determine Pcrit from this PwO2. Upon reaching 2.8 kPa, I converted to the closed-circuit setup and allowed the fish’s respiration to deplete the remaining O2 in the closed system as described previously. This combination of techniques prevented metabolic waste accumulation. As  60 with the rapid RHI Pcrit trials, I used closed-circuit respirometry to make a series of normoxic ṀO2 readings between 25 and 19 kPa prior to the active (but in this case gradual) reduction of PwO2 to aid in my calculation of Pcrit (see below). Upon reaching 19 kPa, I converted the system back to flow-through, reestablished a PwO2 of ~21 kPa, and then commenced the active water volume replacement. The average duration of these combined flow-through and closed-circuit trials was 444±12 min.  4.3.3  Parallel hypoxic exposures for physiological measurements  I ran two separate but identical parallel sets of hypoxic exposures to investigate potentially causal physiological and morphological factors in the time-dependent reduction in Pcrit. These parallel exposures involved manually reducing PwO2 of aquaria from normoxia to anoxia over 60 and 480 min periods to represent rapid and gradual RHIs, respectively. I also ran normoxic control exposures during which PwO2 remained normoxic for 480 min following the habituation period. Each exposure was run in two 10 L aquaria housing four fish each, and each aquarium was fitted with a screen just below the water surface to prevent the fish from accessing the air-water interface. I mimicked the respirometric Pcrit trials described above as closely as possible, with exposures being run at 17°C at the same time of day (each trial commenced at ~9:00am) following a >16 h habituation period, and the aquaria being covered with black plastic so the exposures were done in the dark. Fish from the first set of parallel exposures were sampled to assess gill morphology and haematological parameters, and fish from the second set of parallel exposures were sampled to measure plasma [lactate].  At the end of each exposure, fish were sacrificed by inconspicuously introducing anaesthetic (buffered MS-222, final concentration of 200 mg L-1) to the water. Once the fish reached a surgical plane of anesthesia (~3 min, showed no response to tail pinching), they were individually removed, weighed, and then blood was sampled and gills dissected. To sample blood from the fish in the first set of parallel exposures, the fish’s tail was severed and blood was collected from the caudal preduncle using a 60 µL heparinized capillary tube. Ten µL of blood was pipetted into 1 mL Drabkins reagent for determination of [Hb], 20 µL of blood was mixed with 10 µL of heparinized Cortland’s saline plus 80 µL of 3% perchloric acid for determination of RBC [NTP], and 10 µL of blood was mixed with 5 µL of  61 heparinzed Cortland’s saline for determination of Hb-O2 binding affinity. The entire right gill basket was then removed from the fish and immediately immersed in 1 mL of Karnovsky’s fixative (25% glutaraldehyde, 16% formaldehyde, 0.15 mol L-1 sodium cacodylate, pH 7.4). 24 h later, the gill basket was transferred to 0.15 mol L-1 sodium cacodylate and stored at 4°C until use. This procedure was repeated for all four fish in each tank, and then duplicated for the second tank of four fish yielding N=8 for each treatment. For the second set of parallel exposures, fish were euthanized and blood was collected in the same manner as before, but the plasma was separated from the red blood cells by centrifugation and immediately assayed for plasma [lactate] (see below). The goal of these parallel exposures was to allow me to assess the effects of RHI on morphological and physiological adjustments that may explain differences in Pcrit, but there are differences between the Pcrit trials and the parallel exposures that the reader should be made aware of. The main difference was vessel size (respirometer chamber was 32 mL and exposure aquaria were 10 L), which could have affected the ability of the fish to move throughout the exposure. However, observations of the fish in the 10 L aquaria suggest that goldfish do not increase activity during progressive hypoxia exposure. Furthermore, the parallel exposures were terminated when PwO2 reached 0 kPa. As the samples were taken at this point, the haematology and gill morphology measurements were not taken precisely at the point where I observed differences in Pcrit, and this could affect my ability to relate the two studies. However, the fish used for the haematology and gill morphology analyses were only exposed to an additional ~7 to ~15 min of progressively deepening hypoxia (for rapid and gradual RHI, respectively) beyond what they had induced by the time Pcrit had been reached. Thus I do not believe these relatively minor differences in time would affect my ability to directly relate these components of my study.  4.3.4  Gill morphometrics  Gill samples were randomly assigned an alphanumeric code by an independent party so analysis could be performed blindly. The second gill arch of each gill basket was isolated and its anterior hemibranch imaged using light microscopy (Olympus Stereomicroscope SZX10; 6.3× magnification, 10× zoom; image capture using cellSens Software). The images were used in combination with ImageJ v2.0.0 software to measure filament length and  62 number, and lamellar height (distance from base to the distal edge of the lamellae), length (distance lamellae runs along the filament) and frequency (number of lamellae per unit distance of filament). I made the lamellar measurements by dividing the length of the gill arch into five sections, then isolating a filament from each of these sections. Each filament was imaged from the top and the side, providing clear views of the height and length of its lamellae that I later measured. Specifically, I measured the height, length and width of three lamellae per filament (one from the filament’s base, one from its middle and one from its tip), as well as the width of inter-lamellar channels in these three regions. I then estimated each filament’s lamellar frequency (lamellae µm-1) by dividing filament length by the sum of that filament’s average channel and lamellar widths. Total lamellar surface area for each fish was then calculated according to,  Total LSA = FSA · Ffreq · 16 where FSA is the mean lamellar surface area of the five analyzed filaments, Ffreq is the number of filaments per gill arch, and 16 is the product of 2 hemibranchs per gill arch, 4 gill arches per gill basket, and 2 gill baskets per individual fish (according to Wegner, 2011).  4.3.5  Blood analyses  Haemoglobin-O2 binding affinity was determined within 60 min of blood sampling by constructing an OEC using the thin film spectrophotometric technique (Lilly et al., 2013) and a 96 well microplate spectrophotometer (Molecular Devices, Sunnyvale, CA, USA). A Wostoff gas mixing pump (H. Wösthoff Messtechnik GmbH, Bochum, Germany) mixed compressed O2 and N2 to each of nine PO2s between 0 and 21 kPa PO2, always starting with 0 kPa and working toward 21 kPa, and each PO2 was maintained for 20 min during which Hb-O2 saturation was determined spectrophotometrically. A sigmoidal OEC was fit through the % Hb-O2 saturation versus PO2 data for each fish, and Hb P50 (the PO2 at which Hb is 50% saturated with O2) was determined using SigmaStat 11.0. I measured whole blood [Hb] spectrophotometrically at 17°C and 540 nm after conversion to cyanomethemoglobin using Drabkin’s reagent (Sigma-Aldrch). The measurements were made using a Shimadzu UV-160 spectrophotometer and a millimolar extinction coeffiecient of 11.   63  I measured red blood cell [NTP] spectrophotometrically at 17°C using the GAPDH- and PGK-catalyzed reactions converting glycerate 3-phosphate to glyceraldehyde 3-phosphate, where the oxidation of NADH to NAD+ was measured at 340 nm (Bergmeyer et al., 1983). The measurements were made using a 96 well microplate spectrophotometer (Molecular Devices, Sunnyvale, CA, USA).   Finally, I measured plasma [lactate] spectrophotometrically at 17°C using the LDH-catalyzed reaction converting lactate to pyruvate, where the reduction of NAD+ to NADH was measured at 340 nm (Bergmeyer, 1983). The measurements were made using a 96 well microplate spectrophotometer (Molecular Devices, Sunnyvale, CA, USA).   4.3.6  CO2 and nitrogenous end-product measurements I ran a separate set of closed-circuit Pcrit trials (91±10 min) to measure accumulated CO2 and nitrogenous end-products (NH3 + NH4+). For each of four fish, I took water samples from the respirometer chamber at three time points: at the start of the habituation period, at the end of a 16 h habituation period immediately prior to starting the Pcrit trial, and at the end of the Pcrit trial as soon as the respirometer’s PwO2 reached 0 kPa. PwCO2 was determined using the Henderson-Hasselbalch equation and measurements of total CO2 content in the water (CO2 + HCO3-; Corning 965 Carbon Dioxide Analyzer, Corning, NY, USA) and pH (probe: SaS gK2401C, Radiometer analytical, France; meter: VWR Symphony SB70P, VWR, Radnor, PA, USA). Total ammonia (NH3 + NH4+) was measured using an API ammonia test kit, and unionized ammonia (NH3) was calculated from this value in combination with water pH and temperature (17°C).  4.3.7  Pcrit calculation Pcrit is defined as the PwO2 at which an organism’s stable ṀO2 transitions from being independent of, to being dependent upon, PwO2. There are different methods to calculate Pcrit, but analyses performed by Mueller and Seymour, (2011) suggest that most of the methods used yield comparable values. I therefore decided to use a variation on a two-segment linear regression model (details below) to identify Pcrit as the PwO2 at which the two linear trend lines (one representing the PwO2 range of oxyregulation, the other of oxyconformation) intersect on a graph plotting ṀO2 as a function of PwO2 (BASIC program of Yeager and  64 Ultsch, 1989). This method is employed widely throughout the literature (see Rogers et al., 2016) and has been used by myself and by others for goldfish (Chapter 3; Fu et al., 2011; Dhillon et al., 2013).  I calculated ṀO2 values by measuring the change in PwO2 over sequential time intervals, 5 min between 25 and 5.3 kPa and 2 min between 5.3 and 0 kPa. To standardize my estimates of a stable, oxyregulated ṀO2, I used the mean of each fish’s calculated ṀO2 values between 21 and 18.7 kPa PwO2. This represented a normoxic routine ṀO2 that was likely close to standard ṀO2 as a result of it being made following a ≥18 h habituation period. I then determined Pcrit as the intersection of this horizontal line with a linear regression through the ṀO2 values that were >15% below the mean routine ṀO2 value. This technique was carried out according to McBryan et al. (2016).  4.3.8  Data analysis and statistics I compared all average values of Pcrit, normoxic ṀO2, blood properties, gill morphometrics and accumulated PwCO2 and nitrogenous end-products using one-way, two-tailed ANOVAs with a critical α=0.05 (repeated measures for the water pH, PwCO2 and nitrogenous end-products comparisons). Post hoc Tukey tests were used to test for differences between treatment groups. Any data set that did not meet the assumptions of normality or equal variance was log-transformed prior to analysis. All analyses were performed using SigmaStat 11.0. Finally, values reported in the text are presented as means±sem.  4.4  Results 4.4.1  Respirometry  Long duration Pcrit trials resulted in significantly lower Pcrit values than typical and short duration Pcrit trials (Fig. 4-1; Fig. 4-2A, ANOVA P<0.001). Pcrit values determined by short and typical trial durations did not differ from one another, nor did the Pcrit values determined by the three respirometric techniques used for the long duration trials (Fig. 4-2A). Each of the five respirometric techniques yielded statistically similar normoxic ṀO2 values (Fig. 4-2B; ANOVA P=0.276).   65 4.4.2  Effect of RHI on gill morphology  RHI had a significant effect on the mass-specific lamellar surface areas of goldfish (Fig. 4-3, ANOVA P=0.004), whereby fish exposed to gradual RHIs had significantly larger lamellar surface areas than fish exposed to rapid RHIs and normoxic controls, which did not differ.  4.4.3  Effect of RHI on Hb-O2 affinity, [Hb] and RBC [NTP]  RHI had a significant effect on Hb-O2 binding affinity (Fig. 4-4A,B; ANOVA P=0.007). Rapid RHI significantly reduced Hb-O2 binding affinity relative to normoxic control values as determined by Hb P50, while the gradual RHI had no significant effect. RHI did not affect whole blood [Hb] (Fig. 4-5A; ANOVA P=0.334), but it did affect RBC [NTP] (Fig. 4-5B; ANOVA P=0.001), whereby gradual RHIs resulted in significantly lower RBC [NTP] than rapid RHIs and normoxic control exposures.   4.4.4  Effect of RHI on plasma lactate  Goldfish exposed to rapid and gradual RHIs both accumulated similar concentrations of plasma lactate to a level significantly higher than that observed in normoxic control fish (Fig. 4-5C; ANOVA P=0.001).    4.4.5  Metabolic end-product accumulation  Compared with the start of the habituation period, typical duration closed-circuit Pcrit trials (91±10 min) resulted in a ~2-fold increase in respirometer chamber PwCO2 over the course of the 16 h flow-through habituation period, and a ~13-fold increase over the course of the Pcrit trial itself (Fig. 4-6A, ANOVA P<0.001). Water pH was concomitantly reduced from 7.61 to 6.93 over the course of the Pcrit trial (Fig. 4-6B; ANOVA P<0.001). The concentration of total ammonia (NH3 + NH4+) in the chamber also increased (Fig. 4-6C, ANOVA P<0.001). Unionized ammonia (NH3) accumulated in a different way due to pH changes of the water, with [NH3] increasing ~3-fold over the 16 h habituation period, then falling to an intermediate value by the end of the Pcrit trial habituation period (Fig. 4-6D, ANOVA P<0.001).   66 4.5  Discussion I hypothesized that gradual RHIs would allow goldfish to induce time-dependent plastic phenotypes that enhance O2 uptake. This hypothesis predicted that the Pcrit of goldfish exposed to long-duration Pcrit trials would be lower than those of goldfish exposed to short- or typical-duration Pcrit trials, and my results agree with these predictions regardless of the respirometric technique used. Furthermore, my results suggest that this time-dependent shift in Pcrit is the result of a greater lamellar surface area and a higher Hb-O2 affinity in gradual RHI-exposed fish, leading to an enhanced ability to extract O2 from hypoxic water. Taken together, these results suggest that time (more precisely, RHI) is a significant determinant of Pcrit in goldfish. The vast majority of Pcrit measurements are made using closed-chamber respirometry over the course of 60 to 90 min (Rogers et al., 2016). Here, my representative closed-circuit Pcrit trials lasted ~84 min and resulted in a Pcrit of 2.5±0.2 kPa (Fig. 4-2A). My values are in general agreement with the values previously reported for goldfish [~3.6 kPa (Fry and Hart, 1948); 3.0 kPa (Fu et al., 2011); 3.3 kPa (Dhillon et al., 2013); 3.0 kPa (Chapter 3)], though slightly lower owing to a possible combination of experimental temperature differences and the fact that my study used closed-circuit respirometry as opposed to static closed-chamber respirometry. Reducing the trial duration to ~24 min did not affect Pcrit (Fig. 4-2A), which may not be surprising considering the RHI below a PwO2 of 5.3 kPa was similar between the short and typical duration Pcrit trials (see Materials & Methods for details). However, my results clearly indicate that increasing the trial duration from ~84 min (i.e., reducing its RHI) to ~480 min resulted in significantly lower Pcrit values (Fig. 4-2A). The reasons for this variation could be related to time, technique, or some combination of the two, and I will explore these possibilities below.  4.5.1  Effects of time on the physiology of O2 uptake Goldfish exposed to long duration Pcrit trials displayed a greater ability to extract O2 from hypoxic water than goldfish exposed to short and typical duration Pcrit trials. In line with this, the physiological changes I observed in the fish from the parallel hypoxic exposures were consistent with the gradual RHI fish having improved O2 extraction abilities compared with those of the rapid RHI fish.   67 Goldfish exposed to gradual RHIs had significantly larger lamellar surface areas than those of normoxic controls and goldfish exposed to rapid RHIs, which did not differ. Hypoxia-induced gill remodeling was first observed in goldfish and the closely-related crucian carp (Carassius carassius) 13 years ago (Sollid et al., 2003; Sollid et al., 2005) and in numerous fish species since [e.g., mangrove killifish, Kryptolebias marmoratus (Ong et al., 2007; Turko et al., 2012); African cichlids (Crispo and Chapman, 2010); various carp species (Dhillon et al., 2013); Atlantic salmon Salmo salar (Anttila et al., 2015); Atlantic killifish Fundulus heteroclitus (Borowiec et al., 2015)]. Dhillon et al. (2013) observed a ~2-fold increase in the lamellar surface area of goldfish following 8 h acclimation to a constant PwO2 of 0.7 kPa, but to my knowledge my study is the first time gills have been shown to remodel over such short time scales under progressively decreasing PwO2. Increases to lamellar surface area are typically the result of apoptotic reductions to the ILCM (Sollid et al., 2003). ILCM reductions per se also enhance the gill’s diffusion capacity (Bindon et al., 1994; Greco et al., 1995) and contribute to a reduced Pcrit in crucian carp (Sollid et al., 2003) and Atlantic killifish (Borowiec et al., 2015). However, in a study that examined (among other things) O2 diffusion across the gills of two groups of goldfish with temperature-induced differences in gill surface area, the authors found that the differences in gill surface area had no effect on arterial PO2 when acutely exposed to hypoxia (Tzaneva et al. 2011). While this seems to run counter to what Fick’s first diffusion law would predict, the authors speculated that the goldfish that started hypoxia exposure with a smaller gill surface area may have been rapidly remodeling their gills to increase lamellar surface area over the course of the acute hypoxia exposure. My gill morphometric results lend empirical support to this speculation. Goldfish exposed to gradual RHIs almost halved their Hb P50 values compared with goldfish exposed to rapid RHIs, and these values roughly correlated with Pcrit: gradual RHI fish had an average Hb P50 of 1.6 kPa and an average Pcrit of 1.2 kPa, while the rapid RHI fish had 2.4 and 2.5 kPa (for typical duration trials), respectively. This is consistent with the positive correlation of Hb P50 and Pcrit shown across intertidal sculpin species by Mandic et al. (2009), and with the idea that the O2 binding affinity of Hb is the main factor setting the environmental PO2 when integrated physiological function begins to be lost (Farrell and Richards, 2009). But while the P50 values for the gradual RHI fish were lower than those for the rapid RHI fish, it was the rapid RHI fish that had significantly higher P50 values than both  68 the controls and gradual RHI fish. This suggests that rapid hypoxia induction reduces Hb-O2 affinity, but gradual induction does not. The underlying mechanism(s) might involve RBC [NTP] and/or protons. Nucleoside triphosphates (ATP and GTP, collectively NTPs) reduce Hb-O2 binding affinity by binding to sites on the Hb tetramer that stabilize its deoxygenated conformation and consequently increase the P50 (Jensen et al., 1998; Wood and Johansen, 1972). The significantly lower [NTP]s of my gradual RHI fish (Fig. 4-5B) at least partly explain their lower Hb P50 values compared with those of the rapid RHI fish (Fig. 4-4), but the similar [NTP]s in the rapid RHI and normoxic control fish exclude this mechanism as the cause of the rapid RHI fish’s elevated Hb P50 values. Another possibility is the goldfish’s proton-sensitive Root effect Hbs (Rodewald and Braunitzer, 1984), whereby the higher P50 values in goldfish experiencing a rapid RHI might be the result of a more rapid accumulation of glycolytically-derived protons compared with goldfish experiencing a gradual RHI (as indicated by the similar plasma lactate accumulation following both the rapid and gradual RHI; Fig. 4-5C). However, if goldfish possess an adrenergically-activated RBC Na+/H+ exchanger like the closely related common carp (Cyprinus carpio) (Salama and Nikinmaa, 1988; Salama and Nikinmaa, 1989), then a reduced RBC pH may not explain the increased Hb P50 seen in my rapid RHI fish because such an exchanger would regulate RBC pH under hypoxic conditions. Regardless of the causal mechanism(s), the different Hb-O2 binding affinities of the rapid and gradual RHI fish are likely to at least partly explain their different Pcrit values. Other factors besides lamellar surface area and Hb-O2 binding affinity could potentially contribute to a time-dependent shift in Pcrit, and one such factor may be reduced blood perfusion of the gills resulting in reduced O2 uptake from the water. For example, common carp exposed to a rapid and progressive induction (<1 h) of severe hypoxia (to ~0.7 kPa) showed a 5-fold reduction in cardiac output with a nadir that coincided with the time and PwO2 of the fish’s Pcrit (Stecyk and Farrell, 2002). Similarly, dogfish (Scyliorhinus canicula) exposed to a rapid and progressive induction (rate/time domain not reported beyond “rapid”) of hypoxia (to 4.0 kPa) showed an initial intense bradycardia that was not observed in dogfish exposed to a gradual induction of hypoxia, despite both groups stabilizing at the same heart rate following 30 min (Butler and Taylor, 1971). Finally, in two  69 elasmobranch species that differ in hypoxia tolerance [epaulette shark (Hemiscyllium ocellatum) and shovelnose ray (Aptychotrema rostrata)], the species-specific Pcrit values coincided with the PwO2s at which their cardiac output was reduced compared with normoxia-acclimated fish (Speers-Roesch et al., 2012a, 2012b). These authors also pooled literature values for various fish species and found a significant positive correlation between Pcrit and the PwO2s at the onset of hypoxic bradycardia (Speers-Roesch et al., 2012b). But while these authors highlight the dependence of cardiac function on Pcrit of ṀO2, here I speculate the opposite; that is, Pcrit may partially depend on cardiac function.  4.5.2  Respirometric technique and waste accumulation Respirometric techniques can be broadly categorized as closed (closed-circuit or static closed-chamber), flow-through, or intermittent flow. Though none of these techniques are ideal for all experimental questions, intermittent flow respirometry is generally regarded as superior because it avoids the potential accumulation of metabolic end-products that can occur in closed respirometry and it has greater temporal resolution compared with flow-through respirometry (reviewed by Clark et al., 2013; Steffensen, 1989; Svendsen et al., 2016). It has been suggested that the choice of respirometric technique used to determine Pcrit may influence the results, and indeed Pcrit in shiner perch (Cymatogaster aggregata) shifted from ~9.9 kPa to ~6.1 kPa when using closed-chamber versus intermittent flow respirometry, respectively (Snyder et al., 2016). The authors attribute this to technique, but they also discuss the possibility that duration of the Pcrit trials (~1 h for closed-chamber, ~5 h for intermittent flow) may play a role (Snyder et al., 2016). In the present study, I used modified versions of all three respirometric techniques for my long duration Pcrit trials, and despite technique specific-differences and challenges [e.g., flow-through trials demanded a step-wise reduction in PwO2; closed-circuit trials resulted in higher ṀO2 values in the mid-PwO2 range (Fig. 4-1)], each technique yielded nearly identical Pcrit values, which were all lower than the typical or short duration Pcrit trials. This suggests that the different Pcrit values observed between my short and typical duration Pcrit trials and those of the long duration trials are the result of RHI rather than technique, and this may also be the case with the results of Snyder et al. (2016). However, the fact remains that closed respirometry leads to end-product accumulation, which could still theoretically influence Pcrit.  70 The buildup of metabolic wastes (CO2 and ammonia) in closed-circuit respirometry has for many years been suggested to affect ṀO2 and consequently Pcrit (Keys, 1930; Snyder et al., 2016; Steffensen, 1989). However, to my knowledge, ours are the first reported empirical measurements of these waste products following a closed-circuit Pcrit trial. Chamber PwCO2 was determined to be 0.80±0.02 kPa following typical RHI Pcrit trials (Fig. 4-6A), and this value agreed with my original estimate of 0.71 kPa based on water volume, temperature, salinity and starting O2 content, as well as fish size, average ṀO2, an RQ of 1 and a CO2 solubility coefficient of 0.4224 mmol L-1 kPa-1 (from Boutilier et al., 1984). While this PwCO2 is certainly hypercapnic, it is much lower than the 5.3+ kPa typically tolerated by fishes (Baker et al., 2009; Baker et al., 2015; Brauner et al., 2004; Grøttum and Sigholt, 1996; Hayashi et al., 2004; McKenzie et al., 2002; Takeda and Itazawa, 1983). Furthermore, the three studies I know of that have measured Pcrit as a function of hypercapnia present conflicting, but possibly PwCO2-dependent, results. European eels (Anguilla anguilla) have shown both an increase in Pcrit in hypercapnia (Cruz-Neto and Steffensen, 1997; PwCO2 of 2.7 and 4.0 kPa) and no effect (McKenzie et al., 2003; PwCO2 of 1.3 kPa), while Atlantic killifish and spot (Leiostomus xanthurus) have both shown no effect (Cochran and Burnett, 1996; max PwCO2 of 1.6 kPa). It therefore appears that Pcrit in these species is impacted only by relatively high PwCO2s. If this applies to goldfish too, then a PwCO2 of 0.80 kPa is unlikely to have a significant effect on Pcrit. Finally, the most supportive piece of evidence for mild hypercapnia’s limited effect on Pcrit is the fact that my study’s closed-circuit trials (where CO2 accumulated) and combined flow-through/closed-circuit trials (where CO2 did not accumulate) resulted in nearly identical Pcrit values within the long duration Pcrit trials (Fig. 4-2A). I also measured ammonia accumulation following typical RHI closed-circuit Pcrit trials. While the average total ammonia (NH3 + NH4+) concentration increased from 1.4 to 47 µmol L-1 following the trial, the average concentration of NH3 was not statistically different than the starting concentrations when water pH was taken into account (Fig. 4-6D). It is therefore unlikely that ammonia accumulation affected Pcrit.   71 4.5.3  Implications and recommendations The present study demonstrates that RHI influences the physiological responses of goldfish to hypoxia, which consequently influences their Pcrit values. This influence can be substantial, and in my hands, reducing RHI with Pcrit trials of ~480 min instead of ~84 min reduced the Pcrit of goldfish by up to 2-fold. If this is broadly applicable to other species, then it could undermine recent Pcrit-based modeling efforts that aim to understand how fishes will respond to an increasingly hypoxic world (Deutsch et al., 2015). This is in addition to already existing concerns regarding Pcrit as an overall measure of hypoxia tolerance (Claireaux and Chabot, 2016; Rogers et al., 2016; Salin et al., 2015; Speers-Roesch et al., 2013). The exceptional hypoxia tolerance of goldfish and the mechanisms that underlie it, particularly a highly plastic gill structure, may draw into question the general applicability of my results to other fish species. However, hypoxia-induced gill remodeling has now been described in at least eight species (see above for details), suggesting it may be a more widespread hypoxia-induced response of fishes than previously thought. In addition, and perhaps more importantly, many fish species (if not most or all) have the capacity to alter haematological parameters such as haematocrit and Hb-O2 binding affinity over the course of hypoxia exposure (Wells, 2009), thus possibly affecting O2 uptake and Pcrit. The rates at which these physiological alterations are induced may vary among species, perhaps as a function of the RHI of their natural hypoxic habitats, but all species will likely acclimate to some extent. Thus, it is reasonable to suggest that RHI is an important factor to consider when examining the hypoxic responses of all fish species, but confirmation of this will require more work. If my results are generally applicable across species, then it is important to consider the implications of RHI on the determination of Pcrit. Because longer duration Pcrit trials allow for some degree of acclimation that may consequently reduce Pcrit, shorter duration Pcrit trials likely best represent the innate abilities of a hypoxia-exposed fish to extract and use O2 at the time of analysis. Thus, similar to the standardized rate of temperature change used when determining a fish’s CTmax, an RHI should be chosen that is fast enough to avoid acclimation so as to capture the innate Pcrit of the organism. Because my short (~24 min) and typical (~84 min) duration Pcrit trials yielded nearly identical Pcrit values for goldfish, Pcrit  72 trials of <90 min are advised. In addition, my results also suggest that closed-chamber respirometry is appropriate for the determination of Pcrit.    4.5.4  Conclusions These results demonstrate that RHI significantly alters the Pcrit of goldfish, whereby long duration Pcrit trials (i.e., gradual RHIs) yield lower Pcrit values than short duration Pcrit trials (i.e., rapid RHIs). These reduced Pcrit values are caused by time-dependent effects on mechanisms that enhance environmental O2 extraction, including gill morphology and Hb-O2 binding affinity. Thus, to determine the innate Pcrit of a fish at a moment in time, I recommend shorter duration Pcrit trials carried out using closed-chamber/circuit respirometry. This would avoid any effects of acclimation and remain consistent with the methods used for the majority of Pcrit values in the literature.                    73 Figure 10       Figure 4-1. The effect of water PO2 on the ṀO2 of goldfish exposed to rapid, typical and gradual rates of hypoxic induction. The different rates were achieved through different Pcrit trial durations, and each of the five Pcrit trails used a different respirometric technique to achieve its respective duration. (A) was achieved using a variation on intermittent flow respirometry (N=5; average duration 24±2 min), (B) was achieved using closed-circuit respirometry (N=6; average duration 84±8 min), (C) was achieved using flow-through respirometry (N=6; average duration 562±19 min), (D) was achieved using closed-circuit respirometry with an additional water volume (N=6; average duration 434±56 min), and (E) was achieved using a variation on intermittent flow respirometry (N=4; average duration 444±12 min.0 5 10 15 20 25 3002468Water PO2 (kPa)MO2 (µmol h-1 g-1)0 5 10 15 20 25 3002468MO2 (µmol h-1 g-1)0 5 10 15 20 25 30024680 5 10 15 20 25 3002468100 5 10 15 20 25 3002468Water PO2 (kPa)MO2 (µmol h-1 g-1)Rapid Typical Gradual Gradual Gradual A B C D E  74  Figure                        11  Figure 4-2. The effect of Pcrit trial duration on the average Pcrit and normoxic ṀO2 values of goldfish. (A) shows the average Pcrit values of the individual fish comprising each set of respirometry experiments. (B) shows the average normoxic ṀO2 values of the individual fish comprising each set of respirometry experiments while those fish were exposed to normoxic PwO2 (18 to 26 kPa). Details on the respirometric techniques are included in the Materials and Methods, but briefly, “short” used combined flow-through/closed-circuit intermittent flow respirometry (N=5; 24±2.2 min), “typical” used closed-circuit respirometry (N=6; 84±8 min), and “long” from left to right used closed-circuit respirometry (N=6; 434±56 min), combined flow-through/closed-circuit intermittent flow respirometry (N=4; 444±12 min), and flow-through respirometry (N=6; 562±19 min). Error bars are s.e.m., and bars that share a letter are not significantly different [1-way ANOVA, P<0.001 for (A), and P=0.276 for (B)].0123Short Typicalaab bPcrit trial durationbLongP crit (kPa PO2)0246Pcrit trial durationShort Typical LongNormoxic MO2 (µmol h-1 g-1)a a a a a A B  75 Figure 12                         Figure 4-3. The effect of the rate of hypoxia induction on the mass specific lamellar surface area of goldfish. Mass-specific lamellar surface areas of goldfish exposed to rapid and gradual RHIs (normoxia-to-anoxia in 60 and 480 min, respectively), and normoxic controls (N=8 for each; 1-way ANOVA, P=0.004). Error bars are s.e.m., and bars that share a letter are not significantly different.  0100200300NormoxiccontrolRapid RHIGradual RHIaabLamellar surface area (mm2  g-1 ) 76 Figure 13  Figure 4-4. The effect of the rate of hypoxia induction on the O2 equilibrium curve and Hb P50 value of goldfish. . (A) shows the O2 equilibrium curves (OECs) for the extracted whole blood of goldfish exposed rapid and gradual RHIs (normoxia-to-anoxia in 60 and 480 min, respectively), and normoxic controls. The blood was collected from the fish immediately upon PwO2 reaching ~0 kPa (for rapid and gradual RHIs), and the spectrophotometric determination of Hb-O2 saturation was begun as soon thereafter as possible (<1 h). Each OEC is a trendline through the data points of eight blood samples exposed to nine PwO2s between 0 and 21 kPa PO2. Grey horizontal dashed line highlights the 50% Hb-O2 saturation point. (B) shows the average Hb P50 values (PO2 at which Hb is 50% saturated with O2) for each treatment group (1-way ANOVA, P=0.007). Error bars are s.e.m., and bars that share a letter are not significantly different. 0 5 10 15 200255075100Normoxic controlRapid RHIGradual RHIPO2 (kPa)Hb-O2 saturation (%)0123NormoxiccontrolRapid RHIGradual RHIabaHb P 50 (kPa)A B  77 Figure 14   Figure 4-5. The effect of the rate of hypoxia induction on blood parameters of goldfish. (A) shows average values for whole blood [Hb] measured spectrophotometrically using Drabkins reagent (N=6-8; 1-way ANOVA, P=0.334). (B) shows average values for RBC [NTP] (N=6-8; 1-way ANOVA, P=0.001). (C) shows average values for plasma [lactate] (N=6-8; 1-way ANOVA, P=0.001). Error bars are s.e.m., and bars that share a letter are not significantly different.  0.00.30.60.91.2Whole blood [Hb] (mmol L-1)NormoxiccontrolRapid RHIGradual RHI012345aabRBC [NTP] (mmol L-1)NormoxiccontrolRapid RHIGradual RHI02468ab bPlasma [lactate] (mmol L- 1)NormoxiccontrolRapid RHIGradual RHI A B C  78 Figure 15   Figure 4-6. The effects of closed-circuit respirometry on water chemistry and the buildup of metabolic wastes. (A) shows chamber PwCO2, measured as total CO2 and converted to PwCO2 using water pH and the Henderson-Hasselbalch equation (N=4; 1-way ANOVA, P<0.001). (B) shows water pH (N=4; 1-way ANOVA, P<0.001). (C) shows total ammonia concentration (NH3 + NH4+; N=4; 1-way ANOVA, P<0.001) and (D) shows unionized ammonia concentration, calculated using water pH and temperature (N=4; 1-way ANOVA, P<0.001). Error bars are s.e.m., and bars that share a letter are not significantly different.   0.00.20.40.60.81.0Pcrit trialstartPcrit trialendabcHabituationperiodChamber PCO2 (kPa)6.57.07.58.08.5Pcrit trialstartPcrit trialendHabituationperiodabcWater pHA B D 0204060Pcrit trialstartPcrit trialendabcHabituationperiodChamber [total ammonia] (µmol L-1)C 0.00.20.40.6Pcrit trialstartPcrit trialendHabituationperiodababChamber [ammonia] (µmol L-1)D  79 Chapter 5  Metabolic depression and the rapid evolution of hypoxia tolerance in threespine sticklebacks, Gasterosteus aculeatus      5.1  Summary  Anthropogenic increases in global temperature and agricultural runoff are increasing the prevalence of aquatic hypoxia throughout the world. If aquatic animals like fishes are to continue living where they are, they will need to enhance their hypoxia tolerances over relatively short timescales. I investigated the potential for a relatively rapid evolution of hypoxia tolerance using two isolated (for <11,000 years) populations of threespine sticklebacks, one from a lake that experiences long-term hypoxia (Alta Lake, British Columbia) and one from a lake that does not (Trout Lake, British Columbia). Loss-of-equilibrium experiments revealed the Alta Lake sticklebacks are significantly more tolerant of hypoxia than the Trout Lake sticklebacks, and the enhanced tolerance of Alta Lake sticklebacks is associated with an ability to employ metabolic rate depression (MRD) in hypoxia, something the Trout Lake fish do not do. The two populations do not differ in their capacities for O2 extraction or anaerobic metabolism. These results reveal that significant intraspecific variation in hypoxia tolerance can evolve over relatively short timescales, as can MRD, a complex biochemical response that may be favoured in long-term hypoxic environments. Sticklebacks represent a powerful model to investigate the mechanisms and evolution of hypoxia tolerance in an increasingly hypoxic world.   80 5.2  Introduction The world’s aquatic environments are becoming increasingly hypoxic as a result of elevated water temperatures and increased agricultural runoff (IPCC, 2014; Schmidtko et al., 2017; Smith et al., 2006). Because animals rely on O2 to supply sufficient ATP to meet their metabolic demands, environmental hypoxia threatens the animal’s ability to maintain energy balance, homeostasis and, consequently, life. This threat is compounded in ectothermic animals like fishes by the concomitant increase in metabolic demand that necessarily accompanies rising temperatures. However, many fishes have evolved mechanisms of hypoxia tolerance as a result of adapting to environments that already experience hypoxia (Hochachka and Lutz, 2001), and we can look to the strategies of these species to better understand what might facilitate or hinder an animal’s ability to adapt to a changing world.  Like all animals, fishes have three metabolic mechanisms for balancing energy supply and demand in hypoxia: aerobic metabolism, anaerobic metabolism (primarily in the form of anaerobic glycolysis) and metabolic rate depression (MRD). These mechanisms can be combined in different ways to maintain cellular energy balance, and because hypoxic environments vary in their hypoxic severity, duration and rate of induction, the combination a species uses to survive hypoxia likely stems from the unique hypoxic environment to which that species is adapted.  Fish species that are native to environments that regularly become hypoxic tend to be more tolerant of hypoxia than fish species that are native to less hypoxic environments (Chapman et al., 2002; Mandic et al., 2009b). This relationship may also exist among different populations of the same species as it does for other stressors like temperature (e.g., Fangue et al., 2006), and in fact two subspecies of Atlantic killifish (Fundulus heteroclitus) have recently been shown to differ in hypoxia tolerance (McBryan et al., 2016). If intraspecific variation in hypoxia tolerance exists, defining the underlying mechanisms in the context of the different populations’ natural environments could reveal much about how the hypoxic environment shapes the hypoxic survival strategy. To address this, I collected threespine sticklebacks (Gasterosteus aculeatus) from isolated populations native to two British Columbia lakes: Alta Lake, which experiences long-term hypoxic bouts due to overwinter freezing (Dunnington et al., 2016; Jacques Whitford/AXYS, 2007), and Trout Lake, which does not experience long-term hypoxia. I  81 predicted that Alta Lake sticklebacks would be more hypoxia tolerant than Trout Lake sticklebacks, and that this difference would result from an increased reliance on MRD in the Alta Lake fish. I predicted MRD to be the causal mechanism because of its effectiveness at maintaining cellular energy balance during long, deep hypoxic bouts (Hochachka et al., 1996) and its prevalence of use among ectothermic vertebrate species that inhabit similar winter environments (Chapter 3; Jackson, 1968; Johansson et al., 1995). To test this hypothesis, I used time-to-loss of equilibrium (LOE) experiments to assess the hypoxia tolerances of the two stickleback populations. I also used calorespirometry to simultaneously measure the fish’s use of aerobic metabolism and MRD at severely hypoxic O2 tensions (PwO2), and then ran parallel hypoxic exposures to measure the anaerobic contributions to metabolic rate. I chose threespine sticklebacks as my model species because they are a well-studied example of local ecological adaptation, having invaded coastal British Columbia freshwater habitats ~11 000 years ago following the retreat of the Pleistocene glaciers (Bell and Foster, 1994; Jones et al., 2012; Mathews and Fyles, 1970; McPhail, 1994). As the land rebounded, the populations became isolated from one another in separate freshwater systems and consequently evolved, both in parallel and divergently, at physiological, morphological and behavioural levels (Boughman, 2001; Colosimo et al., 2005; Dalziel et al., 2012b; DeFaveri and Merila, 2014; Jones et al., 2012). In my study, I exploited the different O2 regimes of Alta Lake and Trout Lake to investigate variation in hypoxia tolerance and the mechanisms that underlie it.  5.3  Materials and methods 5.3.1  Lakes Alta Lake (Whistler, BC, Canada; 50°11’42”N 122°98’11”W) and Trout Lake (Sechelt, BC, Canada; 49°50’82”N 123°87’64”W) are similar oligotrophic water bodies (Table 5-1) that differ in elevation and subsequently the number of days each lake is covered in ice each year. Alta Lake is surface-frozen for 128±3.64 d y-1, while Trout Lake does not freeze.   82 5.3.2  Field collection and husbandry  I used minnow traps to collect sticklebacks from both lakes. Traps were placed on the lake bottom at 1 to 2 m depth and 3 to 5 m offshore, far from tributaries. Fish for the calorespirometry experiments were collected in October 2015, and fish for the parallel hypoxic exposures and LOE trials were collected in May 2016. I transported the fish to The University of British Columbia and held them in 100 L recirculating aquaria at a density of <0.3 g L-1 and under 12h:12h light:dark. Dechlorinated Vancouver tap water was aerated and held at 17°C, and 25% water changes were carried out every two weeks. Fish were fed bloodworms (Hikari Bio-Pure) daily to satiation and were held for 3 weeks prior to running experiments to allow adjustment to laboratory conditions of diet and light cycle. I held the water and laboratory temperature at 17°C because I had previously determined this to be the best temperature at which to run calorespirometry experiments (see Chapter 2).   5.3.3  Hypoxic exposures, calorespirometry and time-to-LOE  I withheld food from fish for 24 h and then transferred them to a custom designed calorespirometer (Chapter 2). Fish were allowed to habituate for 18 h after which measurements of normoxic routine O2 uptake rate (ṀO2) and metabolic heat were measured as in Chapter 3. The PwO2 in the calorespirometer chamber was then reduced from normoxia to 2.8 kPa over 1 h using compressed N2, and measurements of metabolic heat and flow-through ṀO2 were made. I chose a hypoxic PwO2 of 2.8 kPa based on preliminary O2 LC50 experiments that revealed 2.8 kPa to be the lowest PwO2 at which ≥75% of sticklebacks survived 8 h of exposure. The hypoxia exposure was maintained for 4 h and then normoxia was re-established for a period of 2 h to allow for recovery in the calorespirometer. Following the recovery period, I then performed closed-chamber respirometry to determine the fish’s critical O2 tension (Pcrit), which is the hypoxic PwO2 below which the fish is unable to extract sufficient environmental O2 to support a stable ṀO2. This stable ṀO2 can represent either standard metabolic rate (i.e., the ṀO2 of an awake, post-absorptive and entirely inactive ectothermic animal in the thermoneutral zone; Chabot et al., 2016) or routine metabolic rate (i.e., ṀO2 under the same conditions as standard metabolic rate but accounting for the small movements that are typical of fishes under experimental conditions). In this  83 chapter, I represented this stable range of ṀO2 as routine ṀO2, and calculated it (and Pcrit) according to Chapter 4.  A parallel set of experiments in which fish were held in opaque 10 L tanks was then carried out for the measurement of whole body concentrations of glycogen (glycolytic fuel) and lactate (glycolytic end-product). Each tank held four fish and, following an 18 h normoxic habituation period, PwO2 was adjusted in two tanks to 2.8 kPa (N=8) over 1 h and the other two remained normoxic to serve as controls (N=8). I repeated this for Alta Lake and Trout Lake fish. At 4 h, I inconspicuously introduced a lethal dose of anaesthetic (buffered MS-222, 0.3 g L-1) and once the fish were unresponsive (~4 min), I removed, weighed and froze each fish in liquid N2 for later metabolite analyses.  I measured hypoxia tolerance of the two populations by determining the time-to-LOE at a PwO2 of 1.3 kPa. Initial experiments to determine time-to-LOE were conducted at 2.8 kPa, but because Alta Lake fish survived for >72 h (Trout Lake fish survived <12 h), I decided to run the trials at 1.3 kPa instead. I ran these trials for each population by placing four fish in each of four tanks (N=16). Following an 18 h normoxic habituation period, I reduced PwO2 to 1.3 kPa over 1 h. I defined time-to-LOE as the time it took after the tank had reached 1.3 kPa for the fish to lose dorsal-ventral equilibrium and become unresponsive to gentle tail prods using a blunt dissection probe. When this point was reached, I removed the fish from the aquarium using a small dipnet, weighed the fish, and transferred it to a well-aerated recovery aquarium.  5.3.4  Metabolite assays  I measured changes in glycogen and lactate concentrations at the whole body level to be consistent with my whole body measurements of metabolic heat and ṀO2. I prepared whole bodies for metabolite extraction according to Chapter 3 and measured glycogen and lactate according to (Bergmeyer et al., 1983).  5.3.5  Statistical analyses  Statistical analyses were performed using SigmaStat 11.0. I used 2-way ANOVAs to analyze anaerobic metabolite concentrations and normoxic ṀO2 values, and 2-way repeated measures ANOVAs to analyze calorespirometry measurements of ṀO2 and metabolic heat.  84 The latter were also analyzed within each population (so as to identify potential use of MRD) using 1-way repeated measures ANOVAs with Tukey post hoc tests. Pcrit and time-to-LOE were analyzed using two-tailed t-tests. Any data set that did not meet the assumption of homogeneity of variance was log transformed prior to analysis. Differences were considered to be significant at P<0.05. All values are presented as means±sem.  5.4  Results  Time-to-LOE at a PwO2 of 1.3 kPa was twice as long in Alta Lake sticklebacks than Trout Lake sticklebacks (t-test, P<0.001, Fig. 5-1).   Closed-chamber respirometry revealed that both populations displayed a typical two-phase response in ṀO2 as PwO2 was decreased. A stable routine ṀO2 was regulated and maintained at PwO2 between 21 and 3.4 kPa in the Alta Lake sticklebacks, and between 21 and 4.1 kPa in the Trout Lake sticklebacks (Fig. 5-2A). At PwO2s below these values, ṀO2 then decreased with decreasing PwO2 (Fig. 5-2A). Routine ṀO2 assessed under normoxic conditions (between 18 and 21 kPa) did not differ as a function of population or respirometric technique (2-way ANOVA, P=0.075 and P=0.135, respectively; interaction P=0.337; Fig. 5-2B). Similarly, Pcrit did not differ significantly between the two populations (t-test, P=0.245, Fig. 5-2C), though these negative Pcrit results should be interpreted with caution due to the small sample size and resulting low statistical power (0.100).  Analysis of the calorespirometry results revealed that the time spent at 2.8 kPa PwO2 had a significant effect on both metabolic heat (P=0.029) and ṀO2 (P=0.007), but the lake of origin did not (P=0.128 and 0.285 for metabolic heat and ṀO2, respectively; interactions P=0.146 and P=0.204, respectively). Exploring each population’s capacity for MRD using 1-way ANOVAs revealed that the Trout Lake fish showed no significant change in metabolic heat compared with normoxic levels (P=0.9634, Fig. 5-3A), and a modest reduction in ṀO2 at 1 h of hypoxic exposure that returned to normoxic levels for the subsequent 3 h (P=0.009, Fig. 5-3B). However, the Alta Lake fish showed significant reductions in both heat (P<0.001, Fig. 5-3A) and ṀO2 (P=0.003, Fig. 5-3B), with a ~33% reduction in metabolic heat relative to normoxic levels by 4 h exposure.  Alta Lake sticklebacks had significantly higher whole body concentrations of glycogen (P=0.044, Table 5-2) than Trout Lake sticklebacks, but these concentrations were  85 unaffected by hypoxic exposure (P=0.355; interaction P=0.292; Table 5-2). Consistent with this, whole body concentrations of lactate did not differ as a function of population or hypoxic exposure (P=0.461 and P=0.068, respectively; interaction P=0.878; Table 5-2).  5.5  Discussion I predicted that sticklebacks native to an environment that experiences long-term hypoxia (Alta Lake) would be more tolerant of hypoxia than sticklebacks native to an environment that does not experience long-term hypoxia (Trout Lake), and that this difference would result from an increased reliance on MRD in the Alta Lake fish. The results agree with these predictions. The time-to-LOE results reveal that Alta Lake sticklebacks are approximately twice as tolerant of hypoxia than Trout Lake sticklebacks (Fig. 5-1), and the calorespirometry results suggest that this difference arises from the Alta Lake sticklebacks’ ability to employ MRD in hypoxic environments, something the Trout Lake sticklebacks are do not do (Fig. 5-3). Furthermore, the Pcrit analyses suggest that the two populations do not differ significantly in the PwO2 at which their ṀO2s start to decline (Fig. 5-2C), and the metabolite analyses suggest that, apart from slightly higher fuel reserves in the Alta Lake fish, the two populations show a similar capacity for and reliance on anaerobic glycolysis (Table 5-2).   5.5.1  Metabolic rate depression and hypoxia tolerance Alta Lake sticklebacks reduced metabolic rate by ~33% relative to normoxic routine values by 4 h exposure to 2.8 kPa PwO2 (Fig. 5-3A). At this same time point and PwO2, Trout Lake sticklebacks had a total metabolic rate approximately twice that of the Alta Lake fish (Fig. 5-3A), and this difference agrees well with the 2-fold longer tolerance time of the Alta Lake fish in the LOE experiment (Fig. 5-1). An MRD of 33% is similar in magnitude to the MRD measured in tilapia (Oreochromis mossambicus) at 3.1 kPa PwO2 (19% reduction; van Ginneken et al., 1997), and is greater than that measured in goldfish (Carassius auratus) at similar PwO2s (Chapter 3; van Ginneken et al., 2004), but is modest compared to the ~80% MRD measured in anoxic goldfish (Chapters 2, 3; Addink et al., 1991; van Ginneken et al., 2004). An MRD of 33% would still greatly benefit hypoxic survival by reducing rates of anaerobic fuel use and deleterious end-product accumulation, and is a magnitude beyond  86 what other hypoxia-native fish species like zebrafish (Danio rerio) are capable of (Stangl and Wegener, 1996). Furthermore, because my analyses were carried out at 17°C (to maximize calorimetric signal:noise) instead of the near-freezing water temperature that would occur in Alta Lake in the winter (see Table 5-1), the 33% MRD measured in my study likely underestimates the total metabolic savings accrued by the Alta Lake fish under long-term hypoxic conditions in the wild. For example, if we assume a 10°C reduction in temperature and a typical Q10 for metabolic rate of 2.5 [though ṀO2 measurements on brook stickleback (Culaea inconstans) at 5°C and 15°C reveal a Q10 of ~4.5 (Klinger et al., 1982)], the total metabolic savings accrued by Alta Lake stickleback during winterfreeze would be ~75%. Larger temperature reductions and Q10 values, both plausible, would result in greater savings. To my knowledge, these results represent the first time MRD use has been shown to vary significantly among geographically isolated populations of the same species. While MRD is believed to be a complex biochemical phenotype requiring a reorganization of cellular processes, the fact that the Alta Lake and Trout Lake populations have been isolated for a maximum of 11 000 y (Bell and Foster, 1994; Mathews and Fyles, 1970) suggests that it is a phenotype that can evolve rapidly. This is consistent with the rapidity with which marine sticklebacks have been shown to naturally evolve their freshwater-distinctive, genetically based morphological features de novo (<50 y; Lescak et al., 2015). However, an intriguing alternative hypothesis might be that MRD is a developmentally plastic neonatal characteristic of the Alta Lake fish, one that is developmentally lost in the Trout Lake fish. Similar developmental plasticity has been shown in the hypoxic ventilatory responses (HVR) of mammals, where exposure to hypoxia during a critical time window following birth can alter the HVR in adulthood [see Teppema and Dahan (2010) for review]. Support for this applying to MRD in fishes is provided by results for zebrafish, where embryos are capable of hypoxia-induced MRD (Padilla and Roth, 2001) but adults are not (Stangl and Wegener, 1996). Hypoxic exposure during a critical window of development might therefore preserve the ability to induce MRD in adulthood. For the stickleback in my study, the long-term hypoxia experienced by the Alta Lake fish could possibly provide this cue for developmental plasticity. This is worth further investigation.   If MRD enhances hypoxia tolerance, then why wouldn’t all fish employ it in hypoxia? The answer may involve environmentally induced developmental plasticity as  87 described above, but could also involve the potential for reduced fitness when MRD is employed in certain environments. Physiological and ecological costs of MRD include oxidative damage (Carey et al., 2000), reduced growth, repair and immunocompetence (Burton and Reichman, 1999), cognitive impairments stemming from neuronal damage (Popov et al., 1992), ceased reproduction (Humphries et al., 2003), and increased susceptibility to predation by aquatic and aerial predators stemming from significantly reduced motor activity (Humphries et al., 2003). For the Trout Lake sticklebacks that do not naturally see long-term hypoxia, these costs, particularly the ecological ones, may outweigh any selection for MRD. But for the Alta Lake sticklebacks that do see long-term hypoxia, the costs of not employing MRD may outweigh the potential costs that come with MRD per se, especially because some of these costs could be mitigated during the wintertime when the sticklebacks were in their metabolically depressed states. For example, the surface layer of ice would eliminate the risk of aerial predation, and the migrations and/or reduced appetites of the resident trout (Oncorhynchus clarkii, O. mykiss) would reduce the risk of aquatic predation (Klemetsen et al., 2003). Moreover, the relatively low hypoxia tolerances of trout (Wagner et al., 2001) tend to restrict them to the uppermost water layers in winter, where O2 levels are highest (Brown et al., 2011; Ultsch, 1989). This could enable the sticklebacks to use the deeper, more hypoxic water layers as refugia from the trout similar to how other species exploit hypoxic environments such as swamps (Chapman et al., 2002; Chapman et al., 1995), coral reef crevices (Nilsson and Östlund-Nilsson, 2004) and oceanic O2 minimum zones (Brill, 1994; Nasby-Lucas et al., 2009; Seibel, 2011) to escape being eaten by their less-tolerant predators. Physiological costs of MRD may still accrue if these sticklebacks have not evolved mechanisms to mitigate them (Humphries et al., 2003), but these costs are generally sub-lethal and may be outweighed by the survival risks that come with not employing MRD during winter freeze. Indeed, long-term deprivation of some essential abiotic factor such as food, water or heat is the common theme underlying diverse animal taxas’ use of hibernation, aestivation and torpor; it is therefore not unreasonable to speculate that long-term deprivations of O2 have driven the evolution of MRD use in fishes.   88 5.5.2  Intraspecific variation in hypoxia tolerance The isolation of these two populations and the variation in their respective natural habitats make for a powerful comparative system with which to study physiological adaptation. A similar approach has been taken with the Atlantic killifish, with various research groups using comparisons of northern and southern populations (subspecies F. h. macrolepidotus and F. h. heteroclitus, respectively) to test a wide range of hypotheses related to environmentally induced physiological adaptation (see reviews by Burnett et al., 2007; Schulte, 2014; Schulte, 2007). Incidentally, to my knowledge, Atlantic killifish are the only other fish species within which significant variation in hypoxia tolerance has been shown to exist. In that recent study, the southern subspecies maintained dorsal-ventral equilibrium for ~1.7-fold longer at 0.4 kPa PwO2 (at 15°C) than the northern subspecies (McBryan et al., 2016). The authors attribute the enhanced tolerance of the southern subspecies to their lower routine normoxic metabolic demand, and this strategy may also contribute to the enhanced tolerance of the Alta Lake stickleback (albeit non-significant; P=0.075, Fig. 5-2B). In any case, comparisons of isolated threepine stickleback populations have revealed variation in the physiology underlying environmentally induced differences in thermal tolerance (Barrett et al., 2011; Gibbons et al., 2016), osmoregulation (Gibbons et al., 2016; McCairns and Bernatchez, 2010; Schaarschmidt et al., 1999) and aerobic performance (Dalziel et al., 2012a; Dalziel et al., 2012b); my results here suggest that these comparisons could also be used to address questions regarding the mechanisms and evolution of hypoxia tolerance.  5.5.3  Conclusions  These results demonstrate two novel findings: first, hypoxia tolerance can vary within a species and evolve over relatively short timescales; and second, MRD, a complex biochemical phenotype, can improve hypoxia tolerance and evolve relatively quickly, particularly in long-term hypoxic environments. Investigating the hypoxia tolerance strategies of other stickleback populations that have independently colonized winterfreeze and non-winterfreeze lakes could shed light on the adaptive value of the responses measured in this study. Furthermore, my results demonstrate that the threespine stickleback, with its isolated populations native to different hypoxia environments, presents a powerful model system with which to investigate the mechanisms and evolution of hypoxia tolerance. Such  89 knowledge will improve our understanding of hypoxia tolerance in general, and importantly, benefit predictive models and conservation efforts that help identify and protect potentially vulnerable species in an increasingly hypoxic world.   Figure 16                            90   Figure 5-1. Time taken for two populations of threespine sticklebacks to lose dorsal-ventral equilibrium and become unresponsive to gentle tail prods when exposed to severely hypoxic water (1.3 kPa PwO2). Water was reduced from ~21 kPa to 1.3 kPa over 1 h following an 18 h normoxic habituation period. Experiments were run separately for each population and in four separate 10 L aquaria housing four fish each (N=16 for each population). Points represent individual values, horizontal line indicates mean value, error bars indicate s.e.m., and asterisk indicates significant difference (t-test, P<0.001).            0306090120Alta Lake Trout LakeTime-to-LOE at 1.3 kPa (min)*  91 Figure 17   Figure 5-2. Closed-chamber ṀO2 measurements as a function of PwO2 (A), normoxic ṀO2 (B), and Pcrits (C) for two populations of threespine sticklebacks. PwO2 was reduced from ~20 to 0 kPa by the fish’s own respiration over the course of ~90 min. (B) Average ṀO2 values at normoxic PwO2 for two populations of threespine sticklebacks as determined using closed-chamber and flow-through respirometry. Normoxic ṀO2 is not significantly affected by population or technique (2-way ANOVA, P=0.075 and P=0.135, respectively; interaction P=0.337). (C) Average Pcrit values for two populations of threespine sticklebacks, calculated for each fish from its closed-chamber ṀO2 trace (t-test, P=0.245). Points represent individual values, horizontal line indicates mean value, and error bars indicate s.e.m. N=5 for Alta Lake and N=4 for Trout Lake. 0 5 10 15 20051015Alta LakeTrout LakeAWater PO2 (kPa)MO2 (µmol g-1 h-1)051015 Alta LakeTrout LakeClosedrespirometryFlow-throughrespirometryBNormoxic MO2 (µmol g-1 h-1)0246Alta Lake Trout LakeCP crit (kPa PO2) 92 Figure 18   Figure 5-3. Flow-through calorespirometric measurements of metabolic heat (A) and ṀO2 (B) as a function of time in severe hypoxia (2.8 kPa PwO2) for two populations of threespine sticklebacks. Normoxic measurements were made following an 18 h habituation period, after which PwO2 was reduced from ~21 kPa to 2.8 kPa over 1 h. Subsequent measurements were made at 1, 2, 3 and 4 h following the point when PwO2 reached 2.8 kPa. Points indicate average values and error bars indicate s.e.m. Statistical tests (1-way ANOVAs) were run within populations and values sharing a letter are not significantly different (Alta Lake: heat P<0.001, ṀO2 P=0.003, N=7; Trout Lake: heat P=0.963, ṀO2 P=0.000, N=4).     0246 Aaabbc cbcAAA A AAlta LakeTrout LakeMetabolic heat (J g-1 )0246810 Baab ab b bAB AB AB ABNormoxia 1h 2h 3h 4hMO2 (µmol g-1 h-1) 93 Table 5-1. Characteristics of Alta Lake and Trout Lake. Data provided by aWhistler Museum, bBritish Columbia Ministry of Environment, cJacques Whitford/AXYS Environmental Consultants, and dSeth Rudman.  Days frozen Elevation Surface area Average depth Maximum depth Surface pH Surface conductivity Fish species Alta Lake 128a 602b 995600b 9.5b 24.4b 6.5 – 7.7c 89c O. clarkiib O. mykissb C. asperb Trout Lake 0 157b 75600b 5.8b 17.4b - 75d O. clarkiib Table 0-1 Days frozen in mean d y-1 since 1942; elevation in m; surface area in m2; depth in m; conductivity in Siemens m-1; fish species (Oncorhynchus clarkii, cutthroat trout; Oncorhynchus mykiss, rainbow trout; Cottus asper, prickly sculpin) in addition to stickleback observed over past ten years.                    94 Table 5-2. Whole body concentrations of glycogen and lactate in two populations of threespine sticklebacks. Measurements were made following a 4 h exposure to normoxia or hypoxia (2.8 kPa PwO2). N=8 for each treatment.     Alta Lake  Trout Lake    2-way ANOVA P values  Normoxia Hypoxia Normoxia Hypoxia Pop. PO2 Interaction [Glycogen] 1.37±0.19 1.32±0.11 0.89±0.17 1.19±0.10 0.044 0.355      0.292 [Lactate] 0.15±0.05 0.30±0.06 0.21±0.07 0.35±0.11 0.461 0.068      0.878 Table 0-2 Concentrations in µmol g-1 tissue; values are mean±s.e.m.    95 Chapter 6  General discussion       The objective of my thesis was to determine how hypoxia-exposed fishes combine aerobic metabolism, anaerobic metabolism and MRD as a total HMR, and how this HMR varies in different types of hypoxic environments. I addressed this using the goldfish (Carassius auratus) and the threespine stickleback (Gasterosteus aculeatus). The results of Chapters 3 and 4 show that the HMR of goldfish is affected by the severity (i.e., PwO2), duration and RHI of the hypoxic exposure, while the results of Chapter 5 show that variation in naturally occurring environmental hypoxia results in variation in the HMRs of two populations of threespine sticklebacks. These results together suggest that the HMR is dynamic, changing as a function of both the biology of the fish and the abiotic aspects of its hypoxic environment. Because natural hypoxic environments vary with respect to PwO2 and time, it is therefore likely that the HMR of a given species (or population) is finely tuned to characteristics of its hypoxic environment.  This Discussion will explore how the naturally occurring O2 dynamics of different hypoxic environments shape the HMRs of their resident species. I will explore this idea using a two-dimensional matrix of environmental hypoxia (Fig. 6-1), and by the end, I hope to identify patterns in the HMRs favoured in different hypoxic environments. But first, I will highlight what my thesis work has contributed to our understanding of hypoxic survival mechanisms and the methods I use to elucidate them.      96 6.1  Thesis highlights and main contributions 6.1.1  Calorespirometry: easier than might be expected  Our calorespirometer is one of only two instruments that have been used to measure the metabolic heat of adult fishes over the past 40 years. The other instrument, belonging to Guido van den Thillart’s group in Leiden, was used for a series of six studies between 1989 and 2004. This is negligible compared to the countless number of studies that have used respirometry over that same time. Given direct calorimetry’s widely recognized superiority over respirometry as a technique of metabolic rate measurement (Kaiyala and Ramsay, 2011; Lighton, 2008; Nelson, 2016; Richards, 2009; van Ginneken and van den Thillart, 2009), it is surprising that so few studies have used it. The reason typically put forward for this involves the difficulty of the technique, an idea originally espoused by leaders in the field (Fry, 1971; Brett and Groves, 1979) and one that has since become accepted as fact. This has prevented many otherwise interested investigators from pursuing calorimetry (or calorespirometry), and perhaps because of the small market for fish-accommodating calorimeters, the one company that offers a turnkey model charges $168,000 for it. If the technique’s perceived difficulty does not prove inhibitory, its known cost very well might!  However, having co-designed, built and used a calorespirometer myself, I can say that the perceived hurdles are not nearly as high as they are purported to be. Building the apparatus is relatively straightforward so long as one has access to machining tools (an experienced machinist helps), as is assembling it with the necessary accessories (e.g., peristaltic pump, O2 equilibration chamber, insulated ice chest). The most difficult step might be optimizing the calorespirometer’s function with respect to its particular environment and the conditions of the desired experiments, entirely the result of the apparatus’s (necessary) thermal sensitivity. I discussed some of these difficulties in Chapter 2, and by and large they can be overcome through trial and error. So long as the appropriate steps are taken to optimize the apparatus’s heat-detecting function, running calorespirometry experiments is no more involved than running typical respirometry experiments. And finally, the same design remains effective when scaled to different sizes, allowing experiments to be performed on animals of various sizes. The only major limitation of our apparatus is a signal-to-noise ratio that precludes precise measurements on ectothermic animals at low temperatures, but this  97 could be mitigated by arranging the inflowing and outflowing waterlines counter-currently (to maximize signal) and/or improving the system’s insulation (to reduce noise). Combined with its low cost of construction ($1300) and a published paper detailing its construction and function, it was my hope to dispel some of calorespirometry’s discouraging aspects and make it more accessible to investigators. Encouragingly, there are at least three groups around the world who have built (or are in the process of building) calorespirometers based on my design to address their own questions.  6.1.2  Hypoxia-induced MRD: a response reserved for extreme environments  My results from Chapters 3 and 5 suggest that, at least in goldfish and sticklebacks, MRD is a hypoxic response that is reserved for extreme environments. This could have only been shown with calorespirometry. ‘Extreme’ hypoxic environments are those that experience particularly low PwO2s and/or particular long duration hypoxic events. Chapters 3 and 4 reveal that goldfish prioritize maintaining routine metabolic rate through different plastic and time-dependent mechanisms (aerobic + anaerobic over short durations and rapid RHIs; aerobic over long durations and gradual RHIs), and reserve their pronounced MRD for near-anoxia. Chapter 5 reveals that sticklebacks native to an environment that experiences deep, long-term hypoxia (a winterfreeze lake) have evolved the ability to use MRD, while sticklebacks native to an environment that does not experience deep, long-term hypoxia have not evolved (or at least do not express) this ability. This pattern of MRD use may be related to the physiological and ecological costs that come with using MRD, and I elaborate on this idea in section 6.2.  Finally, the use of calorespirometry allowed me to determine the PwO2 at which goldfish induce MRD. This PwO2 (~0.5 kPa) coincides with goldfish’s Hb P50 value, and so I hypothesized that tissue O2 delivery via Hb might be an upstream signal for hypoxia-induced MRD in goldfish. With the help of a recently acquired drug (efaproxiral) that significantly increases goldfish’s Hb P50 value, this hypothesis makes some clear predictions that I am now in the process of testing experimentally using cannulated goldfish and a larger version of the calorespirometer.   98 6.1.3  Pcrit: musings on biology and methodology  My results from Chapter 4 show that the RHI used in a Pcrit experiment can significantly alter the Pcrit of the animal in the experiment (goldfish, in my case). Though this idea has been speculated on before (e.g., Rogers et al., 2016; Snyder et al., 2016), my Chapter 4 experiments are the first to provide empirical support for it. The underlying mechanisms involve rapid alterations to steps along the O2 transport cascade, including lamellar surface area and Hb-O2 binding affinity. These alterations may occur more quickly in goldfish than in other species, but because all species have some capacity for environmental acclimation, it is probably true that RHI will affect all species to a corresponding degree and should therefore be controlled for when running hypoxic exposure experiments. Furthermore, because the results showed no change in Pcrit with respirometric method per se, closed-chamber respirometry is a viable and effective method of Pcrit determination despite the modest accumulation of CO2 and ammonia that comes with it.  Were anyone to ask, my recommended method for Pcrit determination would be closed-chamber/circuit respirometry over a ‘typical’ time duration of ~60 to 90 min. This time scale captures a fish’s Pcrit in the present, before hypoxic acclimation is able to exert a significant effect on O2 extraction mechanisms and subsequently Pcrit. While this may underestimate the fish’s full ability to take up and use O2 in hypoxia (i.e., overestimate the fish’s Pcrit), it at least captures the fish’s ability at a known point in the acclimation process (i.e., pre-acclimation). Slowing the RHI by extending the Pcrit trial duration will result in the fish acclimating to hypoxia to some degree, and because this degree will vary among species and individuals, knowing where in the acclimation process a given RHI will place a particular fish is difficult. This jeopardizes the investigator’s ability to standardize among species and individuals, and subsequently, to compare Pcrit values among these groups. And because the majority of studies have determined Pcrits of fishes using closed-chamber respirometry (Rogers et al., 2016), sticking with this method will allow for the most relevant comparisons to literature values.  6.1.4  Threespine stickleback: a potentially powerful model for hypoxia research  My results from Chapter 5 reveal that different populations of threespine sticklebacks vary significantly in their hypoxia tolerances and abilities to use MRD. As far as I know, this  99 is only the second time such intraspecific variation in hypoxia tolerance has been shown to exist in fishes (the other instance being two subspecies of Atlantic killifish; McBryan et al., 2016), and the only time such intraspecific variation in MRD has been shown to exist. The threespine stickleback is therefore an excellent system with which to investigate the mechanisms and evolution of these traits, similar to how intraspecific comparisons of Atlantic killifish subspecies have been used to investigate the mechanisms and evolution of thermal tolerance (see reviews by Schulte, 2007; Schulte, 2014). Adding to the stickleback’s potential is the vast research field that surrounds the species, arising first from the species’ evolutionary history that lends itself to the study of local ecological adaptation, and pushed forward by methodological resources (e.g., a sequenced genome) and an ever-growing body of stickleback literature. Combined with their prevalence on the west coast of North America, their ease of capture, and their adaptability to laboratory conditions, threespine sticklebacks represent an exciting new model system with which to probe the proximate and ultimate causes of hypoxia tolerance.  6.2  The hypoxic environment as sculptor of the HMR  Ultimately, life in hypoxia is about matching ATP supply and demand (Boutilier, 2001; Hochachka and McClelland, 1997). This balance is possible through different combinations of aerobic metabolism, anaerobic metabolism and MRD, and hypoxia tolerance involves any combination of these metabolic strategies. A ‘grand unified theory’ of hypoxia tolerance is therefore unlikely to exist. Rather, the strategy adopted by a species (or even a population or individual) is more likely shaped by its life history and the particular hypoxic environment to which it has become adapted. This hypothesis predicts that different species native to environments with similar O2 characteristics (in terms of severity, RHI and/or duration) will employ similar hypoxic survival strategies, while those native to environments with different O2 characteristics will employ different hypoxic survival strategies.  With the help of a two-dimensional matrix of environmental hypoxia (Fig. 6-1), I will explore how time- and PwO2-related O2 characteristics affect the HMRs of species adapted to three representative natural hypoxic habitats: severe PwO2 + short duration (Q2 in Fig. 6-1), moderate PwO2 + long duration (Q3 in Fig. 6-1), and severe PwO2 + long duration (Q4 in Fig. 6-1). For the sake of simplicity, I have incorporated RHI into the time axis of Fig. 6-1  100 because, though exceptions exist, it is generally the case that the short-duration hypoxic habitats (Q1 and Q2) have rapid RHIs, while the long-duration hypoxic habitats (Q3 and Q4) have gradual RHIs.  6.2.1 Q2: Severe PwO2 + short duration  Tidepools in the intertidal zone become isolated from the ocean as a result of tidal ebbs and flows. Pools located high in the intertidal zone relative to sea level at low tide can become isolated for hours to days. Their small water volumes, combined with their often-dense biota, result in enormous fluctuations in PwO2, reaching anoxia at night and up to 80 kPa in the day and typically taking place over just 6 to 12 h (Richards, 2011; Truchot and Duhamel-Jouve, 1980). These tidepool habitats are therefore typified by hypoxia that is severe (PwO2) and short (duration).  Numerous animal species make their home in these tidepools, and a well-studied example is the tidepool sculpin (Oligocottus maculosus). The tidepool sculpin uses a variety of mechanisms to achieve a very high tolerance of hypoxia, and I can use a series of studies by Mandic and colleagues to put together how it employs some of these mechanisms as environmental PwO2 decreases. First, the sculpin uses a high Hb-O2 binding affinity (P50 of 3.1 kPa) to regulate routine ṀO2 from normoxic PwO2s to 3.5 kPa (Pcrit of 3.5 kPa; Mandic et al., 2009a). As PwO2 falls below this value, the sculpin engages behavioural responses that increase its access to O2, including aquatic surface respiration (ASR) starting at 2.8 kPa, and then aerial emergence starting at 1.2 kPa (Mandic et al., 2009a). Aerial emergence allows fishes that are capable of it to support routine ṀO2 levels amid severely hypoxic (i.e., sub-Pcrit) water (Martin, 1996; Yoshiyama and Cech, 1994), and indeed for tidepool sculpins in particular, 72 h of aerial emergence has been shown to result in no significant accumulation of lactate (Sloman et al., 2008). These responses therefore uncouple the sculpin from its aquatic habitat and allow it to maintain routine ṀO2 levels at sub-Pcrit PwO2 values. However, they also significantly increase the risk of aerial predation (Kramer and Manley, 1983). A perceived threat of predation from above will send the sculpin back into the tidepool’s severely hypoxic water (or delay its emergence from it; Hugie et al., 1991; Shingles et al., 2005; Sloman et al., 2008), and so their survival depends on anaerobic metabolism and/or MRD until the predation threat subsides. The tidepool sculpin’s capacity for hypoxia-induced  101 MRD has never been investigated, but recent work by Mandic et al. suggests that the sculpin’s capacity for anaerobic glycolysis is highly adapted to its hypoxic environment. Specifically, their glycogen reserves are large (Mandic et al., 2013), their glycolytic enzyme activity levels are significantly higher than those of closely-related sculpin species native to less hypoxic sub-tidal environments (Mandic et al., 2013), and the expression of genes involved in anaerobic glycolysis do not change over the course of an 8 h hypoxia exposure, suggesting their high anaerobic capacity is a fixed trait that is capable of being immediately and maximally induced (Mandic et al., 2014). A reliance on anaerobic glycolysis over MRD is practical in this environment because it allows the sculpin to maintain cellular energy balance without impairing predator avoidance behaviour through reduced responsiveness and locomotor abilities. However, there are costs that come with anaerobic reliance. These costs (fuel depletion, end-product accumulation) are positively correlated with time spent in the hypoxic environment, but for the sculpin, this would typically be short; either the threat of predation will subside and allow the fish to reemerge and/or perform ASR, or the tidepool’s water will be replenished with O2 by photosynthesis and/or the rising tide. The costs accrued with anaerobic reliance would therefore be low. The tidepool sculpin prioritizes aerobic metabolism under all possible hypoxic conditions, and likely relies on anaerobic glycolysis (not MRD) when forced to spend time in severely hypoxic water. If this HMR is a product of the tidepool sculpin’s natural hypoxic environment, then I would predict that different species that are native to similar hypoxic habitats (i.e., severe and short) would exhibit similar HMRs. One such example is the zebrafish (Danio rerio).  Zebrafish are native to rice paddies and stagnant water bodies in India that become severely hypoxic as a result of high eutrophication and plant respiration (Cruz et al., 2000; Magneschi and Perata, 2009; Spence et al., 2008). An abundance of aerial predators are typical of these habitats and are believed to be the reason why zebrafish generally avoid the water’s surface (Blaser and Goldsteinholm, 2012; Spence et al., 2008). In many ways, this is a similar hypoxic habitat to the tidepools described above, and perhaps because of this, the HMR of zebrafish is similar to that of the tidepool sculpin. Zebrafish have a low Pcrit value of ~2.7 kPa (Barrionuevo et al., 2010) and will employ ASR to supplement O2 uptake starting at 4 kPa (Abdallah et al., 2015). However, a threat of predation from above would likely cause  102 the zebrafish to retreat to deeper, more hypoxic waters. This zone could theoretically be used as a refuge from aerial predators, but like the sculpin retreating into the tidepool, residency here would require a reliance on anaerobic glycolysis and/or MRD. Calorimetric measurements of hypoxia-exposed zebrafish by myself (unpublished) and others (Stangl and Wegener, 1996) clearly show that zebrafish do not depress metabolic rate at either 2.0 kPa PwO2 or near-anoxia, and this maintenance of metabolic heat at sub-Pcrit PwO2s suggests a strong activation of anaerobic glycolysis. Indeed, glycolysis-related gene expression has been shown to increase in sub-Pcrit-exposed zebrafish (Ton et al., 2003; van der Meer, 2005). Furthermore, anaerobic metabolism (as represented by glycolytic enzyme activities and metabolite concentrations) is believed to be the metabolic strategy underlying variation in hypoxia tolerance among nine related Danio and Davario species (Yao, 2012). Buffering routine metabolic rate with anaerobic glycolysis while in deeper, hypoxic waters would allow the zebrafish to retain some level of activity, and this would be advantageous considering the abundance of air-breathing predatory fishes (e.g. Notopterus notopterus, Xenenetodon cancila, Channa spp.) that inhabit this environment (Engeszer et al., 2007a; Engeszer et al., 2007b; Spence et al., 2008). All told, the zebrafish, like the tidepool sculpin, appears to prioritize the use of aerobic metabolism and resort to anaerobic glycolysis instead of MRD at sub-Pcrit PwO2s. This also appears to be the case with Atlantic killifish (Fundulus heteroclitus), whose estuarine habitats are similar to the sculpin’s and zebrafish’s in terms of O2 regime and predation (Burnett et al., 2007; Kneib, 1982; Schulte, 2007); killifish acclimated to intermittent hypoxia up-regulate mechanisms that enhance glycolytic capacity and the processing of glycolytic end-products, while killifish acclimated to chronic hypoxia do not (Borowiec et al., 2015). That the zebrafish’s and killifish’s natural hypoxic environments are similar to the tidepool sculpin’s suggests that this HMR is well suited—and perhaps selected for—in predator-rich environments experiencing rapid and severe fluctuations in PO2. But what happens to the HMR if predation pressure is removed from these short and severe hypoxic habitats? I can address this question with the help of migratory species of oceanic O2 minimum zones. Oceanic O2 minimum zones (OMZs) occur throughout the world’s oceans at depths between 200 and 1000 m, where certain biological and physical processes combine to reduce dissolved O2 levels to PwO2 values <6.4 kPa around the OMZ’s periphery and often as low as  103 0.5 kPa in its center. Biologically, a high density of aerobic bacteria reduce the OMZ’s O2 levels as they feed upon the organic matter falling from the mixed layer above, while physically, a lack of atmospheric contact and low levels of convective mixing keep these waters low in O2.  There are two types of OMZ resident: permanent and migratory. Permanent residents, which will be discussed later, spend their entire lives in the OMZ and therefore experience chronic moderate-to-severe PwO2s (Q3 and Q4 in Fig. 6-1). Migratory residents on the other hand spend their days in the center of the OMZ and migrate vertically into well-oxygenated surface waters each night to feed in the cover of darkness (Seibel, 2011). This migratory pattern exposes these animals to progressively changing PwO2s, becoming normoxic with upwards migration and hypoxic with downwards migration. The hypoxic exposures experienced by these animals are therefore severe (PwO2) and short (duration; Q2 in Fig. 6-1), similar to those of the sculpins and zebrafish described above. Despite this similarity, migratory OMZ residents use a different HMR. While they tend to possess traits that enhance O2 extraction (e.g., Seibel, 2013; Trueblood and Seibel, 2013) and glycolytic capacity (e.g., Gonzalez and Quiñones, 2002; Torres et al., 2012), migratory OMZ residents rely primarily on MRD while in the deeply hypoxic OMZ during the day (Seibel, 2011; Seibel et al., 2016). For example, the jumbo (or Humboldt) squid (Dosidicus gigas) depresses metabolic rate by 87% when held at 0.6 kPa, the PwO2 at which it typically spends the daytime in the OMZ (Rosa and Seibel, 2010; Trueblood and Seibel, 2013). Migratory krill (Euphausia eximia) from this same OMZ region also employ MRD at this PwO2 (Seibel, 2011; Seibel et al., 2016). These are different HMRs than those employed by tidepool sculpins and zebrafish despite similar environmental O2 characteristics, and the reason might involve predation risk. As discussed, predation risk in the hypoxic habitats of the sculpin and zebrafish is high, and so employing MRD, with its concomitant reduction in locomotor activity and responsiveness (Humphries et al., 2003), would likely increase the chances of being eaten. However, predation risk in the OMZ is relatively low owing to low levels of light and activity, a diffuse distribution of animals (see Childress, 1995; Drazen and Seibel, 2007; Seibel and Drazen, 2007; Seibel et al., 2000), and low O2 levels that tend to keep top ocean predators like sharks, tunas and billfishes out (Brill, 1994; Nasby-Lucas et al., 2009; Vetter et al., 2008). Consequently, animals living in the OMZ—particularly those that migrate into oxygenated  104 surface waters to complete necessary behaviours like feeding and mating—can employ MRD with a relatively low risk of being eaten.   6.2.2 Q4: Severe PwO2 + long duration The Amazon basin floods each year when the Amazon River, which drains the Andes, overflows. When this happens, water spills over the banks of the river and into the surrounding forests and floodplains, bringing with it many of the Amazon’s 5600+ species of fish (Albert and Reis, 2011). At the peak of the wet season, all of the flooded areas are interconnected, allowing fish to move among them. But as the season wears on, the water levels recede and leave behind smaller, isolated water bodies that become hypoxic (even anoxic) as a result of plant and animal respiration, organic decomposition, and a lack of light penetration, remaining deeply hypoxic for months at a time, even chronically. These habitats are therefore typified by hypoxia that is severe (PwO2) and long (duration). Perhaps because of the seasonality (i.e., predictability) of these flood pulses, many of the Amazon’s fish species have independently evolved a high tolerance to hypoxia (Almeida-Val and Val, 1993). Most of these species achieve this by using various behavioural and/or morphological features that maximize their abilities to acquire O2 in their O2-depleted habitats, the prime examples being ASR and air breathing. Many Amazonian fish families have independently evolved morphological features to optimize ASR and air breathing, including extensible lower lips to syphon O2-rich water directly across the gills, and a wide variety of air breathing organs ranging from modified buccal cavities to lungs (Val and Almeida-Val, 1998). These morphological features, and the behaviours they optimize, are believed to have evolved in response to aquatic hypoxia (Graham, 1997; Kramer and McClure, 1982). It is no surprise then that they are widely used among the Amazon’s hypoxia-dwelling fish species. One study collected the resident species of an isolated Amazonian lake that had become severely hypoxic (Camaleao Lake) and determined the primary hypoxic adaptation used by each species to survive in the lake. Of the 11 families caught (numerous species for most), seven used air-breathing as their means of tolerating hypoxia, two used ASR, one used Hb-O2 binding affinity modulation, and one used MRD (Junk et al., 1983). In a similar study in which 20 species were caught in a hypoxic Amazonian lake, ten species used ASR as a primary means of tolerating hypoxia, four used  105 air-breathing, four positioned themselves directly adjacent to O2-secreting plant roots, one combined a large gill surface area with a high Hb-O2 binding affinity, and one used MRD (Astronotus ocellatus, the most tolerant of the group; Soares et al., 2006). Furthermore, while air-breathing and/or ASR behaviours increase the susceptibility of aerial predation (Kramer and Manley, 1983), some of these fishes have evolved complex group behaviours to mitigate this risk (Sloman et al., 2009).  In addition to ASR and air breathing, hypoxia-adapted Amazonian fishes tend to possess a suite of physiological characteristics that enhance O2 extraction and delivery, including increased ventilation, blood-O2 carrying capacity, and Hb-O2 binding affinity via decreased allosteric interactions (reviewed by Val and Almeida-Val, 1995; Val and Almeida-Val, 1998). The obvious trend here is for hypoxia-adapted Amazonian species to prioritize aerobic metabolism over anaerobic metabolism and/or MRD, perhaps unsurprising given the benefits of maintaining routine metabolic rate aerobically and the fact that these species typically have constant access to air. But what about long-term, severely hypoxic habitats that do not allow access to air? Winterfreeze lakes generally occur at high elevations or at far northern or southern latitudes, where wintertime atmospheric temperatures stay below 0°C for sufficient time periods to freeze the lake’s surface layer. The breakdown of organic matter by aerobic microorganisms depletes the liquid water’s dissolved O2 levels, and a lack of water-atmosphere interface and photosynthesis (due to the ice layer reducing or eliminating the passage of light to aquatic plants) prevents them from being replenished until spring thaw (Ultsch, 1989). The hypoxic severity therefore depends on the lake’s depth, biological activity, flowing water supply, and ice cover dynamics and duration, with severe lakes becoming anoxic for months at a time (Barica and Mathias, 1979; Mathias and Barica, 1980). The rate at which O2 is depleted will depend on these same parameters, but even for a severe lake, the descent from normoxia to anoxia typically takes 2+ months (Vornanen et al., 2004). These winterfreeze lakes are thus typified by hypoxia that is severe (PwO2), long (duration) and slow developing.  Various species overwinter beneath the ice of frozen lakes, including reptiles (snakes and turtles), amphibians (frogs and salamanders) and numerous species of fishes (Ultsch, 1989). The reptiles and amphibians tend to bury themselves in the sediment and effectively  106 isolate themselves from the water environment, while the fishes, which I will focus on, tend to maintain some degree of activity and environmental engagement (but see Crawshaw et al., 1982; Loeb, 1964). Species assemblages in winterfreeze lakes are often determined by the magnitude of the winter hypoxic event. The limited tolerances of predatory fishes (e.g., Esocidae, Centrarchidae) restrict their ranges and preclude them from severe winterfreeze lakes, allowing more tolerant fishes (e.g., Cyprinidae, Umbridae) to colonize these lakes with a reduced risk of predation (Magnuson et al., 1989). An excellent example of such a species is the crucian carp (Carassius carassius), a cyprinid native to small lakes in northern Europe and Asia that is “probably the most anoxia-tolerant fish there is” (Vornanen et al., 2009). Similarly tolerant, though slightly less so, is the crucian carp’s cogener, the goldfish (Carassius auratus). The goldfish is native to the same type of habitat as the crucian carp and employs the same suite of anoxia-tolerance traits (though perhaps to a lesser degree; Ultsch, 1989). It too is therefore an excellent species with which to explore adaptations to winterfreeze hypoxia, and I will use the data collected in Chapters 3 and 4 to do so. The HMR of goldfish (described above) is well suited to its native winterfreeze lakes. Exceptional O2 uptake abilities—the plasticity of which reduces osmo-respiratory and O2 extraction-delivery tradeoffs—allow the goldfish to maintain routine metabolic rate at all but near-anoxic PwO2s, preserving some level of routine function for most of the winter hypoxia exposure while simultaneously conserving finite anaerobic fuels until their use is absolutely necessary. As the winter progresses and near-anoxic PwO2 values are encountered, the goldfish induces a significant MRD that reduces the rate at which these fuels are used while simultaneously reducing the rate at which deleterious anaerobic end-products accumulate. However, if the exposure lasts sufficiently long for those end-products to accumulate, they are converted to ethanol and excreted across the gills, thus preventing a metabolic acidosis while delaying the loss of carbon until it is absolutely necessary. Metabolic rate depression is a critical component of this HMR, one that on its own can extend the goldfish’s anoxic survival time by approximately 25-fold when temperature is accounted for (assuming Q10 of ṀO2 of 4.6; Fry and Hart, 1948). It is probably true that MRD is the only mechanism by which a fish can survive anoxia for any appreciable amount of time, regardless of how large their glycogen reserves are or how they may deal with their anaerobic end-products (Hochachka et al., 1996). Even for a species like goldfish, with its  107 exceptionally large glycogen reserves and its ability to avoid a glycolytically-derived acidosis, anoxic survival time would only be ~5 h at routine metabolic rates	  [assuming P:O2 of 6, routine ṀO2 of 3.5 µmol g-1 h-1, P:glycogen of 3 (Hochachka and Somero, 2002), and estimated total body glycogen reserves of 35 µmol g-1 (Mandic et al., 2008)]. With MRD and a reduced water temperature, this time would be extended to approximately 5 days. The combination of goldfish’s native environment becoming anoxic in the winter, the requirement of MRD to survive appreciable periods of time in anoxia, and a low predation risk in these anoxic environments therefore probably explains why MRD was selected for in these animals. So can MRD be expected to play a role in the HMRs of all fish species native to winterfreeze lakes? The results of Chapter 5 suggest it may depend on the type of winterfreeze lake.  6.2.3 Q3: Moderate PwO2 + long duration  The winterfreeze lakes described above are eutrophic lakes, but oligotrophic lakes freeze during winter as well. Oxygen levels in these lakes are reduced and kept low for the same reasons as in eutrophic winterfreeze lakes, but the lower productivity levels inherent to oligotrophic lakes tend to result in less severe PwO2s being reached by winter’s end (Ultsch, 1989). These environments are therefore typified by hypoxia that is moderate (PwO2) and long (duration). Alta Lake in British Columbia is one such lake, and I can use the data collected in Chapter 5 to explore how this hypoxic environment has shaped the HMR of its resident sticklebacks.  The greater hypoxia tolerance of the Alta Lake sticklebacks compared to the Trout Lake sticklebacks appears to result from their ability to employ MRD at sub-Pcrit PwO2s. This is similar to goldfish. But despite both goldfish and the Alta Lake sticklebacks employing a hypoxia-induced MRD, there are notable differences between their respective MRDs and how they are used. In short, goldfish depress metabolic rate by 80% and wait until near-anoxia to initiate it, while Alta Lake sticklebacks depress metabolic rate by 33% and do so at 2.8 kPa PwO2. These differences in MRD might relate to differences in each species’ hypoxic environment. While the native lakes of goldfish likely become anoxic during wintertime (like the native lakes of crucian carps; Vornanen et al., 2004), apart from at the water-sediment interface (Dunnington et al., 2016), Alta Lake only becomes hypoxic (Jacques  108 Whitford/AXYS, 2007). These hypoxic conditions mean two things for the stickleback inhabitants and their use of MRD. First, while 2.8 kPa PwO2 is not ‘severe’ in the context of the goldfish’s natural anoxic habitat, it probably is in the context of Alta Lake. Therefore, selection may be acting on hypoxic survival strategies at higher PwO2s in the Alta Lake environment than in the more severe goldfish environment, and this could be why Alta Lake sticklebacks induce MRD at a higher PwO2 than goldfish. Second, the presence of O2 makes for a less extreme environment that can support other, less tolerant, species during wintertime, including predatory trout. Salmonid species are adept at locating and exploiting microhabitats in frozen lakes and streams (Brown et al., 2011), and can survive in these habitats so long as they can locate an O2 source. For the salmonids that live in Alta Lake (cutthroat and rainbow trout), some move into the small creeks that drain into the Lake while others stay in the lake itself (Tara Schaufele, Resort Municipality of Whistler’s Environmental Supervisor; personal communication). Because salmonids continue to eat during wintertime (albeit at reduced rates; Klemetsen et al., 2003), the stickleback population remains under some level of predation pressure, and this could possibly limit the degree to which the sticklebacks depress their metabolic rates during wintertime (see Chapter 5 for a discussion on the possibility of hypoxic water layers serving as refugia for sticklebacks). This appears to be the case in the OMZ as well, where species that are more likely to be eaten depress their metabolic rates to lesser extents (e.g., MRD for krill of ~40 to 50%) than species that are less likely to be eaten (e.g., MRD for jumbo squid of ~80 to 90%; Seibel, 2011). And in my experience, it is certainly true that metabolically depressed sticklebacks are more responsive to stimuli than metabolically depressed goldfish, which are largely unresponsive. In any case, regardless of whether the hypoxic (as opposed to anoxic) conditions of Alta Lake explain the Alta Lake stickleback’s relatively modest MRD, it very likely explains their lesser tolerance when compared to goldfish. Another environment that is typified by moderate PwO2s and long (chronic) durations is the OMZ from the perspective of its permanent residents. Permanent residents, which include many fish and invertebrate species, tend to live towards the OMZ’s periphery where PwO2s are not as severe as in its center (Childress and Seibel, 1998). Probably owing to the detrimental effects of long-term reliance on anaerobic glycolysis and/or MRD, these animals rely primarily on aerobic metabolism through a suite of highly effective O2 extraction  109 adaptations. Compared with related species from less hypoxic waters, permanent OMZ residents tend to have enhanced ventilatory abilities, larger respiratory surfaces, thinner blood-water diffusion barriers, and higher affinity respiratory pigments. An example is the giant red mysid (Gnathophausia ingens), a permanent OMZ resident that is particularly well studied. The red mysid has a high ventilatory capacity (Childress, 1971), a large mass-specific gill surface area (Childress, 1971), a small blood-water diffusion distance across the gills (Seibel, 2011), a high circulatory capacity (Belman and Childress, 1976), and a haemocyanin with an extremely high affinity for O2 and a large Bohr effect to facilitate tissue O2 delivery (Sanders and Childress, 1990a; Sanders and Childress, 1990b). Combined with an exceptionally low routine metabolic rate (a common trait of permanent OMZ residents; see Childress, 1995), this results in a Pcrit value of 0.8 kPa (Seibel, 2011), coincident with the minimum PwO2 that the mysid typically encounters in the OMZ (Childress and Seibel, 1998). In fact, across a wide range of OMZ residents, Pcrit has been shown to correlate at near unity with the minimum PwO2s each of these animals experience in the wild (Childress, 1975; Cowles et al., 1991; Donnelly and Torres, 1988; Torres et al., 1994). This finely tuned aerobic capacity may preclude a significant reliance on anaerobic glycolysis, and may subsequently explain why permanent OMZ residents tend to have limited anaerobic abilities (Childress and Seibel, 1998).   6.3  Summary and perspectives  We can combine these seven case studies with the environmental hypoxia matrix in Fig. 6-1 to draw some general conclusions on the HMRs used in different hypoxic environments (Fig. 6-2). Figure 6-2 uses a series of circular pie charts to represent HMR. The diameter of a pie chart is proportional to its metabolic rate; therefore, a smaller-diameter pie chart represents a depressed metabolic rate. The diameters of the pie charts are arbitrary; rather, they are qualitative representations to be compared to the normoxic routine metabolic rate shown in Q1. The black-filled portion of a pie chart represents the aerobic contribution to its metabolic rate, and the grey-filled portion represents the anaerobic contribution. All-black and all-grey therefore represent fully aerobic and anaerobic, respectively. The ratio of aerobic:anaerobic within a single pie chart is arbitrary and meant only to portray that aerobic  110 and anaerobic pathways are simultaneously contributing to total metabolic rate at some approximate ratio.  For short and severe hypoxic exposures (Q2), the HMR varies as a function of aerial/surface access and predators. If the air-water interface is accessible (e.g., tidepools, rice paddies), fishes living in these environments tend to prioritize aerobic metabolism by using ASR and/or air breathing; if an aerial predator presents itself, the fish tend to dive into the hypoxic water and buffer routine metabolic rate using anaerobic glycolysis. If the air-water interface is inaccessible (e.g., OMZ), the animals living in these environments tend to use MRD and save their routine activities such as feeding and mating for their daily migration into oxygenated surface waters. These animals use deep MRD if predation risk is low-to-absent (e.g., jumbo squid), and moderate MRD if predation risk is moderate-to-low (e.g., krill).  For long and moderate hypoxic exposures (Q3), the HMR varies as a function of exposure duration. Species that live in seasonally moderate hypoxia (e.g., oligotrophic winterfreeze lakes) tend to use MRD, perhaps because they can accomplish their routine activities during the oxygenated months of the year. My particular example species (threespine stickleback from Alta Lake) employs a relatively modest MRD with only a small anaerobic contribution, possibly due to the presence of predatory trout. Species that live in chronically moderate hypoxia (e.g., OMZ periphery) tend to rely on enhanced O2 extraction abilities so as to support routine metabolic rate aerobically. Because these species never enter fully oxygenated waters, they need to support routine activities like feeding and mating in chronic hypoxia, and this makes an aerobic metabolism-based HMR ideal.  Finally, for long and severe hypoxic exposures (Q4), the HMR varies as a function of aerial/surface access. If it is accessible (e.g., Amazon basin), then the fishes living in these environments tend to exploit its high O2 content using ASR and/or air breathing, effectively uncoupling themselves from their severely hypoxic aquatic environment. If the air-water interface is inaccessible (e.g., eutrophic winterfreeze lakes), then aerobic metabolism is not an option and the fishes living here tend to employ deep MRD instead so as to conserve anaerobic fuel reserves. A general lack of predators in these environments allows these fishes to surrender locomotor performance with minimal threat of being eaten, while the return of  111 O2 with spring thaw allows them to complete routine activities such as feeding and mating in oxygenated water.  Together, these trends suggest that no matter what the hypoxic environment, fishes use a wide variety of adaptive mechanisms to do what they can to preserve aerobic metabolism. If this becomes impossible (as is the case with Q2 and Q4 hypoxic environments), fishes appear to use one of two approaches depending on the duration of the exposure: anaerobic glycolysis is used to buffer routine metabolic rate if the exposure is short, and MRD is used if the exposure is long. The presence of predators and/or aerial access complicates this, as will co-varying abiotic factors like temperature, PwCO2 and pH. But by and large, there appear to be patterns in the HMRs of unrelated species that depend on their native hypoxic environments. Searching for and understanding these patterns is important. The prevalence and severity of hypoxia among the world’s aquatic environments is increasing (IPCC, 2014; Schmidtko et al., 2017; Smith et al., 2006), and so understanding the hypoxic metabolic responses of fishes will help us to better identify potentially vulnerable species, and better predict the ways they may redistribute themselves in an increasingly hypoxic world. 112 Figure 19   Figure 6-1. A matrix of environmental hypoxia portraying various natural hypoxic environments according to their hypoxic severities (PwO2) and hypoxic durations (time). Quadrant (Q) 1 portrays moderate (PwO2) + short (duration) environments, Q2 portrays severe + short environments, Q3 portrays moderate + long environments, and Q4 portrays severe + long environments. The environments listed within each quadrant will vary from one another in severity and duration. For the sake of simplicity, rate of hypoxia induction (RHI) is incorporated along the time duration axis whereby short-duration environments tend to experience rapid RHIs, and long-duration environments tend to experience gradual RHIs. See text for details.Figure 6-1. An environmental hypoxia matrix that portrays various natural hypoxic environments accord ng to th ir hypoxic severities (PwO2) and hypoxic durations (time). Quadrant (Q) 1 portrays moderate + short environments, Q2 portrays severe + short environments, Q3 portrays moderate + long environments, and Q4 portrays severe + long environments. The environments listed within each quadrant will vary from one another in severity and duration. See text for details. !"#$%&'()*+,-$.(/0!12(!"#$%&'(3 456(78$+9( :$.;(7<=<+<(>$)<+,9<(!"# !$#!%# !&#'()*+#,-.*+/012#3((24#52,6(.+(37,8#),-.*+9+**:*#21;*4#5<=#3*+,37*+>#?@A18A2.A+*#3(-04#B,67*+#,-.*+/012#3((24#C,8*#3100,*4#D12.#E1+47*4#5<=#8*-.*+#F+(3,812#+,G*+4#HA.+(37,8#),-.*+9+**:*#21;*4#I,*2#E,6+1.(+>#5<=#/+,#&)2( /;+,)*,:2( 113 Figure 20                 Figure 6-2. The hypoxic metabolic responses (HMRs) of species adapted to natural aquatic hypoxic environments listed in Fig. 6-1 and detailed in the text. The circular pie charts represent HMR and are built according to the multi-species analyses described in the text. Each pie chart’s diameter is a qualitative representation of its total metabolic rate relative to the size of the pie chart in Q1 representing normoxic routine metabolic rate. Therefore, a smaller-diameter pie chart than that shown in Q1 represents a depressed metabolic rate. The black-filled portion of a pie chart represents the aerobic contribution to that HMR’s metabolic rate, and the grey-filled portion represents the anaerobic contribution. The ratios of aerobic:anaerobic are estimations. The low hypoxic magnitudes of the Q1 habitats precluded their analysis (indicated by diagonal grey lines).    !"#$%#&''())*#!"+#$%##&''())*#!"#$%&'()*%+#!"+#$%##&''())*#+&(+*,(-#&.$*+/%&#01%*,20#&.$*+/%&#,-# ,.#,%# ,/#!"*##$%&'()*%+#!"+#$%#&''())*#!"#$%#&''())*#!"#$%&'()*%+#!"*##$%&'()*%+#+&3&%&#4*'&%()&#Figure 6-2. The hypoxic metabolic responses (HMRs) of species adapted to natural aquatic hypoxic environments listed in Fig. 6-1 and detailed in the text. The circular pie charts represent HMR. Each pie chart’s total area is a qualitative representation of its total metabolic rate relative to the size of the pie chart in Q1 representing normoxic routine metabolic rate. The black-filled portion of a pie chart represents the aerobic contribution to that HMR’s metabolic rate, and the grey-filled portion represents the anaerobic contribution. The ratios of aerobic:anaerobic are estimations. The low hypoxic magnitudes of the Q1 habitats pr cluded their analysis (indicated by diagonal grey lines). Th  pie charts depicted in Q2, Q3 and Q4 were built according to the analyses described in the text. 56$*.20#'/%(7*,#895:;#8%($2';# 8<%('/(-;#+1*%)# -*,<#56$*.20#= !>?# 114 REFERENCES  Abdallah, S. J., Thomas, B. S. and Jonz, M. G. (2015). Aquatic surface respiration and swimming behaviour in adult and developing zebrafish exposed to hypoxia. Journal of Experimental Biology 218, 1777–1786. Addink, A., van den Thillart, G., Smit, H. and van Waversveld, J. (1991). A novel 1 liter flow-through calorimeter for heat production measurements on aquatic animals without stress. Thermochimica Acta 193, 41–48. Affonso, E. G., Polez, V. P., Correa, C. F., Mazon, A. 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