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Mechanisms of cardiac pacemaking and temperature-dependent depression of cardiac electrical excitation… Marchant, James 2021

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MECHANISMS OF CARDIAC PACEMAKING AND TEMPERATURE-DEPENDENT DEPRESSION OF CARDIAC ELECTRICAL EXCITATION IN THE ZEBRAFISH  (DANIO RERIO) by  James Marchant  B.Sc., The University of Western Brittany, 2013 M.Sc., The University of Western Brittany, 2015  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)  April 2021  © James Marchant, 2021  ii  The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled:  Mechanisms of cardiac pacemaking and temperature-dependent depression of cardiac electrical excitation in the zebrafish (danio rerio)  submitted by James Marchant in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Zoology  Examining Committee: Dr. Anthony Farrell, Professor, Zoology, UBC Supervisor  Dr. Patricia Schulte, Professor, UBC Supervisory Committee Member  Terrance Snutch, Professor, UBC University Examiner Eric Accili, Associate Professor, UBC University Examiner  Additional Supervisory Committee Members: Dr. Holly Shiels, Associate Professor, The University of Manchester Supervisory Committee Member Dr. Frank Smith, Associate Professor, Dalhousie University Supervisory Committee Member  iii  Abstract  The physiology of ectotherms is profoundly affected by the environmental temperature which governs the rate of physiological processes. Cardiac function, an essential function of all vertebrates, is no exception: during warming heart rate tracks temperature before declining at temperatures beyond maximum optimal temperature, ultimately collapsing with further warming. In fishes, like other vertebrates, intrinsic heart rate is set by pacemaker cells located in the sino-atrial node that spontaneously generate action potentials. At the cellular level, temperature-dependent deterioration of pacemaking mechanisms may contribute to the decline of cardiac function. Nevertheless, the mechanisms of pacemaking and their temperature-dependent deterioration remain elusive. Hence, I explored cardiac pacemaking mechanisms in zebrafish, as well as their relative thermal performance and limits. I validated blebbistatin as an effective excitation-contraction uncoupling agent that did not modify the cardiac action potential properties, thus providing an essential methodology for future cardiac pacemaking research enabling the direct recording of intracellular electrical activity of pacemaker cells.   Using electrocardiograms, I confirmed that cardiac pacemaking involves two major mechanisms. Pharmacological blockade of hyperpolarization-activated cyclic nucleotide-gated (HCN) channels with zatebradine reduced heart rate by up to 60%, suggesting HCN channels play the major role in cardiac pacemaking. Likewise, sarcoplasmic reticulum (SR) calcium cycling was pharmacologically blocked using ryanodine and thapsigargin to block ryanodine receptors and SERCA pumps, respectively, which reduced heart rate by ~40%, suggesting the SR plays a secondarily important role in pacemaking. However, the combination of these pharmacological iv  interventions did not completely stop the heartbeat, suggesting that either mammalian pharmacological agents are less effective in producing total Hcn block in zebrafish, perhaps due to isoform specificity. HCN4, the major HCN channel involved in mammalian pacemaking, was knocked out using CRISPR to explore its role in zebrafish cardiac pacemaking. Heart rate did not differ significantly between mutant and control fish at any test temperature, including fish treated with inhibitors of HCN channels or SR calcium cycling. Thus, alternative Hcn channels compensated for the knockout of Hcn4, presumably contributing to a higher thermal tolerance. In addition, mutant fish had a higher upper thermal tolerance than control fish when SR calcium cycling was inhibited.   v  Lay Summary  The heartbeat is generated by specialized cells in the heart. My research investigated the mechanisms that allow these cells to drive the spontaneous electrical activity behind every heartbeat. I identified two mechanisms involved in cardiac pacemaking and assessed their temperature tolerance in the zebrafish heart. I generated mutant zebrafish, deficient of an important pacemaking gene (Hcn4) for one of these mechanisms, and found no difference in heart rate compared with non-mutant fish, suggesting that fish can cope without Hcn4 across a wide range of temperatures. I also demonstrate proof-of-principal for a method to record electrical activity directly in pacemaker cells.  vi  Preface  A version of chapter 2 has previously been published as Marchant, J. L. and Farrell, A. P. (2019). Membrane and calcium clock mechanisms contribute variably as a function of temperature to setting cardiac pacemaker rate in zebrafish Danio rerio. J. Fish Biol. 95, 1265–1274.  I was the primary contributor to the experimental design. I carried out the collection and data analysis. Frank Smith contributed to the selection of the pharmacological agents used. I prepared the manuscript and Anthony Farrell provided edits. Procedures were approved by the animal care committee of The University of British Columbia under the animal care application A18-0003. For all experiments in chapter 3, I was the primary contributor to the experimental design and data collection. I established the zebrafish knockout line at the University of Manchester under the supervision of Adam Hurlstone and Holly Shiels. Andrew Badrock provided many useful suggestions for the development of the mutant zebrafish line. I was the principal contributor to the conception of the original idea, the principal contributor to the experimental design, data collection, data analysis and manuscript preparation. All procedures adhered to the United Kingdom Home Office Animals Scientific Procedures Act of 1986 and were granted authority under project license P005EF9F9 and 70/9091. Daniel Ripley assisted with the perpetrations of the respirometry experiments, providing the respirometry setup and training. Daniel Ripely performed some of the preliminary respirometry experiments and analyzed the respirometry data. Holly Shiels and Adam Hurlstone provided lab space and equipment at the University of Manchester. Holly Shiels, Andrew Badrock and Anthony Farrell provided feedback on the manuscript. A version of chapter 3 is in perparation for publication.   vii  Data collection for chapter 4 was carried out at the University of British Columbia and at Dalhousie University. The original idea and experimental design were provided by Frank Smith, including the choice and concentration of the pharmacological agent used. I was the primary contributor to data collection and data analysis. Procedures were approved by the institutional animal research authority of The University of British Columbia animal care committee under the animal care application number A18-0014, or by the institutional animal care committee of Dalhousie University under the Ethics Protocol number 15-006. While the data chapters involved several contributors, chapters 1 & 5 are of my own composition, with general input from my thesis supervisor and committee members. All figures without a cited source are of my own original design.   viii  Table of Contents  Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ....................................................................................................................... viii List of Tables ................................................................................................................................xv List of Figures ............................................................................................................................. xvi List of Abbreviations ...................................................................................................................xx Acknowledgements .................................................................................................................. xxiv Dedication ................................................................................................................................. xxvi  Introduction ................................................................................................................1 1.1 Preface............................................................................................................................. 1 1.2 Temperature-dependent excitation and excitation-contraction of the fish heart ............ 3  Thermal acclimation of fishes ................................................................................. 5 1.3 Electrical excitation of the fish heart .............................................................................. 6  The action potential of the working myocardium ................................................... 9  Cardiac pacemaking in the sinoatrial node ........................................................... 13 1.3.2.1 The membrane clock ......................................................................................... 14 1.3.2.1.1 Hyperpolarization-activated cyclic nucleotide-gated (HCN) channels ...... 16 1.3.2.2 The calcium clock ............................................................................................. 20 1.3.2.3 The coupled clock hypothesis ........................................................................... 21 1.4 Pacemaking in fish: role of the membrane and calcium clocks .................................... 24 ix  1.5 Temperature-dependent deterioration of cardiac pacemaking in fish ........................... 30 1.6 The use of zebrafish as a model for molecular and electrophysiological pacemaker research ..................................................................................................................................... 32 1.7 Thesis relevance and aims ............................................................................................ 33  Chapter 2: Thermal limits of the zebrafish heart and contribution of membrane and calcium clock to cardiac pacemaking (Marchant and Farrell, 2019). ............................ 33  Chapter 3: The role of hcn4 in pacemaking of the zebrafish heart and the thermal tolerance of the hcn4 mutant. Physiological characterization of a hcn4 mutant zebrafish. .. 34  Chapter 4: The effect of blebbistatin-mediated uncoupling of excitation-contraction on the cardiac action potential in zebrafish........................................................ 34  Thermal limits of the zebrafish heart and contribution of membrane and calcium clock to cardiac pacemaking .........................................................................................35 2.1 Synopsis ........................................................................................................................ 35 2.2 Introduction ................................................................................................................... 36 2.3 Materials and methods .................................................................................................. 40  Animals and husbandry......................................................................................... 40  Acclimation procedure .......................................................................................... 40  Fish preparation and acute warming protocol ....................................................... 41  Pharmacological agents and solutions .................................................................. 43  Data analysis and statistics.................................................................................... 44 2.4 Results ........................................................................................................................... 46  General effects of acute warming on fHmax ............................................................ 46  The effect of temperature acclimation on fHmax..................................................... 49 x   Effect of Hcn channel blocking with zatebradine ................................................. 54  Effect of SR calcium cycling disruption using ryanodine and thapsigargin ......... 54  Effect of Hcn channel blocking combined with SR calcium cycling disruption .. 58 2.5 Discussion ..................................................................................................................... 60  Extreme acclimation temperatures reduce thermal tolerance ............................... 60  Effect of zatebradine on fHmax ............................................................................... 63  Effect of ryanodine and thapsigargin on fHmax ...................................................... 65  Effect of zatebradine, ryanodine, and thapsigargin on fHmax ................................. 67  Conclusion ............................................................................................................ 67  Limitations and future directions .......................................................................... 68  The role of Hcn4 in pacemaking of the zebrafish heart and the thermal tolerance of the hcn4 mutant - Physiological characterization of a hcn4 knockout zebrafish.........................................................................................................................................................70 3.1 Synopsis ........................................................................................................................ 70 3.2 Introduction ................................................................................................................... 71 3.3 Materials and methods .................................................................................................. 76  Animal husbandry ................................................................................................. 76  Breeding and rearing of embryos .......................................................................... 76  sgRNA design ....................................................................................................... 77  sgRNA synthesis ................................................................................................... 78  Generation of a hcn4 mutant zebrafish line .......................................................... 79 3.3.5.1 Construct injection in one-cell stage zebrafish embryos .................................. 79 3.3.5.2 Identification of INDELS in founder generation (F0) fish ............................... 79 xi  3.3.5.3 F0 outcross and first-generation (F1) incross ................................................... 80  Knockout validation by Western blot ................................................................... 82  Real-time quantitative PCR (RT-qPCR) ............................................................... 83  Critical thermal maximum (CTmax) trials .............................................................. 85  Swimming trials .................................................................................................... 86  Respirometry ......................................................................................................... 87  Electrocardiogram (ECG) recordings ................................................................... 90  Statistical methods ................................................................................................ 92 3.4 Results ........................................................................................................................... 93  Introduction of a premature stop codon into zebrafish hcn4 abrogates protein expression ............................................................................................................................. 93  Identification of INDELS in founder generation (F0) fish ................................... 95  F0 outcross and first-generation (F1) incross of the mutant zebrafish ................. 96  Generation of homozygous hcn4 knockout zebrafish in F2 ................................. 97  Validation of hcn4 knockout zebrafish line – Western blot and RT-qPCR ........ 100  Hcn4 knockout zebrafish are indistinguishable from wild-type fish .................. 101  Loss-of-function of hcn4 mutant increases critical thermal maximum (CTmax) resulting in heat-tolerant zebrafish...................................................................................... 102  hcn4 mutant zebrafish maintained maximum respiratory capacity and swimming performance ........................................................................................................................ 104  Hcn4 mutant fish maintained heart rate during acute warming .......................... 106  Effect of pharmacological blockade of Hcn channels and SR-Ca2+ cycling on fHmax ............................................................................................................................. 109 xii  3.5 Discussion ................................................................................................................... 113  Cardiac pacemaking is maintained in hcn4 mutant fish ..................................... 113  Hcn4 knockout induced no change to the calcium clock .................................... 114  The effect of Hcn4 knockout on the membrane clock ........................................ 114  Alternative Hcn channels have a more prominent role in hcn4 mutant knockout zebrafish   ............................................................................................................................ 116  Hcn4 knockout induced no change to whole-animal aerobic respiration and swimming performance ...................................................................................................... 120  Role of Hcn4 in cardiac thermal tolerance of the zebrafish ................................ 121  Limitations and conclusions ............................................................................... 122  The effect of blebbistatin on the pacemaker action potential ............................126 4.1 Synopsis ...................................................................................................................... 126 4.2 Introduction ................................................................................................................. 127 4.3 Materials and methods ................................................................................................ 131  Animals ............................................................................................................... 131  Heart isolation and tissue preparation ................................................................. 131  Recording of the action potentials ...................................................................... 133  Data analysis ....................................................................................................... 134  Statistical analysis ............................................................................................... 138 4.4 Results ......................................................................................................................... 139  The effect of blebbistatin on the pacemaker action potential characteristics from the SAN region of zebrafish ............................................................................................... 139 xiii   The effect of temperature on the pacemaker action potential with and without blebbistatin in the UBC group ............................................................................................ 145 4.4.3. The effect of blebbistatin on action potential voltage and duration.......................... 157 4.5 Discussion ................................................................................................................... 160 4.6 Limitations and perspectives....................................................................................... 165 4.7 Conclusion .................................................................................................................. 166  General discussion and future directions .............................................................168 5.1 Synopsis ...................................................................................................................... 168 5.2 Thermal acclimation and resetting of the heart rate.................................................... 170 5.3 Mechanisms of cardiac pacemaking ........................................................................... 172  The membrane clock ........................................................................................... 172  The calcium clock ............................................................................................... 174  A coupled clock mechanism ............................................................................... 176 5.4 The importance of Hcn4 in cardiac pacemaking of the zebrafish .............................. 177 5.5 Thermal tolerance of the zebrafish pacemaker and temperature-dependent deterioration of the cardiac electrical signal................................................................................................. 179 5.6 Summary ..................................................................................................................... 183 Bibliography ...............................................................................................................................184 Appendix .....................................................................................................................................229 Appendix 2.1. ECG recordings from a 23°C-acclimated fish incrementally heated from 18°C to 34.4°C. (A) The ECG recording at 18°C (B) the ECG recording at 34.4°C. Red arrows indicate missing QRS complexes............................................................................................ 229 Appendix 2.2. .......................................................................................................................... 230 xiv  Appendix 3.1. Primers used for sgRNA synthesis. ................................................................. 231 Appendix 3.2. Computational model of wild type and mutant truncated form of hcn4 as mediated by CRISPR-Cas9 insertion of a premature stop codon in position 433-435 (TAA).................................................................................................................................................. 232 Appendix 3.4. The weight and CTmax of individual wild-type and hcn4 knockout fish. ........ 234 Appendix 3.5. The percent similarity (%) between the zebrafish and human cDNA sequences and protein sequences for different HCN isoforms. ............................................................... 235 Appendix 3.6. Sequence similarity between the protein sequences of zebrafish Hcn4 and Hcn4l. ...................................................................................................................................... 236 Appendix 4.1. The effect of blebbistatin on action potential duration 50% and action potential duration 80% recorded in reduced heart preparations at room temperature ........................... 238 Appendix 4.3. The effect of blebbistatin and temperature on the action potential threshold potential and the overshoot potential. ..................................................................................... 239 Appendix 4.4. The effect of blebbistatin and temperature on AP amplitude and the maximum hyperpolarization potential. .................................................................................................... 240 Appendix 4.5. A linear regression analysis of the action potential duration 50% and 80%. .. 241 Appendix 4.6. Principal component analysis of voltage-dependent action potential parameters of control (blue circle) and blebbistatin-treated (green diamond) heart preparations recorded at room temperature .................................................................................................................... 242 Appendix 4.7. Principal component analysis of time-dependent action potential parameters of control and blebbistatin-treated  heart preparations recorded at room temperature ............... 243 Appendix 4.8. Sample power plot of required effect size as performed by G*Power 3.1. .... 244  xv  List of Tables  Table 1.1. Activation profiles and cAMP sensitivity of mammalian HCN channels .................. 27 Table 2.1. A comparison of the cardiac performance variables for zebrafish.. ........................... 47 Table 3.1. A list of primers used for generating sgRNA (sgRNA primers), and PCR primers ... 78 Table 3.2. A list of primer pairs used for qPCR. ENSDARG identifier numbers are listed for each of the genes ........................................................................................................................... 84 Table 3.3. A comparison of cardiorespiratory and swimming performance of wild-type and hcn4 mutant zebrafish. ......................................................................................................................... 108 Table 4.1. Table of the number of fish used and action potentials recorded for each treatment and temperature grouping. ................................................................................................................. 136 Table 4.2. Table of action potential parameters with and without blebbistatin in the DU group recorded at room temperature (20°C-23°C) ................................................................................ 143 Table 4.3. A table of all action potential parameters of zebrafish pacemaker cells recorded at room temperature. ....................................................................................................................... 143 Table 4.4. Summary table of the effects of temperature on each of the AP parameters at different temperatures in group UBC. ....................................................................................................... 154 Table 4.5. Pairwise comparison of control and blebbistatin-treated cells with increasing temperature in group UBC. ......................................................................................................... 154     xvi  List of Figures  Figure 1.1. Scheme of the major ion channels generating the action potential and calcium signalling. ........................................................................................................................................ 9 Figure 1.2. Typical action potential (AP) of fish cardiomyocytes. .............................................. 12 Figure 1.3. Schematic of the respirometry Schematic representation of the components of the membrane clock. ........................................................................................................................... 16 Figure 1.4. A functional scheme of the membrane clock and calcium clock mechanisms. ........ 19 Figure 1.5. Schematic of the respirometry Schematic representation of the components of the calcium clock. ............................................................................................................................... 21 Figure 1.6. Schematic of the respirometry Schematic representation of the components of the coupled clock system. ................................................................................................................... 23 Figure 2.1. Effect of acute warming to 33°C on the mean fHmax (± SEM) of anesthetized zebrafish acclimated to 18°C, 23°C, and 28°C ............................................................................. 50 Figure 2.2. Individual responses to acute warming in 1°C increment of anesthetized zebrafish acclimated to either 18°C, 23°C, or  28°C .................................................................................... 52 Figure 2.3. Mean incremental Q10 and Arrhenius breakpoint plot of acclimated zebrafish ........ 53 Figure 2.4.  A comparison of the control responses of fHmax during acute warming in 1°C increments of anesthetized zebrafish acclimated to either 18°C, 23°C ,or 28°C with those following pre-treatment with either zatebradine (4 μg.g-1) or ryanodine (50ng.g-1) and thapsigargin (1.3ug.g-1). ................................................................................................................ 56 Figure 2.5.  The mean percent change in fHmax of zebrafish treated with either zatebradine or ryanodine and thapsigargin ........................................................................................................... 57 xvii  Figure 2.6.    Individual responses of fHmax of anesthetized zebrafish acclimated to 28°C and tested at 28°C. ............................................................................................................................... 59 Figure 3.1. Schematic overview of the CRISPR Cas9 workflow for the generation of a hcn4 knockout zebrafish. ....................................................................................................................... 81 Figure 3.2. Schematic of the respirometry setup. ........................................................................ 89 Figure 3.3. Schematic diagram of CRISPR hcn4 knockout in zebrafish. .................................... 94 Figure 3.4. Agarose gel electrophoresis of DNA isolated from fin tissue of eight-week-old zebrafish. ....................................................................................................................................... 95 Figure 3.5.  Agarose gel electrophoresis of DNA isolated from fin tissue of zebrafish and Sanger sequencing pherogram. ................................................................................................................. 96 Figure 3.6. Validation of hcn4 knockout zebrafish line. .............................................................. 98 Figure 3.7. Validation of hcn4 knockout in zebrafish. ............................................................... 100 Figure 3.8. Morphological appearance and ventricular mass of wild type and hcn4 mutant zebrafish. ..................................................................................................................................... 101 Figure 3.9. Critical thermal maximum (CTmax) of wild-type and hcn4 mutant zebrafish. ........ 103 Figure 3.10. Comparison of swimming performance and respirometry of wild-type and hcn4 mutant zebrafish at 28°C ............................................................................................................. 105 Figure 3.11. Comparison of maximum heart rate (fHmax) of anesthetized wild-type and hcn4 knockout fish ............................................................................................................................... 107 Figure 3.12. A comparison of maximum heart rate (fHmax) in wild-type and hcn4 mutant anesthetized zebrafish during acute 1°C incremental warming. ................................................. 111 Figure 4.1. Schematic diagram of the sagittal view of the zebrafish heart dissection. .............. 132 xviii  Figure 4.2. A schematic protocol of action potential parameter extraction from the raw traces recorded from pacemaker cells. .................................................................................................. 137 Figure 4.3. Overlay of representative action potentials recorded by intracellular microelectrode in reduced zebrafish heart preparations under control conditions and with 10 μM blebbistatin.  ..................................................................................................................................................... 141 Figure 4.4. The effect of blebbistatin on heart rate and action potential period of control and blebbistatin treated cells .............................................................................................................. 141 Figure 4.5.  The effect of blebbistatin on action potential parameters recorded at room temperature from zebrafish heart preparations. .......................................................................... 142 Figure 4.6. Representative action potentials recorded by intracellular microelectrode in a reduced zebrafish heart preparation from different SAN cells in the presence of  10 μM blebbistatin. ................................................................................................................................. 145 Figure 4.7. The effect of blebbistatin and temperature on the heart rate and beat-to-beat period...................................................................................................................................................... 146 Figure 4.8. The effect of blebbistatin and temperature on the diastolic depolarization duration, diastolic amplitude, and the rate of diastolic depolarization. ...................................................... 148 Figure 4.9. The effect of blebbistatin and temperature on the action potential depolarization voltage, action potential depolarization duration, and the action potential duration .................. 151 Figure 4.10. The effect of blebbistatin and temperature on the rate of repolarization and action potential repolarization duration ................................................................................................. 153 Figure 4.11.  Principal component analysis of time-dependent action potential parameters of control and blebbistatin-treated heart preparations recorded at room temperature (20°C-23°C)...................................................................................................................................................... 158 xix  Figure 4.12. Principal component analysis of voltage-dependent action potential parameters of control and blebbistatin-treated heart preparations recorded at room temperature (20°C-23°C)...................................................................................................................................................... 159  xx  List of Abbreviations %: Percent °C: Degrees Celsius  μg: Microgram μL: Microliter  μM: Micromolar  AAS: Absolute aerobic scope ADP: Adenosine diphosphate  ANCOVA: Analysis of covariance ANOVA: Analysis of variance AP: Action potential ATPase: Adenosine triphosphatase BPM: Beats per minute Ca2+: Calcium (ion) [Ca2+]i: Intracellular calcium concentration cAMP: Cyclic adenosine monophosphate Cas9: CRISPR-associated protein 9  cDNA: Complementary deoxyribonucleic acid CHO: Chinese hamster ovary  CI: Confidence interval CICR: Calcium-induced calcium-release CRISPR: Clustered regularly interspaced short palindromic repeats CRISPRsg: CRISPR single guide crRNA: CRISPR RNA Ct: Cycle threshold for qPCR CTmax: Critical thermal maximum Cx43: Connexin 43 DNA: Deoxyribonucleic acid DMSO: dimethyl sulfoxide  Dpf: days post-fertilization ECF: extracellular fluid ECG: Electrocardiogram  xxi  EGFP: Enhanced green fluorescent protein fH: Heart rate fHmax: Maximum heart rate g: Gram GFP: Green fluorescent protein h: hour HCN: Hyperpolarization-activated cyclic nucleotide-gated channel HCN 1 channel: Hyperpolarization-activated cyclic nucleotide-gated channel 1 HCN 2 channel: Hyperpolarization-activated cyclic nucleotide-gated channel 2 HCN 3 channel: Hyperpolarization-activated cyclic nucleotide-gated channel 3 HCN 4 channel: Hyperpolarization-activated cyclic nucleotide-gated channel 4 Hpf: hours post-fertilization ICa: Calcium current  If: Funny current  IK: delayed rectifier potassium current  IK1: Inward rectifier current  IKACH: Acetylcholine-activated inward rectifier current  IKr: Rapidly activating delayed rectifier potassium current IKs: Slowly activating delayed rectifier potassium current INa: sodium current  INDEL: Insertions or deletions Ito: Transient outward potassium current K: Kelvin K+: Potassium (ion) kHz: Kilohertz Ln: Natural logarithm MΩ: Mega Ohm (electrical resistance)  Min: Minute  mL: Milliliter mM: Millimolar MMR: Maximum metabolic rate ṀO2: Rate of oxygen uptake  mRNA: messenger ribonucleic acid  xxii  ms: Millisecond MS-222: Tricaine methanesulfonate mV: Millivolt  n: sample size number Na+: Sodium (ion) NCX: Sodium-calcium exchanger NHEJ: Non-homologous end joining PAM: protospacer adjacent motif PCA: Principal component analysis PCR: Polymerase chain reaction pg: picogram Pi: Inorganic phosphate pL: picoliter P-value: Probability value RMP: Resting membrane potential RNA: Ribonucleic acid RT: Room temperature RT-qPCR: Real-time quantitative PCR RyR: Ryanodine receptor s: Second SAN: Sinoatrial node SD: Standard deviation SEM: Standard error of the mean SERCA: Sarcoplasmic/endoplasmic reticulum calcium ATPase  sgRNA: Single guide RNA SL: Sarcolemma SMR: Standard metabolic rate SR: Sarcoplasmic reticulum Tab: Arrhenius breakpoint temperature Talt: Temperature of first cardiac alternan Tarr: Temperature of first cardiac arrhythmia Tmax: Maximum temperature TQRS: Temperature of the first missing QRS complex xxiii  Ucrit: Maximal critical swimming speed VG: Voltage gated xxiv  Acknowledgements  I would like to thank my supervisor, Anthony Farrell for giving me the opportunity to pursue my Ph.D. and for providing me a lot of freedom in my research. I would like to thank my committee members, Patricia Schulte, Holly Shiels, and Frank Smith for their helpful input, support, and advice throughout my thesis.   I would like to thank Frank Smith for hosting me at Dalhousie University and for showing amazing hospitality during my visit.    I would like to extend my deepest gratitude to Holly Shiels who always made me feel welcome during my research visits at the University of Manchester and always provided me with inspirational mentorship, helpful insight, and friendship throughout my thesis. I would like to thank Alexander Holsgrove, Daniel Ripely, Samantha Hook, Bridget Evans, Ilan Ruhr, Aineura Martins, Charlotte Marris, Sana Yaar, Miriam Thavarjah, Syafiq Musa, Pierre Delaroche, and Shiva Nag Kompella, for everything during my visits to the University of Manchester. In addition, I would like to thank Adam Hurlstone for graciously accepting to supervise me for one of the major chapters of this thesis. I would like to especially thank Andrew Badrock, Federica Bottiglione, and Gemma Davis of the Hurlstone lab for answering my incessant questions and for providing unwavering support and guidance. I would also like to thank all of the other members of the Hurlstone lab, and others of the Michal Smith building for making my visit to the University of Manchester an enjoyable experience.  Between these two labs, my visits to the University of Manchester were made enjoyable and were a major highlight of my Ph.D.  xxv  I would also like to extend my deepest gratitude to Agnès Lacombe and Vivienne Lam for helping me to develop my passion for teaching and for providing an enjoyable teaching environment for me to develop. My UBC teaching experience has been amazing from start to finish, and the guidance and support received from you both helped me immensely in developing my teaching skills.    I would like to thank my supervisor for funding my Ph.D. work through grants awarded to him by NSERC.  I would like to thank my parents and grandparents for their unwavering support and for always reminding me of my achievements despite multiple hurdles that oftentimes felt like mountains.  I save my last acknowledgment for my husband, Christos, who has been my greatest support throughout my thesis. You always knew what to say when I was down, and always knew how to provide the support I needed, when I needed it. You never let me doubt myself, even when my confidence was at its lowest, and you always believed in me. I love you for everything you have done to support and encourage me during my Ph.D., and I hope that our next septs will be filled with adventure and surprise.  xxvi  Dedication  I dedicate this thesis to my parents who taught me to pursue my dreams in the quest of happiness.  1   Introduction 1.1 Preface Regardless of the species considered, the primary function of the heart — pumping blood through the body, transporting respiratory gases and metabolic substrates —is conserved among all vertebrates (Farrell and Jones, 1992). Specialized cardiac pacemaker cells are responsible for the initiation of every heartbeat and thus support the essential function of cardiac pacemaking, which is conserved among all vertebrates. The heartbeat would not occur without the spontaneous electrical excitation of pacemaker cells, which are central to the heart's response to temperature and are directly affected by temperature; increasing temperature directly increases the action potential firing rate (Farrell and Smith, 2017; Haverinen and Vornanen, 2007; Randall, 1970). Therefore, the underlying pacemaking mechanisms are central components of the temperature response of the fish heart and, indeed, of the whole animal, to temperature fluctuations (Farrell et al., 2009; Haverinen and Vornanen, 2007).  For ectothermic animals such as fishes, where the body temperature parallels the environmental temperature, maintaining sufficient cardiac output to meet oxygen demands can become problematic under varying environmental conditions, notably, at extremes of temperature. Fish increase heart rate (fH) in response to increasing temperature with a functional ratio over a 10°C change in temperature (Q10) of 2, until a point of no return where fH can only decline with further warming (Casselman et al., 2012; Marchant and Farrell, 2019; Safi et al., 2019; Sidhu et al., 2014). Therefore, heat tolerance of cardiac function has been suggested to be a limiting factor 2  for whole animal upper thermal tolerance limits of fish (Badr et al., 2018; Eliason et al., 2011; Gollock et al., 2006; Vornanen, 2016).    Although there is no consensus within the scientific community, the cause of the temperature-dependent decline in cardiac performance may be caused by a decline in contractile capacity of the myofilaments, insufficient oxygen delivery to the heart, or caused by the deterioration of electrical excitation and signalling of the heart (Farrell, 2009; Haverinen and Vornanen, 2020a; Vornanen, 2016; Vornanen, 2020). Furthermore, temperature-dependent deterioration of electrical excitation could be caused by propagation failure of electrical excitation or by the failure of impulse generation in cardiac pacemaker cells at high temperatures (Vornanen, 2016; Vornanen, 2020). Despite the essential role of the heart in thermal responses of cardiac output, the ionic mechanisms of cardiac pacemaking, and the influence of temperature on pacemaking mechanisms remain poorly understood in fish, including their temperature-dependent deterioration (Vornanen, 2016).  Given the vital importance of the heart and the threat posed to cardiac function by global warming, I chose to study the temperature-dependent depression of cardiac electrical excitation to further our understanding of the effect of temperature, notably extreme high temperatures on the mechanisms generating the cardiac action potential. Fisheries and aquaculture represent a multibillion-dollar industry and an essential food source for human populations (Balami et al., 2019; Tacon and Metian, 2013). However, global warming endangers wild population and cultured fish (Hoegh-guldberg, 2010; Pörtner et al., 2010) as physiological performance is negatively impacted beyond the upper thermal limit of the fish (Sandblom et al., 2016) but phenotypic plasticity may allow fish to adapt to the new environmental temperatures (Seebacher et al., 2015; Somero, 2010). Deterioration of cardiac performance at high temperature has previously been 3  recorded, but the mechanisms underpinning the decline in physiological performance remain unknown, and the mechanism by which the decline in heart rate may occur is debated within the field but may originate from either impulse propagation of the action potential or impulse generation at the level of the pacemaker cells. Thus, better understanding the mechanisms of temperature-dependent depression of cardiac function furthers our understanding of cardiac thermal tolerance and may be useful in informing environmental conservation efforts and aquaculture. I specifically chose the zebrafish for a number of reasons, the first being its relevance as a model for human cardiac physiology, electrophysiology and cardiac pathology. As the mechanisms of cardiac pacemaking in the zebrafish are not fully understood, nor their thermal preference and tolerance, current research may not be optimized. Furthermore, the acclimation capacity and thus the timeframe within which any acclimation occurs is also understudied in the zebrafish which may also impact isolated heart, tissue and cell studies. The zebrafish is also amenable to genetic engineering technologies allowing investigation of the role of different proteins, essential for dissecting complex mechanisms such as cardiac pacemaking.    1.2 Temperature-dependent excitation and excitation-contraction of the fish heart Temperature governs the rates of biological and physiological processes and particularly affects the performance and fitness of ectotherms, including fish (Hochachka and Somero, 2002). It has been termed the ecological master factor as the effects of temperature manifest at every level of biological organization, from molecular interactions and stability of DNA duplexes and proteins to whole-animal metabolic rates and population distributions (Brett, 1971; Dickson and Graham, 2004; Shelford, 1931). Elevated temperature exponentially increases the rate of physiological 4  processes and biochemical reactions, including the gating properties of ion channels, pumps, and exchangers (Hille, 2001). Increasing temperature, for example, increases the rate of ion channel gating properties as the conformation changes to the channel for opening are temperature sensitive, accelerating changes in electrochemical gradients and membrane potential, even though conductance sees little increase with temperature (Hille, 2001).  A central component in a fish’s response to temperature is the temperature-dependent increase in fH, a response observed in all fish studied to date (Clark et al., 2008; Clark et al., 2011; Drost et al., 2014; Eliason et al., 2011; Ferreira et al., 2014; Gilbert et al., 2019; Gollock et al., 2006; Harper et al., 1995; Haverinen and Vornanen, 2007; Heath and Hughes, 1973; Lin et al., 2014; Safi et al., 2019; Sidhu et al., 2014; Steinhausen et al., 2008; Vornanen, 2016; Vornanen et al., 2014). Increasing fH is the fish’s primary cardiac response to increasing temperature because stroke volume changes little (Mendonça and Gamperl, 2010; Steinhausen et al., 2008). The temperature-dependent increase in fH is mediated by the acceleration of pacemaker action potential (AP) generation, which increases the firing rate of cardiac pacemaker cells (Farrell and Smith, 2017; Haverinen and Vornanen, 2007; Randall, 1970). At temperatures below those acutely lethal to fish (Critical thermal maximum, CTmax), fH ceases to increase, capping the oxygen supply to tissues, even though tissue demand continues increasing with temperature (Eliason et al., 2011; Fry, 1947; Fry and Hart, J, 1948; Steinhausen et al., 2008). Cardiorespiratory collapse has been observed in adult migrating sockeye salmon when the temperature goes beyond these thermal limits for increasing heart rate (Eliason et al., 2013; Farrell et al., 2008). Furthermore, cardiac excitability can determine the minimum as well as the maximum temperature tolerance of fish (Drost et al., 2014; Haverinen and Vornanen, 2020a; Safi et al., 2019; Sidhu et al., 2014; Vornanen, 2016). Thus, the temperature-dependence of cardiac excitability is of growing importance amid 5  concerns for fish populations to cope with increasing global temperatures. In my thesis, I focus only on the temperature-dependent depression of cardiac pacemaking as the limiting factor to cardiac thermal tolerance.    Thermal acclimation of fishes  The thermal history of the fish is important for determining cardiac excitability as temperature can induce resetting of the intrinsic firing rate of the cardiac pacemaker and modify the levels of proteins and different isoforms (Sutcliffe et al., 2020). Acclimation (a reversible physiological change in response to a change in an environmental parameter) or acclimatization (a change in physiological functions as a result of complex environmental changes) can induce temperature-dependent responses that provide important physiological changes (phenotypic plasticity) for the fish (Gamperl and Farrell, 2004; Schaefer and Ryan, 2006). The cardiac responses generally differ for acute and chronic temperature exposures. Changes in gene expression following acclimation can result in compensatory mechanisms or expression of protein isoforms adapted for the new environmental temperature and result in improved whole-animal performance than if the thermal change were acute (Goldspink, 1995; Graham and Farrell, 1989; Schulte et al., 2011; Seebacher et al., 2015; Sutcliffe et al., 2020; Vornanen et al., 2005).  In general, warm acclimation increases both the minimum and maximum temperature tolerated (Beitinger and Bennett, 2000), thereby increasing upper cardiac thermal tolerance but sacrificing lower thermal tolerance (Aho et al., 1999; Farrell et al., 1996; Gamperl and Farrell, 2004; Vornanen et al., 2002a). With temperature acclimation, many fishes, but not all, can reset the intrinsic fH set by pacemaker cells (Farrell, 1991; Farrell, 2009), and hence modify their cardiac response to acute warming (Drost et al., 2016; Ferreira et al., 2014; Klaiman et al., 2011; 6  Vornanen, 2016). For example, with warm acclimation, intrinsic fH is lower at a given temperature than with an acute exposure to the same warm temperature without prior acclimation (Safi et al., 2019; Vornanen et al., 2002b; Vornanen et al., 2002a). Despite their importance, direct recordings of pacemaker APs during warming are still missing from the fish literature but would provide direct insight into the mechanisms of pacemaking and its thermal regulation. As a result, the direct effect of temperature on pacemaking mechanisms and the mechanism underpinning the resetting of fH observed in some species during thermal acclimation remains largely unexplored. Resetting of fH during thermal acclimation has never been investigated in the zebrafish. The aim of my thesis is to investigate the thermal acclimation capacity of the zebrafish and assess the effect of acclimation on the maximum fH. Furthermore, in my thesis, I aimed to provide proof-of-principle for a methodology enabling the recording of the electrical activity of in situ pacemaker cells in a reduced cardiac preparation using intracellular microelectrodes.    1.3 Electrical excitation of the fish heart Excitation of pacemaker cells and cardiomyocytes is controlled by the flow of ions across the sarcolemma (SL) (Figure 1.1) which, is driven by an electrochemical and concentration gradient generated by the unequal distribution of ions across the SL. This electrochemical gradient is maintained largely by SL sodium (Na+), calcium (Ca2+), and potassium (K+) ion channels, and the Na+/K+ ATPase, maintaining a negative intracellular potential across the SL and providing the driving force for the transmembrane movement of ions, notably, Na+, Ca2+, and K+ (Monfredi et al., 2013; Vornanen, 2016). Typical intracellular concentrations of these ions in teleost cardiomyocytes are around 13, 150 and 0.0001 mM for Na+, K+, and Ca2+, respectively, and 155, 7  4 and 2 mM for extracellular concentrations of Na+, K+, and Ca2+, respectively, resulting in a negative resting membrane potential (RMP) between -70 and -90 mV (phase 4; Figure 1.2. A) (Houston and Koss, 1984). In the zebrafish, the resting membrane potential of ventricular cardiomyocytes at room temperature (20 – 23°C) has previously been reported as -70.4 ± 2.8 mV (Brette et al., 2008), and – 71.5 ± 1 mV at 28°C (Nemtsas et al., 2010). APs are generated through the coordinated opening and closing of multiple ion channels that orchestrate the depolarization and subsequent repolarization of the membrane. Transmembrane movement of ions depends on the open state of the ion channels, which are either voltage-gated (VG), opening when the membrane potential is equal to their activation voltage, or ligand-gated, in which case their open state depends on the binding of the ligand to the binding domain of the channel (Bezanilla, 2005). Open channels allow a current to flow across the SL, modifying the membrane potential of the cell, which can depolarize (the membrane potential becomes more positive) or repolarize (the membrane potential becomes more negative). The directionality of the ionic flow depends on both the concentration gradient of each ion and the electrical gradient across the SL.  Working cardiomyocytes of the atrium (atrial cardiomyocytes) and the ventricle (ventricular cardiomyocytes) require external stimulus from an electrically coupled neighbouring cell or neuronal stimulation to initiate depolarization and generate an AP. Depolarization occurs when cations (positively charged ions), such as Na+ and Ca2+, enter the cell. Should the membrane potential sufficiently depolarize to reach the threshold potential for activation of VG-Na+ channels, the ion channels responsible for triggering the depolarization phase of the AP, the VG-Na+ channels will open. In the zebrafish heart, Nav1.5 is the principal cardiac isoform generating INa (Vornanen, 2017), and when expressed in Chinese hamster ovary cells has been shown to have a 8  half time activation of -47.5 ± 1.6 mV and a halftime inactivation of -79.3 ± 1.0 mV when recorded at room temperature (Chopra et al., 2007). In addition, the pool of Na+ channels available for activation is dependent on the RMP (lower RMP increases the pool of available channels for activation) and thus rectifying K+ channels have an indirect effect on INa as they contribute to determining the RMP and thus regulate the number of Na+ channels available for activation (Vornanen, 2017). Rapid entry of Na+ into the cell will depolarize the membrane potential. The AP is then propagated throughout the heart as a voltage wave through gap junctions connecting excitable cardiomyocytes of the working myocardium.   Cardiomyocyte contraction is initiated by the transient increase in the intracellular Ca2+ concentration ([Ca2+]i) resulting from the opening of depolarizing Ca2+ channels (Bers, 1993; Morad and Goldman, 1973; Vornanen, 2016). Ca2+ binds to troponin C, causing a conformational change in the troponin complex, thereby allowing myosin to bind to actin. Relaxation occurs when [Ca2+]i is reduced to the diastolic level by the removal of free Ca2+ from the cytosol, which is mediated by reuptake via sarcoplasmic reticulum Ca2+ ATPase (SERCA) pumps and the Na+-Ca2+ exchanger (NCX) (Bers, 2001). Therefore, temperature-dependent changes in depolarizing Ca2+ currents and repolarizing K+ currents in cardiomyocytes, including pacemaker cells, result in changes to the AP waveform and the frequency of AP generation (Hassinen et al., 2007; Haverinen and Vornanen, 2009; Vornanen, 2016).  9   Figure 1.1. Scheme of the major ion channels generating the action potential and calcium signalling. If, funny current; ICa-T, T-type calcium current; ICa-L, L-type calcium current; INa, sodium current; IK1, Inward rectifier current; IKr, rapidly activating delayed rectifier potassium current; IKs, slowly activating delayed rectifier potassium current; INCX, Na+-K+ exchanger; RyR, ryanodine receptor; and SERCA, Sarcoplasmic/endoplasmic reticulum calcium ATPase.  The action potential of the working myocardium The voltage wave arriving from neighbouring cells initiates a chain of coordinated, voltage-gated, and time-delayed reactions that result in the depolarization and subsequent repolarization of the cell. Most, but not all of what we know concerning AP mechanisms comes from mammalian studies. The zebrafish ventricle shares the main characteristics of the mammalian AP, including 10  that of humans, as many ion channels are conserved and have similar gating properties (Sedmera et al., 2003; Verkerk and Remme, 2012; Vornanen and Hassinen, 2016). With the exception of the rapid phase-1 repolarization, which is absent in the zebrafish heart, all other phases (0-4) of the cardiac AP are shared between mammals and zebrafish and the ion channels that generate each phase are largely conserved (Vornanen and Hassinen, 2016). The electrical stimulus depolarizes the SL to the threshold potential of the VG-Na+ channels, causing them to open and allowing rapid entry of Na+ into the cell, which results in depolarization of the SL to a peak depolarization voltage (phase 0; Figure 1.2. A). Phase 1 repolarization is either highly reduced or totally absent from all fish hearts investigated thus far, including the zebrafish, due to the absence of the transient outward K+ current (Ito) (Alday et al., 2014; Vornanen and Hassinen, 2016) (Figure 1.2. A). The phase 0 depolarization also activates VG Ca2+ and K+ channels, which are slower to activate. Ca2+ channels are activated before K+ channels and provide a voltage-dependent entry of Ca2+ into the cell, driving depolarization and providing Ca2+ for Ca2+-dependent excitation-contraction of cardiomyocytes. There are two Ca2+ channels that generate a Ca2+ current (ICa): L-type and T-type Ca2+ channels. The T-type Ca2+ channels are activated at around -60 mV generating the T-type Ca2+ current (ICaT) (Nemtsas et al., 2010), whereas L-type Ca2+ channels are activated at -40 mV generating the L-type Ca2+ current (ICaL) (Hove-Madsen and Tort, 1998; Vornanen, 1997; Vornanen, 1998). In the zebrafish heart, the major Ca2+ channel transcript is the T-type channel (64.1%) and the L-type channel is the non-dominant isoform (33.8%) (Haverinen et al., 2018a), generating a large ICaT, in both atrial and ventricular cardiomyocytes, unique to the zebrafish heart (Nemtsas et al., 2010; Vornanen et al., 2018). In the zebrafish heart, ICaT reaches peak current density at -30 mV before the AP is fully depolarized and decays as ICaL reaches peak current density at 0 mV (Haverinen et al., 2018a). This delay assures continuity of depolarizing Ca2+ current to 11  oppose the rising time-delayed K+ current (IK) activated during the depolarization of the sarcolemma. Opposing the depolarizing ICa and repolarizing IK results in a plateau phase of the AP (phase 2; Figure 1.2. A,). The IK opposing ICa in phase 2 is comprised of IKs and IKr. IKr activates at -15 mV and rapidly inactivates at +23 mV and initiates the repolarization, whereas IKs inactivates close to RMP. However, zebrafish embryo cardiomyocytes do not have a functional IKs (Alday et al., 2014; Nemtsas et al., 2010) but IKs has recently been recorded in the adult zebrafish heart and is only present in the ventricle (Abramochkin et al., 2018). Finally, IK1, an inwardly rectifying K+ channel drives the tail end of the AP back to RMP in phase 3 of the AP (Luo and Rudy, 1991; Zeng et al., 1995). In zebrafish, IK1 in ventricular cardiomyocytes had a reversal potential of -81 ± 1.1 mV (Hassinen et al., 2015). Other than this difference between zebrafish embryos/larvae in cardiac ion channels, all other channels that have thus far been recorded are the same in the embryonic/larval and adult stages (Alday et al., 2014; Nemtsas et al., 2010).         12   Figure 1.2. Typical action potential (AP) of fish cardiomyocytes. (A) Five phases of the AP and the major ion currents that generate them are shown in parentheses. (B) Pacemaker cell AP from an enzymatically isolated pacemaker cell of the brown trout (Salmo trutta fario). The AP phase and the major ion channels of the vertebrate heart are shown. IK1, inward rectifier K+ current; INa, Na+ current; ITo, transient outward current; ICaL, L-type Ca2+ current; ICaT, T-type Ca2+ current; INCX, Na+-Ca2+ exchanger current; If, funny current; IKr, the rapid component of the delayed rectifier. Taken with permission from Vornanen 2017.   13   Cardiac pacemaking in the sinoatrial node The fH in all fish is determined by pacemaker cells of the sino-atrial node (SAN), where the cardiac action potential (AP) is generated. The SAN was first identified in mammals in 1907 over 100 years ago (Keith and Flack, 1907); only three years later, the SAN was then identified in fish (Keith and Mackenzie, 1910) where it is situated between the sinus venosus and the atrium (Newton et al., 2014; Tessadori et al., 2012; Yamauchi and Burnstock, 1968). The SAN is a ring-like structure at the base of the atrial valves generating the cardiac AP in specialized pacemaker cells (Haverinen and Vornanen, 2007; Newton et al., 2014; Vornanen et al., 2010; Yamauchi and Burnstock, 1968). In the zebrafish, the SAN can be identified by the co-expression of the transcription factor Islet-1 and the pacemaker ion channel Hcn4 (Newton et al., 2014; Stoyek et al., 2015).  Pacemaker cells are self-exciting cells; they possess voltage-gated ion channels, exchangers, and pumps, that enable the rhythmic generation and propagation of APs (Irisawa, 1978; Vornanen, 2016). The voltage change of the sarcolemma (SL) triggers a transient increase in [Ca2+]i which leads to the generation and propagation of an AP and the subsequent contraction of the atrial and ventricular chambers (Bers, 2002; Vornanen et al., 2002b; Vornanen et al., 2002a).  Pacemaker cells generate the pacemaker AP and thus differ in their electrophysiological properties from cardiomyocytes of the working myocardium (Baker et al., 1997; Brown and DiFrancesco, 1980; DiFrancesco, 2010; Irisawa, 1978; Monfredi et al., 2013). Pacemaker cells are defined by the spontaneous depolarization of phase 4 which produces the autorhythmicity of pacemaker cells of the myogenic heart. This pacemaker current replaces the RMP (Figure 1.2; phase 4 depolarization of the AP) found in excitable cardiomyocytes of the working myocardium (Harper et al., 1995; Hassinen et al., 2017; Irisawa, 1978; Monfredi et al., 2013; Tessadori et al., 14  2012; Vornanen, 2016). The spontaneous depolarization phase begins directly after repolarization and involves several SL ion channels and intracellular Ca2+ release from internal stores (see sections 1.3.1 and 1.3.2), which constitute two distinct pacemaking mechanisms, that slowly depolarize the membrane potential. Once the spontaneous depolarization reaches the threshold potential for Ca2+ channels, SL depolarization is driven by Ca2+ entry through the open Ca2+ channels. Repolarization of the pacemaker cell occurs directly after peak depolarization since pacemaker cells do not present an AP plateau phase despite involving the same ion channels as in the working myocardium (omitting IK1 that is responsible for maintaining RMP) which result from differential expression of ion channels between the different cardiac chambers and the SAN (Hassinen et al., 2021).    1.3.2.1 The membrane clock The term “membrane clock” refers to an ensemble of electrogenic proteins specific to the SL that generate spontaneous depolarization resulting in pacemaking APs. Although several ion currents are involved in the membrane clock, the principal component responsible for the spontaneous depolarization of the SL is the funny current (If), which is generated by hyperpolarization-activated cyclic nucleotide-gated (HCN) channels (DiFrancesco and Noble, 2012; DiFrancesco et al., 1986; Yampolsky et al., 2019). HCN channels are triggered upon hyperpolarization, between -60 and -40 mV, close to the RMP of most cardiomyocytes (Accili et al., 2002; Baker et al., 1997; Baruscotti et al., 2005; Robinson and Siegelbaum, 2003). If is a time-dependent, mixed inward Na+ and K+ current, which provides slow entry of a depolarizing current that gradually approaches the threshold potential of T-type Ca2+ channels (Baker et al., 1997; Brown and DiFrancesco, 1980; Brown et al., 1979; DiFrancesco, 1993; DiFrancesco, 2010) 15  (Figure 1.3; Figure 1.4). The rapid and brief opening of these channels occurs at voltages below AP threshold potential (-50 mV to -40 mV) (Hagiwara et al., 1988) and generates the initial rapid depolarization of the SL that drives the membrane potential towards the threshold potential of L-type Ca2+ channels. The opening of these Ca2+ channels allows further entry of Ca2+, driving depolarization of the SL and upstroke of the AP. Peak depolarization amplitude is determined by the opposing depolarization-activated repolarization currents and the Ca2+-dependent inactivation of the depolarizing current (Cros et al., 2014).   Depolarization of the SL also triggers the time-delayed opening of the rectifier K+ current, which orchestrates the repolarization and is comprised of a rapid component (IKr) and a slow component (IKs) (Hassinen et al., 2008; Nerbonne and Kass, 2005; Vornanen et al., 2002b). IKr is responsible for the initial phase of repolarization and inactivates rapidly, whereas IKs provides a longer repolarizing current and plays an important role in controlling the repolarization phase that results in hyperpolarization of the SL. In addition to IKr and IKs, an acetylcholine-activated inward rectifier current (IKACH) is activated under parasympathetic tone via muscarinic cholinergic receptors and can significantly reduce the repolarization duration. With this hyperpolarization, HCN channels are activated, and the cardiac pacemaker current generates the spontaneous depolarization, initiating the following AP and thus perpetuating the autorhythmicity of the membrane clock.   16   Figure 1.3. Schematic of the respirometry Schematic representation of the components of the membrane clock.     1.3.2.1.1 Hyperpolarization-activated cyclic nucleotide-gated (HCN) channels HCN channels are central to the membrane clock hypothesis. These voltage-dependent ion channels belong to the six-transmembrane segment channel superfamily, which include VG K+ channels, and conduct both Na+ and K+ (Biel et al., 2009; Sartiani et al., 2017). Vertebrates express four HCN isoforms (HCN 1-4), which assemble in homo- or hetero-tetramers, with channels forming via associations of HCN2 and HCN4 proteins (Much et al., 2003; Whitaker et al., 2007). HCN channels have a pore located between segments S5 and S6 of the four HCN proteins of the channel. Upon hyperpolarization, the pore of HCN channels open after the displacement of the voltage-sensitive S4, which displaces the link between S4 and S5 and dissociates S5 from S6 (Männikkö et al., 2002) although debate remains on the mechanisms of voltage sensing and channel opening (Mobli et al., 2017). Opening of the HCN channels results in a net inward depolarizing current and the relative conductance of K+ is four times higher than that of Na+, 17  although, the presence of a K+ ion in the channel pore may hinder Na+ conductance (DiFrancesco, 1981; Lee and MacKinnon, 2017).  In the heart, HCN channels are expressed in the pacemaking cells of the SAN and generate the depolarizing If current. The expression of HCN channels in the SAN provides strong evidence of the role of If in cardiac pacemaking (Newton et al., 2014; Tessadori et al., 2012; Yamauchi and Burnstock, 1968). Each of the four HCN isoforms possesses distinct electrophysiological properties and kinetics (Stieber et al., 2006), with HCN4 being the slowest activating channel with the highest cAMP sensitivity, producing the largest effect among the HCN channels measured as a shift in the half-maximal activation voltage (Table 1.1). Cilobradine, ivabradine, and zatebradine block all HCN channel isoforms and subtypes with a similar half-maximal inhibitory concentration (IC50; 0.99, 2.25, and 1.96 μM respectively) and thus do not provide HCN subtype-specific block (Stieber et al., 2006). With no pharmacological separation of HCN channels to determine their contribution to generating If, the relative contributions of the different HCNs to If are yet to be fully elucidated. In mammals, strong evidence supports that HCN4 is the primary pacemaker channel as HCN4 is the major HCN isoform expressed in the mammalian SAN (Satoh, 2003). Additionally, HCN4 has been identified as the primary carrier of If in multiple species, including rabbits (Altomare et al., 2003; Brioschi et al., 2009; Ishii et al., 1999; Shi et al., 1999; Tellez et al., 2006), mice (Garcia-Frigola et al., 2003; Marionneau et al., 2005; Moosmang et al., 2001), dogs (Zicha et al., 2005), and humans (Chandler et al., 2009; Ludwig et al., 1999; Thollon et al., 2007). Loss of function mutations in human HCN4 results in bradycardia and arrhythmia (Milanesi et al., 2006; Schulze-bahr et al., 2003; Verkerk and Wilders, 2015) while HCN4 gain of function mutations 18  conferring increased cyclic adenosine monophosphate (cAMP) sensitivity are associated with sinus tachycardia (Baruscotti et al., 2017).              19   Figure 1.4. A functional scheme of the membrane clock and calcium clock mechanisms. Pacemaker action potential of a rabbit sinoatrial nodal cell (red trace; top). The different phases of the AP are labelled. Schematic representation of the timing and magnitude of the different components of the membrane clock (middle) and the Ca2+ clock (bottom). During phase 4, local Ca2+ release from the sarcoplasmic reticulum (SR) gradually increases total cytosolic [Ca2+]. L-type Ca2+ channels activate at threshold potential and cause Ca2+-induced Ca2+ release from the SR via ryanodine receptors. Cytoplasmic Ca2+ is removed by both the SR Ca2+ pump, SERCA, and the sarcolemmal Na+-Ca2+ exchanger. MDP, maximum diastolic potential; DD, diastolic depolarization; ICa,T, T-type voltage-dependent Ca2+ current; ICa,L, L-type voltage-dependent Ca2+ current; INCX, Na+-Ca2+exchange current; IK, delayed rectifier K+ current; If, funny current; SERCA, sarco-endoplasmic reticulum ATPase; LCRs, local Ca2+ releases. Taken with the Copywrite holder’s permission from Monfredi et al., 2013.  20  1.3.2.2 The calcium clock   The calcium clock is a second, independent pacemaking clock, known to be involved in mammalian cardiac pacemaking, and is supported by diastolic intracellular Ca2+ release from the sarcoplasmic reticulum (SR) (Figure 1.5) (Maltsev and Lakatta, 2007; Maltsev and Lakatta, 2008). Local periodic Ca2+ release through ryanodine receptors (RyR) occurs spontaneously in the form of sparks, which increase the [Ca2+]i (Hüser et al., 2000; Ju and Allen, 2000; Maltsev and Lakatta, 2007). This spontaneous local SR-Ca2+ release (Figure 1.4) is independent of membrane depolarization (Vinogradova et al., 2004) and occurs even with inhibition of ICaT (Hüser et al., 2000). Ca2+ sparks occur through the accumulation of open state RyR, originating from the opening of a single RyR, resulting in the activation of RyR clusters in a Ca2+-induced Ca2+ release (CICR) manner, which generate a Ca2+ wave (Cheng et al., 1993). Blocking the RyR with ryanodine reduces the rate of spontaneous depolarization by inhibiting SR-Ca2+ release (Maltsev and Lakatta, 2007; Rubenstein and Lipsius, 1989). Continued SR-Ca2+ release increases [Ca2+]i, which activates the Na+-Ca2+ exchanger (NCX) and generates a small inward depolarizing current as more Na+ enters the cell than Ca2+ is extruded via the exchanger (3:1 ratio of Na+/Ca2+), further depolarizing the cell which triggers the opening of L-type Ca2+ channels, greatly increasing [Ca2+]i (Lakatta et al., 2010). Ca2+-mediated activation of the RyR channels triggers CICR emptying of SR Ca2+ stores. The large increase in [Ca2+]i depolarizes the SL, driving the membrane potential to threshold potential of voltage-gated T-type Ca2+ channels and initiating an action potential (Monfredi et al., 2013). The SERCA pump then restores SR Ca2+ whilst NCX in reverse mode extrudes Ca2+ across the SL (Monfredi et al. 2013; Vornanen, 2016).   21   Figure 1.5. Schematic of the respirometry Schematic representation of the components of the calcium clock.     1.3.2.3 The coupled clock hypothesis Several lines of evidence point towards a coupled clock pacemaking mechanism resulting from the crosstalk between the membrane and calcium clocks rather than two independent clock mechanisms (Figure 1.6), which led to the establishment of the coupled clock hypothesis in 2009 (Maltsev and Lakatta, 2009). One argument for this is the activation of the SL protein NCX during the later phase of the diastolic depolarization in response to increased [Ca2+]i (Bogdanov et al., 2001; Maltsev and Lakatta, 2007). Furthermore, pharmacological inhibition of NCX exacerbates the bradycardic effect of If inhibition (Kohajda et al., 2020). The NCX can operate in forward and reverse modes, allowing three Na+ ions to enter the cell whilst extruding one Ca2+ ion in forward mode, or the opposite in reverse mode (Shigekawa and Iwamoto, 2001). Under resting physiological conditions of cardiomyocytes, the reversal potential for NCX is around -20 mV, corresponding to a large driving force for Na+ entry when the membrane potential is hyperpolarized at around -50 mV. In dynamic conditions including cardiac stimulation, Ca2+ influx 22  is favoured and Na+ is extruded at potentials positive of -20 mV (Baartscheer et al., 2011). Furthermore, late phase diastolic depolarization leads to the activation of T-type Ca2+ channels (at more negative potentials than L-type Ca2+ channels), which contribute to the pacemaking potential (Mangoni et al., 2006), connecting spontaneous slow depolarization with rapid depolarization. Although the calcium clock spontaneously generates local Ca2+ release and is thus believed to be an independent clock (Lakatta and DiFrancesco, 2009; Vinogradova et al., 2004), the calcium clock activates SL NCX making the calcium clock a coupled clock (Yaniv et al., 2015). Furthermore, evidence suggests, that T-type Ca2+ channels may be the trigger for SR-Ca2+ sparks by providing the RyR-activating Ca2+ (Hüser et al., 2000) in contradiction to the autonomous Ca2+ cycling activity assumed by the Ca2+ clock and thus debate remains over the existence of the coupled clock (DiFrancesco, 2020).  The depolarizing current generated by SL If favours the activity of NCX in forward mode, which further depolarizes the SL and activates T-type and L-type Ca2+ channels (Lakatta and DiFrancesco, 2009; Monfredi et al., 2013). Ca2+-activated RyR channels open with the L-type Ca2+-mediated increase in [Ca2+]i, coupling membrane, and calcium clock in the generation of the Ca2+ wave (Monfredi et al., 2013).  The dual affiliation of NCX and L-type Ca2+ channels mechanistically links the membrane and calcium clocks in a coupled clock. In the coupled clock, the membrane and calcium clocks are capable of independently creating a depolarizing pacemaking current, and their level of integration may be species-dependent and modulated by abiotic environmental factors, including temperature (Joung et al., 2011).  23   Figure 1.6. Schematic of the respirometry Schematic representation of the components of the coupled clock system. Red lines indicate coupling of the two clock mechanisms.  Few studies have addressed the relative contribution of the membrane and calcium clock of cardiac pacemaking, and the relative contribution of the clocks have not been quantified in fish. The coupled clock hypothesis may not fully explain the underlying mechanisms as the relative contribution of its components do not always account for the fH observed in wild-type and unmanipulated animals (Logantha et al., 2016). Furthermore, mitochondrial Ca2+ stores and buffering capacity have been suggested to play a role in pacemaking (Maltsev et al., 2014; Zhang et al., 2015). The relative significance of the membrane and calcium clocks in the fish cardiac pacemaking cells remains unknown (Vornanen, 2016).    24  1.4 Pacemaking in fish: role of the membrane and calcium clocks Pacemaking action potentials of the fish heart were first recorded in the hagfish (Jensen, 1965) followed by teleosts, including the European carp (Cyprinus carpio) (Saito, 1969; Saito, 1973; Saito and Tenma, 1976), and, more recently, the European plaice (Pleuronectes platessa) (Harper et al., 1995), rainbow trout (Oncorhynchus mykiss) (Haverinen and Vornanen, 2007) and the brown trout (Salmo trutta) (Hassinen et al., 2017). In addition, pacemaker APs have been recorded from isolated zebrafish pacemaker cells by patch clamp and stimulated at 3 Hz (Tessadori et al., 2012). As noted above, the fish pacemaker AP strongly resembles those of mammals as the orthologues of the major ion channels and ionic mechanisms are conserved in the fish heart (Vornanen, 2016; Vornanen and Hassinen, 2016). Evidence in some species suggests a membrane clock-based pacemaking mechanism (hagfish), whereas others suggest that Hcn channels play a reduced or no role in cardiac pacemaking (brown trout) (Hassinen et al., 2017; Wilson et al., 2016). The evidence for spontaneous depolarization of fish pacemaker cells being, in part, generated by Hcn channels involves the pharmacological block of Hcn channels. For example, zatebradine, an HCN antagonist, abolishes atrial contractions in the Pacific hagfish (Eptatretus stoutii) (Wilson and Farrell, 2013) and reduces resting fH by 11% and maximum fH by 33% in steelhead trout (Keen and Gamperl, 2012). Furthermore, ivabradine, another HCN antagonist, reduced fH in isolated zebrafish hearts by 60% (Lin et al., 2014). Zebrafish homozygous for the recessive slow mo mutation provide genetic evidence for Hcn channel-mediated pacemaking as the mutant fish have a reduced fH due to a reduction in the If generated by Hcn channels (Baker et al., 1997). Contrary to this, recording of the If in pacemaker cells of the brown trout (Salmo trutta fario) showed a 25  surprisingly small current (compared to the large If current generated in the Hcn4 expression systems) of only 1.2 pA/pF-1 at the voltage range of If, inconsistent with the membrane clock pacemaking and despite a strong Hcn mRNA expression (Hassinen et al., 2017).   Currently, very little is known about the role of the calcium clock in fish cardiac pacemaking. In mammals, SR-Ca2+ cycling is considered to be a key contributor to cardiac pacemaking (Bers, 1993; Bers, 2002; Joung et al., 2011; Lakatta and DiFrancesco, 2009; Lakatta et al., 2008; Maltsev and Lakatta, 2007). In ventricular cardiomyocytes of zebrafish, studies estimate that Ca2+ release from the SR during development of the Ca2+ transient is less than 20% of the total Ca2+ fraction, and Ca2+ sparks occur at a lower frequency and have a lower amplitude than Ca2+ sparks observed in mammals (Bovo et al., 2013; Shiels and White, 2005). In addition, when compared with rabbit ventricular cardiomyocytes, less Ca2+ is released from the zebrafish SR (Bovo et al., 2013). Inhibition of the calcium clock using ryanodine and thapsigargin in rainbow trout results in a 44% reduction in sinoatrial beating frequency, but only at high test temperatures near the upper thermal limit (18°C) suggesting a role in cardiac pacemaking (Haverinen and Vornanen, 2007). However, application of ryanodine in the hagfish does not affect fH, suggesting that the calcium clock plays little to no role in pacemaking of the hagfish heart (Wilson and Farrell, 2013). SR Ca2+ handling in the teleost heart is known to be both species- and temperature-dependent (Hove-Madsen et al., 1999; Shiels et al., 2004; Vornanen, 1998). The relative role of L-type Ca2+ channels and SR Ca2+ handling differs among species and greater SR Ca2+ release for cardiac pacemaking is perhaps characteristic of species with high swimming performance (Aho et al., 1999; Rivaroli et al., 2006; Shiels and Farrell, 2000; Shiels et al., 1998; Shiels et al., 1999). Mechanisms of cardiac pacemaking may differ between species and vary with temperature. Furthermore, resetting of fH during thermal acclimation may involve shifts in pacemaking 26  mechanisms or changes in ion channel isoform expression. My thesis will focus on dissecting the relevance of the membrane and calcium clocks in cardiac pacemaking of the zebrafish and investigate the acclimation potential of the zebrafish with regard to these mechanisms.27  Table 1.1. Activation profiles and cAMP sensitivity of mammalian HCN channels.  HCN1 HCN2 HCN3 HCN4  Activation kinetics  Fastest  HCN3/4<HCN2<HCN1  HCN4<HCN3<HCN1/2  Slowest τ (associated hyperpolarization voltage) 13 ms (-100 mV)1 98 ms (-130 mV) 2   144-330 ms (-140 mV)3-5  179 ms (-110 mV)5  470 ms (-140 mV)4  500 ms (-100 mV)1 461-659 ms (-140 mV)3,5 679 s (-110 mV)5   Midpoint activation (V0.5)  -71 mV6 -78 mV (-75 mV)7  -100 mV (-100 mV)2   -78 mV6 -95 to -99 mV (-140 mV)3-5    -95 mV (-140 mV)4  -99 to -109 mV (-140 mV)3,5     Response to cAMP   Poorest8,9   Intermediate-high8,9   Poor- intermediate9  Highest8,9  cAMP activation voltage (associated hyperpolarization voltage) -98 mV (-100 mV)2   -86 mV (-140 mV; 100μM)3  -85 mV (-140 mV; 0.5 mM)4 -81 mV (-140 mV; 1 mM)5  -100 mV (-140 mV; 0.5 mM)4  -85 mV (-140 mV; 100μM)3 -94 mV (-140 mV; 1mM)5  τ cAMP  98 ms (-130 mV)2   165 ms (-140 mV; 0.5 mM)4 69 ms (-140 mV; 1 mM)5 510 ms (-140 mV; 0.5 mM)4  447 ms (-140 mV; 1mM)5 Mouse 1,2,4,6,7, human3,5 , rabbit1   28 A list of references for Table 1.1. 1. (Ishii et al., 2001); 2. (Santoro et al., 1998); 3. (Stieber et al., 2003a);  4. (Mistrik et al., 2005) ;5. (Ludwig et al., 1999); 6. (Santoro et al., 2000);  7. (Azene et al., 2005); 8. (Kaupp and Seifert, 2001); 9. (Lewis et al., 2010)                        29 In fish, the major isoform expressed in the brown trout heart was Hcn3, followed by Hcn4 (Hassinen et al., 2017), and in the rainbow trout, the major isoform expressed in the SAN was Hcn4  (Sutcliffe et al., 2020). However, expression of brown trout Hcn3 in Chinese hamster ovary (CHO) cells did not produce a current when stimulated across the voltage range of If, in contrast to Hcn4 (Hassinen et al., 2017). Interestingly, If recordings from primary pacemaker cells in the brown trout only produced a small current, and was determined inconsistent with pacemaking driven by a membrane clock mechanism, despite previous reports of 1 pA/pF in sinoatrial cardiomyocytes in the rabbit providing sufficient current to initiate an action potential (DiFrancesco, 1991). In addition, most of the cells (12 out of 16) identified as pacemaker cells did not produce any If current at all, despite optimized If recording conditions, further supporting the idea that If plays no role, or a reduced role, in cardiac pacemaking of the brown trout (Hassinen et al., 2017). Given the importance attributed to HCN4 in mammals, the uncertainty of its role in cardiac pacemaking in fish, and the fact that different Hcn isoforms cannot be blocked using pharmacology, I wanted to establish an Hcn4 knockout fish that would allow me to better dissect the membrane clock and the role of Hcn4 in cardiac pacemaking in the zebrafish.  Furthermore, as resetting of the fH is an intrinsic property of the heart, Hcn channels may mediate changes in intrinsic fH with acclimation. Only one recent study has investigated the role of Hcn expression with regard to thermal acclimation and resetting of the heart in rainbow trout (Sutcliffe et al., 2020). To fully understand the role of Hcn channels in cardiac pacemaking and resetting of the intrinsic fH, and given the absence of Hcn isoform-specific pharmacological agents, we must first establish an experimental model that is amenable to genetic modification, and that shows a temperature acclimation response.  Proteins such as Hcn can be targeted by CRISPR and a mutant line of Hcn-knockout fish can be generated, and their cardiac physiology examined.  My thesis targeted hcn4 in the zebrafish to investigate its role  30 in cardiac pacemaking and to determine the effect of knocking out an important gene for cardiac pacemaking on the whole-animal performance.   1.5 Temperature-dependent deterioration of cardiac pacemaking in fish Acute warming increases rates of physiological and biochemical processes exponentially until a maximum rate is achieved. In almost all fish species studied to date, acute warming increases both fH and cardiac output in parallel up to peak values with increasing temperature, before then declining  (Brett, 1971; Casselman et al., 2012; Clark et al., 2008; Clark et al., 2011; Drost et al., 2014; Ferreira et al., 2014; Fry, 1947; Gilbert et al., 2019; Gollock et al., 2006; Heath and Hughes, 1973; Sidhu et al., 2014; Steinhausen et al., 2008). Thus, the scope to increase fH dramatically decreases beyond an optimum temperature before falling close to zero at the critical temperature (Steinhausen et al., 2008). Consequently, fH could become a limiting factor for whole-animal performance at high temperatures.  The temperature-dependent increase in fH is thought to cascade from a direct effect on cardiac pacemaker cells (Randall, 1970) as isolated perfused heart preparations show an increase in fH despite being isolated from all neuronal and hormonal input (Farrell et al., 1988; Gilbert et al., 2019; Sutcliffe et al., 2020). Furthermore,  an isolated perfused heart retains the capacity to respond to modulatory agents, such as adrenaline and noradrenaline, demonstrating the intrinsic nature of the cardiac pacemaker to modify fH in response to stimuli (Farrell et al., 1988). Whole animal ECG recordings and isolated perfused heart preparations both show thermal ceilings of cardiac function (Badr et al., 2017; Casselman et al., 2012; Gilbert et al., 2019; Marchant and Farrell, 2019) demonstrating that the temperature-dependent deterioration of fH above its thermal maximum is an intrinsic property of the heart.   31 Extreme warming results in the widening of the QRS-complex corresponding to a delay of the AP conduction through the ventricle and ultimately to a complete loss of the QRS-complex and the periodic preservation of the P-wave, thus indicating loss of conduction to or within the ventricle (Badr et al., 2016; Marchant and Farrell, 2019; Vornanen et al., 2014). Cardiac arrhythmias appear as warming continues above thermal optimal of the fish and could be due to either failure of the impulse generation (initiation of the AP at the pacemaker cells) or failure of the impulse propagation (AP relay failure) (Haverinen and Vornanen, 2020a; Vornanen, 2016). Thermal effects on cardiac electrical excitation can easily be recorded by means of ECGs. ECGs provide information on the rate of AP generation and impulse propagation between cardiac chambers and possible disturbances of either impulse generation or impulse propagation (Badr et al., 2016).  The underpinning cause for pacemaker failure and indeed cardiac failure at high temperature remains unknown, in part due to the fact that the exact mechanism of SAN automaticity and the associated thermal response remain unclear (Vornanen, 2016). Of particular interest in determining temperature-dependent mechanisms of cardiac failure are the major ion channels, pumps, and exchangers that constitute the mechanisms of cardiac pacemaking which is known to be supported by at least two mechanisms: the membrane clock and the calcium clock. Debate remains, however, over the relative contribution of these mechanisms to pacemaking and the existence of crosstalk between the two mechanisms, resulting in a coupled clock mechanism (DiFrancesco, 2020). My thesis provides insight into the relative importance of the membrane and the calcium clocks in cardiac pacemaking of the zebrafish, and the relative importance of these mechanisms with incremental warming.     32 1.6 The use of zebrafish as a model for molecular and electrophysiological pacemaker research The zebrafish has become a valuable model organism in cardiovascular research due to the ease of genetic manipulation of its entirely sequenced genome and the high conservation of gene function between humans (Giardoglou and Beis, 2019). The transparency of the zebrafish embryo provides insight into cardiovascular development, and its small size allows the embryo to pursue development despite severe developmental defects in the cardiovascular system, which is not possible using mammalian models (Bakkers, 2011). Furthermore, the zebrafish embryo can pursue development during the first week of life without a properly functioning heart due to their small size and integumentary oxygen uptake capacity (Bang et al., 2004; Chen et al., 1996; Sehnert et al., 2002; Stainier et al., 1996; Strecker et al., 2011). While little is known about cardiac pacemaking mechanisms in fish, the zebrafish is emerging as a promising model due to its amenability to genetic engineering technologies, such as CRISPR, and unlike most mammalian cardiac knockout models, some zebrafish cardiac mutants can be raised to adulthood (Baker et al., 1997).  Cardiac pacemaker cells have been located in the SAN of the zebrafish and identified by immunohistochemistry (Stoyek et al., 2016). The zebrafish represents an excellent model for cardiac ECG recordings as the heart is situated close to the surface, and ECG recordings can easily be obtained using surface electrodes (Milan et al., 2006; Sidhu et al., 2014). Because zebrafish naturally inhabit a wide spectrum of temperatures ranging from 6°C in the winter to 38°C in the summer with daily temperature fluctuations of up to 5.6°C (Payne and Temple, 1996; Spence et al., 2008), they offer a wide temperature range for thermal acclimation, facilitating the detection of acclimation effects.     33 1.7 Thesis relevance and aims Although cardiac pacemaking in mammals has been extensively studied, debate remains on the relative importance of the two primary mechanisms. In fish, cardiac pacemaking is virtually unexplored, and most of what is currently known is inferred from the mammalian literature. Therefore, the overreaching goal of my thesis was to demonstrate the relevance of the membrane and calcium clocks in cardiac pacemaking of the zebrafish, establish the role of Hcn4 in the membrane clock, and investigate the thermal acclimation potential of the zebrafish with regard to these mechanisms. An additional goal of this thesis was to establish a methodology for recording APs from pacemaker cells in ex vivo zebrafish heart preparations, thus generating a tool to investigate some of the questions raised by this thesis work. The thesis is comprised of three chapters, each of which is outlined below with a brief rationale leading to the hypothesis of each chapter. Throughout the thesis, unless otherwise stated, the term zebrafish refers to the adult fish.    Chapter 2: Thermal limits of the zebrafish heart and contribution of membrane and calcium clock to cardiac pacemaking (Marchant and Farrell, 2019). Given that debate remains over the two mechanisms of cardiac pacemaking in the SAN tissue (a membrane clock driven by Hcn channels and a calcium clock driven by SR Ca2+ cycling) and that the involvement of SR Ca2+ cycling in contractile function varies as a function of temperature as well as among fish species, I investigated both the acclimation potential of the zebrafish heart at 18°C, 23°C, and 28°C and the two pacemaking mechanisms by pharmacologically blocking SERCA and RyR over an acute thermal ramp from 18°C to 40°C. I hypothesized that cardiac pacemaking would be largely dependent on the membrane clock, with warm acclimation and acute warming leading to lower intrinsic fH and that warm acclimation and acute warming would increase the participation by SR Ca2+ cycling.    34   Chapter 3: The role of hcn4 in pacemaking of the zebrafish heart and the thermal tolerance of the hcn4 mutant. Physiological characterization of a hcn4 mutant zebrafish. Given the discovery in Chapter 2 of the prominent role played by Hcn channels setting fH in zebrafish and given that the Hcn4 subtype of brown trout can produce a large If current when expressed in CHO cells, I used genome engineering to provide a physiological characterization of a hcn4-knockout zebrafish by characterizing its cardiac, thermal, swimming, and respiratory performances. I hypothesized that Hcn4 knockout zebrafish would have a reduced fH that was less responsive to incremental warming compared with wild-type fish. Consequently, mutant fish would have a reduced peak fH, which would compromise swimming capacity, reduce aerobic scope, and reduce thermal tolerance.   Chapter 4: The effect of blebbistatin-mediated uncoupling of excitation-contraction on the cardiac action potential in zebrafish. Given that future work in fishes on cardiac pacemaking and the transmission of the cardiac AP will require an electrically active, ex vivo heart preparation that is non-contractile, I validated blebbistatin as a pharmacological method of excitation-contraction uncoupling by characterizing the effects of blebbistatin on AP parameters over temperatures ranging from 28°C to 35°C. I hypothesized that blebbistatin would inhibit contraction of the working myocardium but not significantly alter AP parameters, regardless of acute test temperature.  35  Thermal limits of the zebrafish heart and contribution of membrane and calcium clock to cardiac pacemaking1 2.1 Synopsis Cardiac pacemaking is a vital function that must be maintained at all temperatures encountered by the fish. Prolonged periods of exposure to a new temperature may result in changes to cardiac pacemaking mechanisms, resulting in a resetting of the heart rate which may involve changes in gene expression. As outlined in chapter 1, two mechanisms are known to support pacemaking: the membrane clock involving Hcn channels, and the calcium clock involving SR Ca2+ cycling. Chapter 2 determines the thermal limits of the zebrafish heart when acclimated to 18°C, 23°C or 28°C. In this chapter, I show that the zebrafish is dependent upon two pacemaking mechanisms and it possesses a limited ability to reset the cardiac pacemaker with temperature acclimation. I took electrocardiogram recordings to follow the response of maximum heart rate (fHmax) to acute, 1°C incremental warming, from 18°C until signs of cardiac failure appeared (up to ~40°C) to assess the acclimation potential of the zebrafish heart. I also investigated the relative contribution of the membrane and calcium clocks to cardiac pacemaking using pharmacological agents to block the clocks and evaluate the corresponding decrease in fHmax. I found that the membrane clock possessed a slight dominancy over the calcium clock mechanism which plays an additional role in setting pacemaker activity that was independent of temperature, whereas the membrane clock was more dependent on temperature.    1 Chapter 2 has previously been published as Marchant, J. L. and Farrell, A. P. (2019). Membrane and calcium clock mechanisms contribute variably as a function of temperature to setting cardiac pacemaker rate in zebrafish Danio rerio. J. Fish Biol. 95, 1265–1274.  36 2.2 Introduction Projections for global warming include a predicted increase in extreme environmental fluctuations, accentuating the increasingly urgent need to address the effects of both acute and long term thermal stress to predict species’ capacity to survive acute exposures and adapt to long term temperature regimes (Somero, 2005).Temperature has been termed the ecological master factor as its effects manifest at every level of biological organization, from molecular interactions and stability of DNA duplexes and proteins to metabolic rate and population distribution (Brett, 1971; Dickson and Graham, 2004; Shelford, 1931). Temperature governs the interactions between molecules involved in biochemical reactions, which form the foundations of physiological mechanisms and processes (Brett, 1971; Dickson & Graham, 2005). Temperature, thus, modifies the rate of biochemical reactions and profoundly influences biochemical and physiological processes and alters the whole-body metabolic rate of an organism. Most organisms face variations in temperature, the duration of which can range from minutes (acute experiences, e.g., foraging in colder water) to months (chronic experiences, e.g., seasonal). Ectothermic animals are particularly susceptible to temperature variations as their body temperature varies with the temperature of the surrounding environment. Fish are, for the most part, ectotherms; they are exposed to both acute and chronic changes of environmental temperature, inducing short term or prolonged modifications to the body temperature. In order to survive the thermal stress and maintain the functionality of physiological processes, fish must adapt to the new temperature through phenotypic plasticity and changes in gene expression (Hochachka and Somero, 2002; Schulte et al., 2011; Seebacher et al., 2015). A central component in a fish’s response to changing temperature is the temperature-induced change to the heart rate, which, in almost all fish studied to date, increases in response to acute warming (Drost et al., 2014; Ferreira et al., 2014; Fry and Hart, J, 1948; Gollock et  37 al., 2006; Heath and Hughes, 1973; Sidhu et al., 2014; Vornanen et al., 2014). Acute increases in temperature increase heart rate to a peak followed by a decline, ultimately leading to arrhythmia and cardiac collapse (Farrell et al., 2008; Eliason et al., 2011; Verhille et al., 2013; Anttila et al., 2014; Vornanen et al., 2014). Due to a temperature-dependent acceleration of biochemical reactions, temperature elevation also increases oxygen demands (Clark et al., 2008; Steinhausen et al., 2008; Eliason et al., 2011; Keen & Gamperl, 2012; Ekström et al., 2016a). Additionally, the temperature-dependent increase in heart rate increases oxygen supply to tissues and is the primary means of meeting the increased energy demands, as the cardiac stroke volume is largely independent of temperature (Gollock et al., 2006; Steinhausen et al., 2008). However, it remains unclear if the temperature-dependent cardiac collapse is caused by a temperature-dependent disruption of oxygen supply to the cardiac tissue, decreasing the heart’s capacity to circulate oxygenated blood to all tissues, or if cardiac collapse is caused by the temperature-dependent deterioration of cardiac electrical excitability.  In many fish species, chronic exposure to a temperature leads to temperature acclimation, which is known to cause changes in the cardiac response to temperature, ultimately leading to intrinsic heart rate resetting (Drost et al., 2016; Ferreira et al., 2014; Klaiman et al., 2011; Vornanen, 2016). Although heart rate resetting has been shown in some species, not all fish reset their intrinsic heart rate, thus revealing a species-dependent response to thermal acclimation. For example, cold-active fish, such as rainbow trout (Oncorhynchus mykiss) and killifish (Fundulus heteroclitus), can increase heart rate after cold-acclimation to compensate for the direct effects of the cold (Aho and Vornanen, 1998; Aho et al., 1999; Safi et al., 2019). On the other hand, cold-dormant species, such as the crucian carp Carassius carassius L., reduce heart rate in response to cold-acclimation (Tiitu and Vornanen, 2002a; Tiitu and Vornanen, 2002b), a response which seems preparatory for a winter hibernation under ice cover. However, it has recently become apparent that species that are known to reset their  38 intrinsic heart rate, may not always do so when acclimated to a new temperature (Sutcliffe et al., 2020) and the recent thermal history of the fish may play a significant role in thermal acclimation capacity. Further, acclimation may entail dynamic and complex changes in intrinsic heart rate that may be in part be governed by varying cholinergic inhibition of resting heart rate (Ekström et al., 2016b). Central to the resetting of heart rate are cardiac pacemaker cells, which generate the action potential and are known to be directly influenced by temperature (Randall, 1970; Haverinen & Vornanen, 2007; Farrell & Smith, 2016). Cardiac pacemaking in fish is known to be supported by two pacemaking mechanisms that have also been identified in mammalians and amphibians, to varying extents. The membrane clock, driven by HCN (hyperpolarization-activated cyclic nucleotide-gated) channels, is responsible for the generation of the funny current (If), whereas local calcium release from the sarcoplasmic reticulum (SR) drives the calcium clock. Despite the vertebrate sinoatrial node (SAN) being discovered over 100 years ago as the cardiac site for pacemaking activity, debate remains on the origins of pacemaker activity in mammals and the relative importance of the membrane and calcium clocks, including the possibility that pacemaking may be a coupled clock resulting from crosstalk between mechanisms (DiFrancesco, 2020).   In fish, although little is known about the cardiac pacemaking mechanisms, Hcn channels (that carry If) and SR-Ca2+ cycling (that involves Ca2+ release from the SR) have been shown to be involved in cardiac pacemaking and both generate a depolarizing increase in intracellular Ca2+ concentrations. Zatebradine block of Hcn channels in the rainbow trout significantly reduced heart rate by up to half and reduced peak heart rate by around 33% rate (Altimiras and Axelsson, 2004; Gamperl et al., 2011). Additionally, zebrafish (Danio rerio) carrying the slow mo mutation have chronic bradycardia due to a reduction of the fast component of the funny current, possibly carried by Hcn1 or Hcn2, demonstrating the  39 importance of If in cardiac pacemaking of the zebrafish (Baker et al., 1997). Nevertheless, in the brown trout, If was found only in a small subpopulation of sinoatrial cells and produced a very small current, inconsistent with that of a membrane clock-driven pacemaker (Hassinen et al., 2017). The calcium clock also has been shown to be involved in cardiac pacemaking in the rainbow trout as the pharmacological block of RyR and SERCA pumps with ryanodine and thapsigargin, respectively, reduced the rate of spontaneous depolarization of isolated pacemaker cells by 44% in 18°C-acclimated fish (Haverinen and Vornanen, 2007). Consequently, the mechanisms and origin of cardiac pacemaking in fish, as well as the mechanisms of temperature-induced resetting of heart rate, remain elusive (Vornanen, 2016). The aim of chapter 2 is to determine the mechanisms involved in cardiac pacemaking and their relative contributions to pacemaking in the zebrafish by recording maximum heart rate (fHmax) during acute warming whilst pharmacologically blocking Hcn channels with zatebradine and blocking ryanodine receptors and SERCA pumps with ryanodine and thapsigargin respectively. I chose to record fHmax rather than routine heart rate to obtain the maximum heart rate at each test temperature which eliminates heart rate variation which would reduce the capacity to detect differences between the acclimation groups. Furthermore, chapter 2 aims to determine the acclimation capacity of the zebrafish heart and its response to acute incremental warming. I hypothesized that after temperature acclimation, the fHmax of fish acclimated to different temperatures would reset and that fHmax at different test temperatures would differ between the fish of different acclimation groups. I hypothesized that zatebradine would reduce maximum heart rate in vivo, indicating the involvement of Hcn channels in pacemaking and that Hcn channels would be the principal component driving pacemaking. I further hypothesized that the combination of ryanodine and thapsigargin also would reduce maximum  40 heart rate in vivo but to a lesser extent, indicating an involvement of a calcium clock in cardiac pacemaking but one of lesser importance than Hcn channels.  2.3 Materials and methods  Animals and husbandry A total of 90 wild-type AB strain zebrafish (Danio rerio; 6-9 months post-fertilization, mixed sex) were used in this study. All experiments were performed in accordance with the University of British Columbia Animal Care Committee (permit A18-0003). Fish were obtained from a local pet store (Noah’s Pet Ark, Vancouver, BC, Canada) where they were reared at 23°C, and were transferred to the UBC holding facility where they were maintained in 40 L aquaria in laboratory conditions. Fish were fed daily with fish flakes (Nutrafin max, Rolf C. Hagen Inc., QC, Canada); food was withheld for at least 12 h prior to experiments. Fish mass did not differ significantly (P ≥ 0.05, one-way ANOVA) among the three acclimation groups: 18°C-acclimated fish, 0.595 ± 0.017 g (n=33); 23°C-acclimated fish, 0.510 ± 0.076 g (n=30); and 28°C-acclimated fish, 0.589 ± 0.023 g (n=26).   Acclimation procedure Zebrafish were acclimated to three different acclimation temperatures (18°C ± 0.1°C, 23°C ± 0.1°C and 28°C ± 0.1°C) for a minimum of four weeks. Thirty fish per temperature acclimation group were held in 40 L aquaria fitted with a recirculating filtration system (AquaClear power filter, Rolf C. Hagen Inc., QC, Canada) with a 14:10 day to night photoperiod and continuous water aeration (>80% air saturation). The desired acclimation temperature was reached by adjusting 1°C per day, starting at their holding temperature of 23°C, until the desired temperature was reached.   41 There were three independent test groups for each acclimation temperature: untreated (control), zatebradine pre-treated (Hcn channel blocker to test the membrane clock mechanism), and ryanodine/thapsigargin pre-treated (RyR and SERCA blockers, respectively, to test the calcium clock mechanism). For each of the three acclimation temperatures, individual fish were acutely warmed from 18°C until heat-induced cardiac failure was observed.   Fish preparation and acute warming protocol Each fish was fitted with electrocardiogram (ECG) wires to record fHmax, which was pharmacologically induced in anesthetized fish, a technique first developed by Casselman et al. (2012) and subsequently refined for zebrafish (Sidhu et al., 2014). Individual fish were anesthetized at their acclimation temperature using a 100 mg L-1 tricaine methanesulfonate (MS-222) solution buffered with 50 mg L-1 sodium bicarbonate until loss of equilibrium and slowing of ventilation rate. Fish were weighed prior to intraperitoneal injection of atropine sulphate (1.2 μg g-1) and isoproterenol HCl (7.8 ng g-1) to induce fHmax by blocking cardiac vagal tone and maximally stimulating cardiac β-adrenoreceptors, respectively (Sidhu et al., 2014). In fish treated with pharmacological blockers to inhibit either the membrane clock or the calcium clock, the corresponding pharmacological agent was co-injected with atropine sulphate and isoproterenol HCl before the fish was placed into the recording chamber (see pharmacological agents and solutions). To ensure sufficient contact with the skin and reduce mucus production over the course of the subsequent incremental warming protocol, the scales directly over the heart were gently removed from the ventral surface using a scalpel blade. Subsequently, the fish was placed dorso-ventrally in a small opening cut into a sponge base inside a custom-made, double-walled Plexiglas water-bath (1 L), containing a maintenance dose of buffered MS-222 (80 mg L-1) for prolonged anesthesia. Continuous irrigation of the  42 gills was ensured throughout the experiment using a small water pump that provided 10 mL min-1 water flow to a modified pipette tip that was placed into the mouth of the fish and that directed the flow of aerated (>80% air saturation) water out of the mouth across the gills. In a pilot study, the fish could be maintained in this experimental setup for over three hours with continuous recording of high-quality ECGs at 23°C. Once the fish was in place, a custom-made ECG electrode (30 AGW copper wire sealed into a glass Pasteur pipette and exposed only at the point of contact with the fish) was placed on the ventral surface of the exposed skin, midline to the heart. A second electrode was placed caudal to the first on the body of the fish and served as the reference electrode. Electrodes were placed using micromanipulators and positioned to achieve a clear ECG signal. The ECG signal was amplified (Grass P55 AC; Astro-Med Inc., RI, USA) and recorded electronically using Powerlab 8/35 (ADInstruments Inc., CO, USA). Subsequent analysis was performed off-line using the heart rate analysis package of LabChart 8 (ADInstruments Inc., CO, USA).   Once a clean ECG signal was being recorded, the temperature was adjusted to 18°C at a rate of 1°C every 3-4 min if the acclimation temperature of the fish was different to 18°C. A dual temperature control system using two heater chillers (VWR 1160S, PA, USA) regulating the temperature of the water circulating within the walls of the experimental chamber and in the experimental tank provided accurate control over the fish’s temperature, which was continuously recorded using a temperature probe (accuracy ±0.1°C) coupled to an oxygen and temperature sensing unit (Fibox 3 trace, preSens). Heart rate (fH) was stabilized at 18°C for at least 30 min, or until stable fH was achieved for all fish, regardless of the acclimation or treatment groups. The experimental test temperature was then incrementally increased by 1°C every 5 min, and the ECG recordings were continuously monitored. The temperature was increased until signs of cardiac failure appeared on the ECG recording (cardiac arrhythmia and/or alternans; Appendix 2.1), at which point the fish was removed from the recording  43 chamber and euthanized by a lethal overdose of MS-222 followed by severing of the spinal cord and pithing of the brain. The number of fish used for each treatment group is provided in Table 2.1.   Pharmacological agents and solutions All chemicals were obtained from Sigma-Aldrich exception for zatebradine, which was obtained from Caymen Chemicals. The pharmacological agents were prepared fresh from stock solutions prior to each experiment. Zatebradine and ryanodine stock solutions were made in DMSO, thapsigargin was prepared in ethanol, and isoproterenol and atropine stock solutions were made directly in saline. All drug stock solutions were diluted in saline solution (in mM: NaCl 124.2; KCl 5.1; Na2HPO4; MgSO4 1.9; CaCl2 1.4; NaHCO3 11.9; pH 7.2) to injectable concentrations that could be intraperitoneally injected into the fish with the final injection volume ≤ 20 μL (equivalent to 5% of the fish’s body mass). Atropine and isoproterenol were used to induce fHmax. Zatebradine (4 μg g-1) was used to block all isoforms of cardiac Hcn channels (Altimiras and Axelsson, 2004), and ryanodine (50 ng g-1) in combination with thapsigargin (1.3 μg g-1) was used to block cardiac RyR and the SERCA pumps, respectively. The pharmacological blockers were injected after intraperitoneal injection of atropine and isoproterenol. Drug concentrations and efficiency were tested in a pilot study by two successive intraperitoneal injections of either zatebradine or ryanodine and thapsigargin (Appendix 2.2). The concentrations of atropine and isoproterenol HCl required for fHmax in the zebrafish were previously determined by performing subsequent drug injections that did not alter fH  (Sidhu et al., 2014) and the concentrations of the other pharmacological agents used were determined using the same protocol where the concentration was deemed sufficient if no change in fHmax was observed within the 5 min after subsequent drug injection. As described previously, fish  44 were anesthetized (100 mg L-1 MS-222) and injected with atropine sulphate (1.2 μg g-1) and isoproterenol HCl (7.8 ng g-1), before being placed in the recording chamber, with a maintenance dose of 80  mg L-1 MS-222. fHmax was recorded for a minimum of 20 min, or until stable, prior to intraperitoneal injection of zatebradine or ryanodine and thapsigargin. fH was recorded for 20 min after the injection, or 5 min after a stable fH was reached. A second injection of the same drug was administered and fHmax was recorded for a further 20 min.    Data analysis and statistics fHmax of individual fish was determined by analyzing the R-R interval (LabChart 8) over a 1-min period at each stable test temperature. Peak fHmax was determined as the highest fHmax attained during the incremental warming protocol and Tmax was the temperature at which peak fHmax was first attained. After reaching Tmax, further warming would result in either a plateau of fHmax (a change in fHmax of < 5 beats min-1) or fHmax would immediately decline. Continued warming would result in the appearance of signs of cardiac collapse, and the temperatures at which the first missing QRS complex (TQRS) arrhythmia (Tarr) and cardiac alternans (Talt) occurred were assigned. For each fish, the first Arrhenius breakpoint temperature (Tab) for fHmax was determined by fitting a two-segment linear regression to the natural log of fHmax against the inverse of temperature in K (1000 K-1) using GraphPad Prism 8.0 (GraphPad Software Inc., CA, USA) and the intersection of the two linear regressions was determined. The Arrhenius break point temperatures were used to determine the first point of deviation from the linear increase in fHmax, indicating a slowing of fHmax. The mean values for Tab were graphically represented for each acclimation group and only in the absence of pacemaker clock blockers.      45 The incremental Q10 for fHmax was calculated for each increment in temperature using the formula: 𝑄10 = (𝑓𝐻𝑚𝑎𝑥 2𝑓𝐻𝑚𝑎𝑥1)(10𝑇2−𝑇1)  where T1 and T2 designate the temperature increments, and fHmax 1 and 2 designate the fHmax recorded at those temperatures. As a result of increased metabolic rate, acute warming should result in a Q10 > 2 for heart rate. Therefore, threshold incremental Q10 was assigned when Q10 fHmax decreased below 1.9, and statistical significance between the mean values was determined using a one-way ANOVA with a Tukey post hoc test, using P ≤ 0.05 as the level of significance. The percentage decrease in fHmax from control was calculated for each drug treatment group and presented as the mean ± SEM for each acclimation temperature. The data were arcsine transformed prior to statistical analysis and before comparing all acclimation groups within a drug treatment group. Statistical significance was determined by fitting a mixed model with a Geisser-Greenhouse correction as implemented by GraphPad Prism 8.0, followed by a Sidak post hoc test, using P ≤ 0.05 as the level of significance. The mixed model for repeated measures using a residual maximum likelihood approach was used to determine the effect of treatment group and acclimation temperature in groups of declining numbers of individuals by modelling the individual subject variables whilst modelling the influence of nonlinear differences from the individuals included in the analysis over time (Krueger and Tian, 2004). In addition, the Geisser-Greenhouse correction was used to reduce the type 1 error rate in the mixed model analysis due to reduced sphericity of the fHmax data in response to increasing temperature between acclimation groups caused by different individual responses to incremental warming.  Finally, the Sidak post hoc test was used to control for type 1 errors of the pairwise comparisons at each test temperature. Interaction between treatment groups and acclimation temperature was performed using a two-way ANOVA.   46 2.4 Results  General effects of acute warming on fHmax Acute warming in 1°C increments progressively and continuously increased fHmax to a peak value (Figure 2.1), which ranged from 235 ± 9 beats min-1 to 296 ± 12 beats min-1 among the three acclimation groups (Table 2.1). While fHmax in all individual fish ultimately ceased to increase at 34°C (independent of acclimation temperature), and either plateaued or declined with further warming, some individuals reached a peak fHmax at a much lower temperature, especially in the 28°C acclimation group (Figure 2.2). Independent of acclimation temperature, the transition temperatures for fHmax were ordered similarly: Tab < Tmax < Talt < Tarr (Table 2.1) but TQRS, Tarr, and Talt were all found to be independent of the acclimation temperature (Table 2.1). These comparisons may have been confounded by the varying length of the plateau phase between individuals, which tended to be longer and more variable at warmer acclimation temperatures (Figure 2.2 A-C) and the low n values for some of the groups (Table 2.1) because not all fish showed missing QRS, alternans or arrhythmia. The temperature at which the first individual fish to show cardiac failure with a missing QRS complex (34°C) was also independent of acclimation temperature.           47 Table 2.1. A comparison of the cardiac performance variables for zebrafish. Cardiac performance variables are derived from the response of fHmax to acute warming at three acclimation temperatures for the three treatment groups (control conditions and two independent blocker treatments): Temperature of first Arrhenius breakpoint (Tab); Temperature at which the maximum value for fHmax was recorded (Tmax); Temperature of the first missing QRS complex (TQRS); Temperature of appearance of arrhythmia (Tarr); Temperature of appearance of alternan (Talt) and Maximum fHmax value (Peak fHmax). For each cardiac performance variable within a treatment group, whenever a statistically significant difference (P ≤ 0.05; one-way ANOVA) exists among the acclimation temperatures it is indicated by a dissimilar uppercase superscript. Likewise, for each cardiac performance variable within an acclimation temperature, whenever a statistically significant difference (P ≤ 0.05; one-way ANOVA) exists among the treatment groups it is indicated by a dissimilar lowercase superscript. For clarity, the letter superscript is omitted when no such differences exist for these comparisons (Table on the next page).                  48         Acclimation Temperature (°C) Peak fHmax  (min-1) Tab °C Tmax °C TQRS °C Tarr °C Talt °C Control 18 235 ± 9a,A 27.1 ± 0.007a 31.7 ± 0.8A 35.3 ± 0.4 35.4 ± 0.4 35.9 ± 1.0 n 9 9 9 9 9 3 23 296 ± 12a,B 22.4 ± 0.012 34.8 ± 0.7a,B 37.5 ± 0.8 37.8 ± 0.8 38.2 ± 1.1 n 11 11 11 10 7 5 28 248 ± 20a,A 25.4 ± 0.009 33.7 ± 0.9A,B 36.4 ± 1.7 37.6 ± 1.3a,b 35.9 ± 3.0 n 8 8 8 4 6 2 Zatebradine 18 175 ± 11b 28.7 ± 0.012b,A 32.7 ± 1.2 35.0 ± 1.2 35.3 ± 1.3 36.4 ± 2.2 n 13 13 13 7 7 2 23 173 ± 19b 26.0 ± 0.004B 32.0 ± 1.7b 36.44 ± 2.5 39.0 ± 1.5 38.54 n 6 6 6 4 2 1 28 141 ± 23b 26.6 ± 0.009A,B 33.0± 0.8 36.7 ± 0.7 38.0 ± 0.6a 38.0 ± 2.1 n 8 8 8 5 5 2 Ryanodine and thapsigargin 18 182 ± 16b 25.0 ± 0.014a 33.5 ± 0.7 34.7 ± 0.9 36.6 ± 0.5 35.6 ± 1.3 n 9 9 9 7 5 2 23 203 ± 25b 25.3 ± 0.008 30.2 ± 1.7b 35.1 ± 1.6 35.6 ± 0.9 31.4 ± 3.8 n 9 9 9 3 4 3 28 199 ± 12a,b 25.6 ± 0.020 32.4 ± 1.3 34.3 ± 1.5 33.8 ± 1.3b 33.4 ± 1.2 n 7 7 7 6 5 6  49  The effect of temperature acclimation on fHmax Mean fHmax was similar at almost all the incremental test temperatures, with significant differences only at 18°C and above 30°C (Figure 2.1). Specifically, at 18°C, mean fHmax of 23°C-acclimated fish (91 ± 2 beats min-1) was significantly lower than the mean fHmax of 18°C-acclimated fish (106 ± 3 beats min-1; P ≤ 0.05), but not the 28°C-acclimated fish (97 ± 3 beats min-1; P ≥ 0.05) indicating a small acclimation effect in the 18°C-acclimated fish. All fish in the 23°C acclimation group linearly increased  fHmax up to 33°C before temperature-induced cardiac collapse began to be observed; 10 out of 11 fish still had a regular heart rhythm at 37.5°C (Figure 2.1, 2.2). In contrast, temperature-induced cardiac collapse in all 18°C-acclimated fish occurred at the mean temperature of 35.3°C, and all but one fish showed cardiac collapse by 37°C.  The mean maximum temperature of the three acclimation groups was 33°C as fHmax decreased at 34°C, with the exception of those in the 28°C acclimation group.  Nevertheless, peak fHmax in 23°C-acclimated fish was significantly higher than in 18°C- and 28°C-acclimated fish (Table 2.1; Figure 2.1). In addition, Tmax for peak fHmax in the 18°C-acclimated fish was lower than either 23°C- or 28°C-acclimated fish; Tmax did not differ significantly between 23°C- and 28°C-acclimated fish (Table 2.1). Fish acclimated to 23°C reached a higher fHmax (by 21%) and a higher peak Tmax (by 3°C) compared to the 18°C-acclimated fish.    50  Figure 2.1. Effect of acute warming to 33°C on the mean fHmax (± SEM) of anesthetized zebrafish acclimated to 18°C (n=9), 23°C (n=10) and 28°C (n=8). Also shown is the temperature of the first missing QRS complex for individual fish and is expressed as a percentage of fish in each acclimation group. (P ≤ 0.05 calculated using a mixed model with Geisser-Greenhouse correction with a Sidak post hoc test). An asterisk indicates a significant difference between 18°C-acclimated fish and 23°C-acclimated fish.        51  Individual variability in fHmax was found to differ among acclimation groups with incremental warming. The individual variability in peak fHmax and the temperature at which fHmax started to plateau was greater in 28°C-acclimated fish, with peak fHmax ranging from 178 to 332 beats min-1 and the plateau phase commencing from 29°C to 37°C (Figure 2.2 C).  Further, the first Tab of the 23°C acclimation group was significantly lower than the two other acclimation temperature groups (Figure 2.3 B) indicating that cellular pacemaking mechanisms in the 23°C-acclimated fish limit the increase in fHmax at a lower temperature than fish acclimated to either 18°C or 28°C. In addition, a second Tab at 29.4°C was present in the Arrhenius plot of the 23°C acclimation group (data not shown). The incremental Q10 of fHmax remained at or above 1.9 up to 31°C for the 23°C-acclimation group, whereas the Q10 of the other two temperature acclimation groups dropped below and remained below 1.9 from 27°C and above (Figure 2.3 A). Lastly, all 18°C-acclimated fish and all but one of the 23°C-acclimated fish presented missing QRS complexes, whereas only half of the 28°C-acclimated fish presented missing QRS complexes at a high temperature. In the remaining 28°C-acclimated fish fH simply slowed and the ECG signal faded until no signal could be detected, perhaps indicating a different mechanism of cardiac failure (Figure 2.2). The effects of temperature acclimation on cardiac performance were, thus, found to be minimal as all acclimation groups linearly increased fH.        52   Figure 2.2. Individual responses to acute warming in 1°C increment of anesthetized zebrafish acclimated to either (A) 18°C (n=9), (B) 23°C (n=10) or (C) 28°C (n=8). The red dashed line indicates 33°C. Each line represents individual fish and are coloured to visualize the fHmax of individuals with increasing temperature.    53  Figure 2.3. Mean incremental Q10 and Arrhenius breakpoint plot of acclimated zebrafish. (A) Mean incremental Q10 of heart rate during incremental warming ± SEM. (B) Arrhenius plot of fHmax for 18°C, 23, and 28°C-acclimated zebrafish. The first Arrhenius breakpoint temperature was calculated from individual Arrhenius plots and the mean ± SEM for these calculations is presented for each acclimation group. Arrows indicate the temperature acclimation groups of the Arrhenius breakpoints. (P ≤ 0.05 calculated using a one-way ANOVA with Tukey post hoc test). An asterisk denotes a significant difference between 23°C acclimated fish and the 18°C and 28°C-acclimated fish.         54  Effect of Hcn channel blocking with zatebradine Zatebradine significantly reduced fHmax by at least 38% at all test temperatures and for all three acclimation temperatures (Figure 2.4 A-C; Figure 2.5 A). However, the percentage reduction in fHmax induced by zatebradine was influenced by the acclimation temperature. While zatebradine reduced mean fHmax by ∼65% in 28°C-acclimated fish across all test temperatures, the mean reduction in fHmax in 23°C-acclimated fish was significantly lower (around 40%) at almost all test temperatures (Figure 2.5 A). In contrast, the zatebradine response in the 18°C-acclimated fish was temperature-dependent, with zatebradine reducing mean fHmax by ∼65% at test temperatures from 18°C to 23°C, but only by ∼40% at test temperatures >30°C (Figure 2.5 A), thereby behaving similar to 28°C-acclimated fish at cold test temperatures and similar to 23°C-acclimated fish at warm test temperatures. Consequently, at the test temperature of 18°C, zatebradine significantly lowered mean fHmax in 23°C-acclimated fish to 34 ± 3 beats min-1 which was significantly lower than in 28°C-acclimated fish (68 ± 7 beats min-1; P ≤ 0.05) and in 18°C-acclimated fish (46 ± 5 beats min-1; P ≤ 0.05). Finally, there was a significant interaction between acclimation temperature and zatebradine treatment at all test temperatures from 18°C to 27°C, but not above 27°C, and is driven by the higher fHmax in zatebradine treated 23°C-acclimated fish. This interaction suggests that zatebradine treatment is less effective in 23°C-acclimated fish. However, the interaction detected may be due to the lower number of replications in this treatment group and may not have any biological significance.    Effect of SR calcium cycling disruption using ryanodine and thapsigargin Treatment with ryanodine in combination with thapsigargin significantly reduced fHmax at almost all test temperatures, and the reduction was largely independent of the acclimation temperature (Figure 2.4 A-C; Figure 2.5 A). Moreover, the percent reduction in fHmax in response to ryanodine and thapsigargin, which was around 40% at most test temperatures with  55 the exception of those >26°C for the 23°C-acclimated fish, which was significantly lower than the percent reduction in fHmax in zatebradine-treated fish at almost all test temperatures in fish acclimated to 18°C or 28°C (Figure 2.5 A, B). However, for fish acclimated to 23°C, the effects of zatebradine and ryanodine with thapsigargin in fHmax were similar for all test temperatures except at 33°C, even though the percentage decrease in fHmax with ryanodine and thapsigargin was closer to 20% at test temperatures above 26°C. Furthermore, three fish among the 23°C-acclimated fish treated with ryanodine and thapsigargin failed to maintain cardiac function above 26°C. Finally, no interaction between acclimation temperature and ryanodine and thapsigargin treatment were detected for any of the test temperatures suggesting that pharmacological block of the calcium clock is not affected by acclimation temperature.            56  Figure 2.4.  A comparison of the control responses of fHmax during acute warming in 1°C increments of anesthetized zebrafish acclimated to either  (A) 18°C, (B) 23°C or (C) 28°C with those following pre-treatment with either zatebradine (4 μg.g-1) or ryanodine (50ng.g-1) and thapsigargin (1.3ug.g-1). Dissimilar letters indicate a significant difference between the two types of pharmacological blockade at a given test temperature. (P ≤ 0.05 calculated using a mixed model with Geisser-Greenhouse correction with a Sidak post hoc test).  57  Figure 2.5.  The mean percent change in fHmax of zebrafish treated with either zatebradine or ryanodine and thapsigargin.  (A) The mean percent change in fHmax produced by zatebradine during acute warming of 18°C)-, 23°C- and 28°C-acclimated fish. (B) The mean percent change in fHmax was produced by ryanodine and thapsigargin during acute warming of 18°C-, 23°C- and 28°C-acclimated fish. Dissimilar letters indicate a significant difference between the acclimation groups at a given test temperature. (P ≤ 0.05 calculated using a mixed model with Geisser-Greenhouse correction with a Sidak post hoc test).        58  Effect of Hcn channel blocking combined with SR calcium cycling disruption Zatebradine combined with ryanodine and thapsigargin significantly reduced fHmax in all six test fish acclimated to and tested at 28°C. Mean fHmax was decreased by 88.5% from 231 ± 11 beats min-1 to 29 ± 3 beats min-1 (Figure 2.6. A), whereas zatebradine alone resulted in a reduction of fHmax of 60% and ryanodine and thapsigargin alone a reduction of 40%. The effect of zatebradine combined with ryanodine and thapsigargin produced sub-additive effects, as the mean fHmax does not cross the isobole line between the zatebradine and the ryanodine and thapsigargin response intercepts (Figure 2.6. B) (Tallarida, 2011), and resulted in an unexpected residual low fHmax. Intriguingly, after injection with zatebradine and ryanodine with thapsigargin, the hearts continued beating for up to one hour before the experiment was terminated, possibly suggesting an alternative pacemaking mechanism not previously described in fish. However, maximum tissue penetrance, and thus blockade of pacemaker mechanisms may not be achieved in the heart, and a complete block of the pacemaking mechanisms may not be achieved. Alternatively, the primary pacemaker is indeed fully blocked and the secondary pacemaker region, located in the atrioventricular node (Stoyek et al., 2016), continues to drive pacemaking at a reduced rate. This second possibility entails that there is either a different penetrance of the pharmacological agents at this tissue, or that the ion channels responsible for pacemaking have lower sensitivity to the blocking agents, or both.     59  Figure 2.6.    Individual responses of fHmax of anesthetized zebrafish acclimated to 28°C and tested at 28°C. (A) Comparison between control fish (n=6) and fish intraperitoneally treated with pharmacological pacemaker blockers zatebradine (Z), ryanodine (R) and thapsigargin (T) (n=6). (B) Isobole plot of the fHmax response to Z and to R with T showing the sub-additive effect of Z+R+T  ****; P ≤ 0.0001  (P ≤ 0.05 calculated by Students t-test).            60 2.5 Discussion Increasing global temperature has driven the need to better understand species capacity to adapt to long-term temperature regimes (Somero, 2005) and to better understand mechanisms of temperature-dependent deterioration of physiological performance. This chapter assessed the in vivo cardiac function of zebrafish as a function of both thermal acclimation and acute incremental warming. Only one previous study has investigated the effects of acute incremental warming on fHmax in the zebrafish (Sidhu et al., 2014). However, the effect of temperature acclimation and cardiac resetting was not investigated. Thus, this work builds on the existing literature, adding the effect of acclimation and, for the first time, dissects the pacemaking mechanism of the fish heart. At comparable acclimation (25°C-27°C) and test (28°C) temperatures, Sidhu et al. (2014) found that peak fHmax was somewhat higher than the peak fHmax reported in the present study (247.3 ± 4.5 beat min-1 vs. 202.0 ± 10.4 beats min-1). Other studies found fH between 120 and 180 beats min-1 at 28°C (Huang et al., 2010; Lee et al., 2016; Sampurna et al., 2018). All of these studies employed MS-222, which can lower fH, decrease ventricular contraction and lower transition temperatures (Denvir et al., 2008; Santoso et al., 2019; Sun et al., 2009a; Sun et al., 2009b). However, the effect of MS-222 is mediated by increased vagal tone (Lochowitz et al., 1974), which is inhibited by atropine in the fHmax induced fish and is thus not a concern for this data.   Extreme acclimation temperatures reduce thermal tolerance One of the aims of chapter 2 was to evaluate the effect of temperature acclimation on cardiac resetting in zebrafish using fHmax. Thermal acclimation was found to have little effect on resetting the heart rate as no significant difference was observed among all three acclimation temperatures at almost all equivalent test temperatures. This non-compensatory, or inverse acclimation, has previously been reported in the zebrafish where 18°C and 28°C acclimated  61 fish had similar fH at 18°C (78–79 bpm) and 28°C (162–169 bpm) for both cold- and warm-acclimated fish (Lee et al., 2016). This acclimation response can also be observed in the crucian carp (Carassius carassius) where the classical compensatory acclimation (i.e. a higher metabolic rate of cold-acclimated fish at warm temperature compared to warm acclimated fish) is not observed (Vornanen et al., 1992).  Despite the lack of cardiac resetting, thermal acclimation affected cardiac thermal tolerance in important ways. Thermal acclimation was found to limit the cardiac thermal tolerance of both the 18°C and 28°C acclimation groups since fHmax failed to linearly increase above 30°C, indicated by the Q10 < 1.9 in the 18°C and 28°C acclimation groups. Acute exposure to elevated temperatures reduced fHmax to a greater extent in the cold and warm acclimation groups than in the intermediate 23°C-acclimated temperature group. The warmest acute exposure temperature attained by all acclimation groups before the mean fHmax started to decline, was 33°C, regardless of the acclimation temperature. In a previous study, Sidhu et al. found that zebrafish held between 25°C and 27°C had a fHmax 313 ± 9 beat min-1, which was attained at 33.6 ± 0.7°C (Sidhu et al., 2014), consistent with the findings of this study and suggesting a possible thermal ceiling for the cardiac function of the zebrafish heart. The upper thermal tolerance of cardiac function in the brown trout (Salmo trutta fario) has been suggested to be linked to the heat-induced failure of sodium channels, which may limit cardiac function at a high temperature (Vornanen et al., 2014). Sodium channels are one of the primary ion channels involved in the electrical excitation of the heart and have been suggested to be more heat-sensitive than other ion channels. Effectively, Vornanen et al. have shown that the maximum thermal tolerance of sodium channels in the brown trout is 20.9°C, which is lower than that of IK1, ICa, or IKr, and may, to some extent, determine the upper thermal limits of the heart (Vornanen et al., 2014). The mechanisms underlying thermal tolerance of ion channels  62 in the zebrafish heart, however, remain unexplored, but clearly involve different ion channel protein isoforms with a greater thermal tolerance than those of the brown trout. The thermal history of the zebrafish may confer thermal resilience to arrhythmia as missing QRS complexes were observed at a lower temperature in the 18°C acclimation group than the 23°C-acclimated fish. However, not all of the 28°C-acclimated fish presented missing QRS complexes, and not all maintained cardiac function at a lower temperature than the 23°C-acclimated fish, further indicating the limited effect of thermal acclimation in zebrafish. Fish thermal tolerance is generally shifted to warmer temperatures for both the upper and lower thermal tolerance limits after a period of warm acclimation (Beitinger and Bennett, 2000). My data suggest, however, that the thermal acclimation capacity of the zebrafish is limited. Also, my findings of a limited thermal capacity and a common thermal ceiling among fish with different thermal histories are in accordance with current zebrafish literature.   Overall, my findings on the absence of an effect of temperature acclimation on cardiac resetting in the zebrafish and limited effects on thermal tolerance limits, and transition temperatures, are consistent with those found in the literature. Specifically, I found that the transition temperatures of cardiac function all occur in the same order with incremental warming as Sidhu et al (2014), and limited acclimation effect with regard to shifts in thermal tolerance, as CTmax only varied < 2°C for fish acclimated to temperatures ranging from 20°C to 30°C (Cortemeglia and Beitinger, 2005; Schaefer and Ryan, 2006; Sidhu et al., 2014). Collectively, my data suggest that zebrafish possess a low acclimation capacity, which is consistent with other tropical species that possess a low thermal acclimation capacity, perhaps linked to the limited thermal variation in their natural environment (Tewksbury et al., 2008). However, Previous studies on zebrafish acclimated to 20°C and 30°C have shown a critical thermal maximum (CTMax) of 39.2 ± 0.3°C and 41.7 ± 0.4°C respectively, (Cortemeglia and Beitinger, 2005), and 39.9 ± 0.1°C when acclimated between 25°C and 27°C (Sidhu et al.,  63 2014). Zebrafish acclimated to 24°C, 28°C, and 32°C do significantly increase CTMax with warm acclimation (Schaefer and Ryan, 2006). Although CTmax reported in the literature is much higher than all of the transition temperatures reported here (Tab, Tmax, Talt, TQRS, and Tarr), with increasing acclimation temperature, the temperature difference decreases between the transition temperatures and CTMax, indicating an increased upper thermal limit in the warm acclimated fish. Thus, at least for the zebrafish, fHmax may not be a substitute for determining optimum temperature and aerobic scope as suggested for salmonids by Casselman et al. (Casselman et al., 2012). However, comparing fHmax transition temperatures of atropine-treated, β-adrenergic-stimulated, and anesthetized fish to free-swimming non-treated fish for CTmax may result in an increased temperature difference between these parameters. Furthermore, Schaefer and Ryan found that a fluctuating thermal regime induced greater thermal tolerance as CTMax significantly increased with warm acclimation (Schaefer and Ryan, 2006). Heterogenous environments have been shown to induce greater phenotypic diversity, resulting in a greater diversity of physiological traits (Mittelbach et al., 1992; Trexler et al., 1990; Watters et al., 2003). Therefore, thermal life-history may have a greater influence on the cardiac performance of fish than recent acclimation. Not all studies, however, have found that fluctuating temperatures improve thermal tolerance. Fangue et al (2011) observed a lower thermal tolerance of tidepool sculpins (Oligocottus maculosus) reared in fluctuating temperature compared to those reared in standard laboratory conditions (Fangue et al., 2011).      Effect of zatebradine on fHmax Another aim of chapter 2 was to evaluate the relevance of two cardiac pacemaking mechanisms (the membrane clock and the calcium clock) and their temperature dependence. Blocking Hcn channels with zatebradine significantly reduced fHmax. Zatebradine is a non-specific HCN blocker inhibiting all isoforms of cardiac HCN channels (Stieber et al., 2006), of which HCN4 is the dominant subtype in mammals and principal component of the HCN  64 channel-driven membrane clock that generates the funny current (If) (Stieber et al., 2004; Wahl-Schott et al., 2014). In the rainbow trout, Hcn4 is also suggested to be the predominant Hcn isoform contributing to the generation of the If (Hassinen et al., 2017). However, Hcn3, the most abundant Hcn isoform expressed in the SAN, atrium and the ventricle, has been reported not to produce a current when stimulated within the activation range of the If (Hassinen et al., 2017). Furthermore, in the brown trout, Hcn4 has been reported not to play a role (or to play a minimal role) in fH regulation due to the small (1.2 ± 0.37 pA/pF) current it produces (Hassinen et al., 2017). Nevertheless, in zebrafish, Hcn channels have been previously reported to play a role in cardiac pacemaking as fH of isolated zebrafish hearts was decreased by 60% using ivabradine (10 μM), another HCN channel blocker, thus suggesting a role for Hcn channel function in zebrafish cardiac pacemaking (Lin et al., 2014). My results for 18°C and 28°C-acclimated fish are in accordance with this finding because block of Hcn channels with zatebradine resulted in a 60% decrease in heart rate at all test temperatures below 26°C. At the 18°C test temperature, zatebradine reduced mean fHmax by 37% in 23°C-acclimated fish, whereas fHmax was decreased by 56% and 65% in 18°C- and 28°C-acclimated fish respectively. These findings suggest a hitherto undescribed effect of thermal acclimation on the involvement of Hcn channels in zebrafish cardiac pacemaking. However, a caveat to this conclusion is the possible off-target effect of zatebradine on potassium currents, which drive the repolarizing Ik currents. The relative block of HCN channels (and HCN isoforms) and potassium channels is currently unknown but may be of significant relevance to this study as zatebradine block of Ikr would prolong the action potential, thereby decreasing fH (Carmen et al., 1996; Van Bogaert and Pittoors, 2003). If this were the case, I would have overestimated the relative contribution of Hcn channels to cardiac pacemaking in zebrafish. HCN4 knockout mice are reported to have a 37% lower fH and can develop atrioventricular block that is fatal in utero (Stieber et al., 2003b). Similarly, fH was reduced by  65 ∼50% in inducible cardiac-specific Hcn4 knockout mice, and If was reduced by 75-90% in isolated cardiomyocytes (Baruscotti et al., 2011). The heart of the slow mo mutant zebrafish beats 37% lower than control zebrafish due to a defective If current, providing strong genetic evidence that If produced by Hcn channels is involved in cardiac pacemaking in zebrafish (Baker et al., 1997). Baker et al. (1997) also showed that the If current recorded from isolated cardiomyocytes of the slow mo mutant was reduced by 85% compared with the If current in cardiomyocytes of wild-type fish when stimulated at -130 mV. They further describe the current as being composed of a fast and a slow component, of which the fast component is defective in the slow mo mutant. Thus, If is important for cardiac pacemaking in the zebrafish, but its elimination (or elimination of the fast component) only results in a less than half reduction in fH, suggesting redundant or backup mechanisms for pacemaking, such as the remaining slow If component (Baker et al., 1997), functional redundancy of Hcn isoforms, a calcium clock, or an alternative form of pacemaking are yet to be identified. Four Hcn isoforms are known in the zebrafish heart, all of which may contribute to pacemaking and may all be blocked to some extent by zatebradine (Stieber et al., 2006). Therefore, a partial block of one or more of the Hcn channels in zebrafish would underestimate the contribution of the Hcn channel-driven membrane clock to cardiac pacemaking.   Effect of ryanodine and thapsigargin on fHmax At the 18°C test temperature, blocking RyRs and SERCA pumps resulted in a 40% reduction in fHmax that was largely independent of the acclimation temperature across all test temperatures below ~26°C (with the exception of the 23°C-acclimated fish at 23°C). In the 23°C-acclimated fish, the reduction in fHmax was close to 20% and was significantly lower than the reduction in fHmax in the 28°C-acclimated fish at test temperatures above 29°C (with the exception of 33°C), suggesting a reduced SR handling mechanism for pacemaking in the 23°C- 66 acclimated fish at a high temperature. Furthermore, this finding suggests that the zebrafish heart increases the involvement of the SR in cardiac pacemaking at a high beating frequency for fish acclimated to warm temperatures. However, cold-acclimated zebrafish may have enhanced SR-Ca2+ handling, providing a depolarizing current at high temperatures. Cold-acclimated rainbow trout exhibited enhanced SR-Ca2+ uptake rate at 22°C, whereas warm-acclimated fish had a 44% reduced rate velocity (Aho and Vornanen, 1998). Although cardiac SR-Ca2+ cycling is considered to be a key contributor to cardiac pacemaking in mammals (Bers, 1993; Bers, 2002), the relative contribution of the SR to cardiac function in the teleost heart is known to be both species- and temperature-dependent (Hove-Madsen et al., 1999; Shiels et al., 2004; Vornanen, 1998) and might be a characteristic of high swimming performance species (Aho et al., 1999; Rivaroli et al., 2006; Shiels et al., 1999). My study provides insight into the importance of the SR for cardiac pacemaking in zebrafish and the independence of the calcium clock to temperature. The phylogenetic conservation of spontaneous SR-Ca2+ release between zebrafish and mammals has previously been highlighted (Llach et al., 2011), although this was not reported by an earlier study in rainbow trout (Shiels and White, 2005). Despite the functional conservation in the SR, RyR expression levels are reported to be 78% lower in zebrafish than in rabbit ventricular myocytes, and action potential-induced calcium transients in the zebrafish were only 20% mediated by calcium release from the SR (Bovo et al., 2013). Zebrafish calcium transients were demonstrated to be mediated mainly by Ca2+ influx from L-type Ca2+ channels and NCX (Bovo et al., 2013; Shiels and Galli, 2014). The findings of this study suggest that calcium transients in the zebrafish are likely mediated by Ca2+ influx from L-type Ca2+ channels and NCX in the pacemaker cells because of the greater effect of zatebradine on fHmax.   67  Effect of zatebradine, ryanodine, and thapsigargin on fHmax The additive effect of zatebradine, ryanodine, and thapsigargin significantly reduced fH to a greater degree than zatebradine or ryanodine with thapsigargin alone suggesting simultaneous block of both the membrane and calcium clocks. Although pharmacological block of both Hcn channels and SR-Ca2+ cycling did not result in a complete cessation of the heart, it resulted in a strong and sustained reduction in fHmax, possibly indicating the existence of an undescribed pacemaking mechanism. Similar results were observed in SERCA2 knockout mice, in which pharmacological block of If (2 mM CsCl - an HCN channel blocker) equally reduced fH in wild-type and knockout mice, but did not stop the heart (Logantha et al., 2016). However, the application of ryanodine resulted in a 10% reduction in fH of SERCA2 knockout mice, suggesting residual SERCA2 driving the calcium clock. Additionally, it has been reported that residual (< 5%) SERCA2 protein in SERCA2 knockout mice was capable of partially replenishing SR-Ca2+ content (Andersson et al., 2009; Louch et al., 2010). Therefore, if the pharmacological block of RyR and SERCA pumps is not total, residual functional calcium clock components may be sufficient to drive cardiac pacemaking. Further, the volume of the SR in fish is greater than that in mammals (Shiels, 2017), and plays a more significant role in the zebrafish pacemaking mechanism than in mammals (Lakatta and DiFrancesco, 2009).   Conclusion In chapter 2, the thermal acclimation capacity of the zebrafish heart was shown to be limited, and in particular, the intrinsic fHmax did not reset with thermal acclimation. Furthermore, extreme temperatures reduced the fish’s upper thermal tolerance as fish that were acclimated to the intermediate temperature of 23°C were somewhat more thermally tolerant than colder and warmer temperature acclimation groups. A test temperature of 33°C was  68 identified as the thermal ceiling for zebrafish since peak fHmax did not increase further in all three of the acclimation groups as fHmax either plateaued or declined with further warming. The zebrafish’s capacity to cope with increasing temperatures predicted by global warming may be limited by the heart in particular due to its limited acclimation capacity. However, the limitation in thermal tolerance of the heart appears to occur in the working myocardium rather than in the pacemaker cells. This research significantly increases our current understanding of pacemaking mechanisms in fish, but mechanisms of pacemaking are likely species-dependent and thus these findings will not be widely applicable to all fish species.  Cardiac pacemaking of the zebrafish heart clearly involved both the membrane and calcium clock mechanisms, with a dominance of the membrane clock in the 18°C and 28°C-acclimated fish, but no clear dominant mechanism in the 23°C-acclimated fish. The membrane clock was found to be largely independent of both acclimation and acute test temperatures, whereas the calcium clock was temperature-dependent. As neither the membrane nor calcium clock blockers completely stop pacemaking, mechanistic crosstalk may be less profound in zebrafish than in mammals and may involve a third, yet undescribed pacemaking mechanism.   Limitations and future directions I acclimated fish for four weeks, which could be insufficient for cardiac thermal remodelling in the zebrafish. The impaired cardiac performance of the 28°C-acclimated fish at a high temperature may have resulted from exposure to a considerably colder (18°C) test temperature prior to warming. In addition, fish were acclimated to 33°C but did not survive the decrease in temperature down to 18°C followed by incremental ramping and were thus not included in this study. Investigation into the effect of acute temperature changes prior to incremental thermal challenges should be further investigated as these may result in an acute response rather than an acclimation response. As such, the methodology of such experimental  69 design needs to be reconsidered and the thermal history of the fish taken into consideration, as well as the species-dependent response to thermal acclimation. Future work is required to quantitively dissect thermal sensitivity and resilience of the membrane and calcium clock mechanisms to enhance our understanding of heat-induced cardiac failure in fish. A significant challenge associated with in vivo experimentation is control over the amount of pharmacological agent reaching and acting upon the organ and mechanism of interest. Due to the in vivo nature of this work, I cannot certify the molarity of the pharmacological agents acting on the heart and the pacemaker cells, nor can this be standardized between fish beyond injecting a mass-adjusted volume of pharmacological agents for each fish. Pharmacological agents targeting specific ion channels with no off-target effects as well as organ-specific drug delivery systems will drastically improve the efficiency of in vivo cardiac physiology investigation. Furthermore, genome editing technologies specifically targeting genes encoding for ion channels and different isoforms of channels can provide insight into the roles of ionic currents in fish cardiac pacemaking.  Chapter 3 further investigates the membrane and calcium clock mechanisms in cardiac pacemaking by investigating the specific role played by Hcn4, using a CRISPR hcn4 knockout zebrafish line. I generated the hcn4 knockout fish to resolve the relative importance of Hcn4 in generating the If current of the spontaneous depolarization phase of the pacemaker AP, an unachievable goal using pharmacological agents as all currently available pharmacological agents affect all of the Hcn isoforms.      70  The role of Hcn4 in pacemaking of the zebrafish heart and the thermal tolerance of the hcn4 mutant - Physiological characterization of a hcn4 knockout zebrafish. 3.1 Synopsis  The pacemaker current of the heart in all chordates studied so far involves hyperpolarization-activated cyclic nucleotide-gated (HCN) channels that open on hyperpolarization and are a crucial component of the membrane clock hypothesis for the generation of cardiac pacemaking. Since HCN4 channels are key components in mammalian pacemaking, I investigated the importance of Hcn4 in the zebrafish (Danio rerio) cardiac pacemaking and its consequences to the whole animal by generating a CRISPR-mediated hcn4 knockout line. In addition, I investigated the role of Hcn4 in cardiac thermal tolerance as Hcn4 may play a role in the temperature-dependent deterioration of electrical excitation. I hypothesized that the knockout would result in chronic bradycardia, reduced swimming performance, reduced aerobic scope and reduced thermal tolerance. Hcn4 mutant zebrafish exhibited total compensation of pacemaking activity, exhibiting the same maximum heart rate across a wide thermal range (28°C-40°C) as wild-type zebrafish. However, hcn4 mutant zebrafish maintained maximum heart rate at a significantly higher temperature and with a higher frequency when the calcium clock, driven by the sarcoplasmic reticulum was pharmacologically blocked. Lastly, hcn4 mutant zebrafish maintained swimming capacity (Ucrit), cardiac function (electrocardiogram), and had a higher thermal tolerance (CTmax). The role played by Hcn4 in zebrafish cardiac pacemaking can be completely compensated for by alternative pacemaking mechanisms, presumably, other Hcn isoforms as metrics of cardiac function investigated here were not significantly impacted by the Hcn4 knockout.  71 3.2 Introduction At the cellular level, temperature-dependent deterioration of mechanisms underpinning vital functions may contribute to the decline of functions at the level of the organism. In particular, cardiac function in fish is highly temperature-dependent (Randall, 1970), increasing with increasing temperature and then rapidly declining as fish approach their upper thermal limits, limiting thermal tolerance (Eliason et al., 2011; Farrell et al., 2008; Verhille et al., 2013; Vornanen et al., 2014). Novel thermal conditions can result in physiological changes to cardiac function and performance (Nyboer and Chapman, 2018). As such, thermal tolerance can be modulated during thermal acclimation or acclimatization, resulting in changes in gene expression, modifying whole animal temperature tolerance (López-Olmeda and Sánchez-Vázquez, 2011; Schulte et al., 2011; Seebacher et al., 2015; Somero, 2005). Changes in gene expression that occur during thermal acclimation, and that lead to a modification of the intrinsic heart rate, are likely to include genes encoding for proteins involved in pacemaking. Moreover, autonomic interventions may play a protective role in fish hearts at high temperature, with increased vagal tone slowing heart rate (Eliason et al., 2013; Farrell et al., 1996; Hanson et al., 2006) and β-adrenergic stimulation increasing the maximum thermal performance of the heart (Gilbert et al., 2019). Nevertheless, the mechanisms underlying temperature-dependent deterioration of cellular pacemaking remain elusive.  Temperature-dependent deterioration of cardiomyocyte excitation and subsequent contraction is thought to be an important cause of temperature-induced deterioration of cardiac function. Elevated temperature-induced cardiac arrhythmia and missing QRS complexes (representing ventricular contraction on the ECG) ultimately lead to asystole (Anttila et al., 2013; Badr et al., 2016; Casselman et al., 2012; Ferreira et al., 2014; Fry, 1947; Gollock et al., 2006; Heath and Hughes, 1973; Marchant and Farrell, 2019; Sidhu et al., 2014; Vornanen, 2016; Vornanen et al., 2014).   72 Cardiac pacemaking is generated by spontaneous depolarization of specialized pacemaker cells located in the sinoatrial node (SAN) (Haverinen and Vornanen, 2007; Newton et al., 2014). This spontaneous AP sets the intrinsic heart rate which is then modulated primarily by autonomic controls (Stoyek, 2016; Vornanen, 2017). In fishes, pacemaking is highly temperature-sensitive and allows temperature-dependent modulation of cardiac output by increasing heart rate (fH) to meet the increasing oxygen demands due to temperature-induced increases in metabolic rate and oxygen consumption (ṀO2). Debate remains, however, over the mechanisms that generate pacemaking activity in fish as two mechanisms are known to support pacemaking: a membrane clock involving Hcn channels and a calcium clock driven by SR-Ca2+ cycling (Hassinen et al., 2017; Marchant and Farrell, 2019). However, the mechanisms of cardiac pacemaking and their relative importance appear to be species-specific among fishes. For example, pacemaking in the Pacific hagfish is solely reliant on Hcn channels as pharmacologically blocking them completely stops the heartbeat (Wilson et al., 2013). As shown by chapter 2, zebrafish apparently use both pacemaking mechanisms regardless of the test temperature across an acute thermal range (Marchant and Farrell, 2019), In contrast, Hcn channels in brown trout, play a reduced role in cardiac pacemaking as the current produced by stimulating Hcn channels expressed in Chinese hamster ovary (CHO) cells across the voltage range of If was small and inconsistent with a membrane clock pacemaking-dependent model. In both the brown trout and the pacific hagfish, Hcn3 was found to be more highly expressed than Hcn4 in the atrium and the ventricle and was the highest expressed Hcn isoform in the SAN of the brown trout (Hassinen et al., 2017; Wilson and Farrell, 2013). However, only Hcn4 expressed in CHO cells produced a current when stimulated across the voltage range of If (Hassinen et al., 2017). Hcn3 was also the highest expressed Hcn transcript in the SAN in the rainbow trout (Sutcliffe et al., 2020).   73 HCN channels, carry the cardiac pacemaker current (If), which is a mixed inward Na+ and K+ current (DiFrancesco, 1993; Hassinen et al., 2017; Ludwig et al., 1999; Moosmang et al., 2001; Pape, 1996; Robinson and Siegelbaum, 2003; Schweizer et al., 2009). Four HCN subtypes (HCN1-4) have been identified and functionally expressed in HEK293 cells and were shown to have very different activation kinetics (Moosmang et al., 2001). Although multiple ion channels may be expressed in cardiac tissues, channel activation and kinetics differ between isoforms and were shown to vary with temperature and hypoxia in the turtle heart (Stecyk et al., 2007), including Hcn channels (Stecyk, personal communication).  In mammals, the membrane clock is the major cardiac pacemaking mechanism and the calcium clock plays a reduced, secondary role and is not able to generate APs alone (Bers, 2002; Shinohara et al., 2010). In the pacemaker region of mammals, HCN4 is considered the principal carrier of If due to its high expression levels (Brioschi et al., 2009; Ishii et al., 1999; Stieber et al., 2003b; Stieber et al., 2004; Wahl-Schott et al., 2014). The importance of HCN4 in mammals is demonstrated by cardiac-specific Hcn4 knockout mice which have an 85% reduced If and the knockout was embryonically lethal between days 9.5 and 11.5 (Harzheim et al., 2008; Stieber et al., 2003b). Furthermore, a tamoxifen-inducible HCN4 knockout in adult mice resulted in no difference in fH but induces sinus pauses and reduces the If current in SAN cells, which had functional characteristics of the HCN2 and HCN1 currents (Herrmann et al., 2007). Taken together, these data suggest that HCN4 plays a vital role in mammalian development and a major role in cardiac pacemaking.  Given the importance of HCN4 for pacemaking in mammals, the apparently variable role of Hcn channels in cardiac pacemaking among fish, and the unknown relevance of Hcn channel isoforms in fish cardiac pacemaking, this chapter focuses on the role of Hcn4 in cardiac pacemaking of the zebrafish. While little is known about cardiac pacemaking mechanisms in fish, zebrafish are emerging as a promising model organism due to their amenability to  74 knockout technologies, their entirely sequenced genome and the high level of gene function conservation between humans and zebrafish. The transparency of the zebrafish embryo provides insight into cardiovascular development and its small size allows the embryo to pursue development despite severe developmental defects of the cardiovascular system, which would result in premature death of mammalian models (Bakkers, 2011). For example, the slow mo mutant zebrafish, which carry a defective pacemaker current which significantly reduces fH by 37% and reduces If current by 85%, is not a lethal mutation (Baker et al., 1997; Warren et al., 2001). However, the underlying genetic components of the slow mo mutant zebrafish have not been identified (Burkhard et al., 2017). Nevertheless, direct evidence of the functional significance of Hcn4 in cardiac pacemaking remains to be demonstrated. However, pharmacological agents specific to each of the zebrafish Hcn channels have yet to be identified. Therefore, the CRISPR-Cas9 system was used to generate a homozygous line of Hcn4 knockout zebrafish to determine the role of Hcn4 in cardiac pacemaking. The CRISPR-Cas9 system is a genome-editing tool derived from a prokaryotic immune system that is highly versatile and precisely targets and cleaves target genes (Jinek et al., 2012; Sander and Joung, 2014). A short single guide RNA (sgRNA) of 20 base pairs, complexed to the Cas9 endonuclease of Streptococcus pyogenes, binds to the complementary target DNA guiding the Cas9 complex to the target site. The Cas9 endonuclease cleaves the target DNA downstream of the protospacer adjacent motif (PAM) site resulting in a double-stranded break at the target genomic locus, which is subsequently repaired by error-prone non-homologous end joining (NHEJ) repair machinery of the cell, which can lead to insertions or deletions (INDELs) (Kosicki et al., 2018). The resulting gene inactivation, caused by INDELs, frameshifts in the coding sequence and premature stop codons, can lead to nonsense-mediated decay of mRNA or truncated, dysfunctional proteins (Tuladhar et al., 2019).  75  Thus, the aim of this study was to assess cardiac performance and thermal tolerance in a hcn4 knockout zebrafish line utilizing electrocardiogram (ECG) recordings, swimming trials, respirometry, and critical thermal maximum (CTmax) trials to provide definitive insight into the role of Hcn4 in cardiac pacemaking and determining thermal tolerance of the zebrafish. Given the important role of HCN4 played in mammals and the phenotype of the slow mo mutant zebrafish, I hypothesized that hcn4 mutant zebrafish would have a reduced maximum heart rate (fHmax), limiting swimming performance and reducing thermal tolerance and ṀO2. Using the CRISPR-Cas9 system, I generated a homozygous line of Hcn4 knockout zebrafish and assessed the role of Hcn4 in cardiac pacemaking of the zebrafish.                76 3.3 Materials and methods All procedures adhered to the United Kingdom Home Office Animals Scientific Procedures Act of 1986 and were granted authority under project licenses P005EF9F9 and 70/9091.   Animal husbandry Zebrafish were housed at the animal holding facility at The University of Manchester in recirculating aquaria and were maintained at 28°C with a 14 h:10 h light:dark photoperiod. Fish were fed twice per day to satiation.      In-house bred wild-type zebrafish and the SqEt33Mi59B reporter line which express EGFP in pacemaker cells under the influence of the promotor of fibroblast growth homologous factor 2a (fhf2a) (Poon et al., 2016) were used to generate the knockout line to ease the identification of pacemaker cells for any subsequent cellular work out of the scope of this thesis. All fish were three months of age at the time of all experiments and the parental SqEt33Mi59B line was used as wild-type controls for all experiments. At the time of experimentation, hcn4 knockout fish were third generation, whilst the control fish were second generation.   Breeding and rearing of embryos Fish were bred by placing one male and one female in a flowthrough breeding tank overnight, separated by a partition. Shortly after the light is switched on in the morning, the divider was removed, the water was partially drained, and the tank was placed at an incline to create a shallow segment to promote laying. Fertilized embryos were collected from the bottom of the tank, separated from the adult fish by a false bottom meshing. Embryos were raised at  77 28°C in E3 standard medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, 0.00001% (v/v) Methylene Blue).  sgRNA design Three sgRNAs targeting exon 1 of the hcn4 gene were designed using the webtool “CHOPCHOP” (https://chopchop.cbu.uib.no) (Labun et al., 2019). The zebrafish hcn4 coding sequence (Ensembl(GRCz11):ENSDARG00000061685) was used as the target gene and was obtained from Ensemble (http://uswest.ensembl.org/index.html). CHOPCHOP parameters were set to Danio rerio (DanRer11/GRCZ11), the nucleotide length set to 20 base pairs, starting with GG and the PAM set to NGG. Finally, the efficiency score was calculated using the Doench efficiency score  (Doench et al., 2014).  The target sequences were reviewed and chosen according to their position in the sequence, their efficiency score and the number of off-targets to exclude unwanted off-target cleavage. Three sgRNAs were designed to target different sites in the gene (Table 3.1). To the sgRNA primers were added the T7 promoter sequence (5’ - TAATACGACTCACTATA - 3’) and the tracerRNA (5’ – GTTTTAGAGC – 3’) to the 5’ and 3’ ends of the target sequence respectively. The final sequences of the sgRNAs are provided in Appendix 3.1 and were obtained as DNA oligonucleotides from ThermoFisher (Paisley, UK).           78 Table 3.1. A list of primers used for generating sgRNA (sgRNA primers), and PCR primers.    sgRNA synthesis The sgRNA oligomers were synthesized from single-stranded DNA oligomers and annealed with a CRISPR single guide (CRISPRsg) universal reverse primer that codes for the entire scaffold sequence in reverse complement by heating to 100°C and then slowly decreasing the temperature to 50°C. The double-stranded DNA was then amplified by PCR with pfusion high-fidelity DNA polymerase (NEB; M0531) and the resultant PCR product was purified from agarose gel using QIAEX II gel extraction kit (QIAGEN; 20021). The purified DNA templates were used for in vitro transcription using Ambion MEGAscript T7 transcription kit (ThermoFisher; AM1334) following the manufacture’s protocol. Following in vitro transcription of RNA, 1 μl DNase was added and incubated at 37°C for 10 mins to remove the DNA template. Finally, RNA was precipitated by using sodium acetate (3M, 1/10 volume) and two volumes of 100% ethanol at -20°C for 15 min followed by 15 min centrifugation at 13,000 rpm. The RNA pellet was resuspended in 15 μL of RNase-free sgRNA primers Primer Sequences (5’-3’) Predicted off targets  Predicted efficiency  sgRNA 1 GGGCAGAAAGCGTCCATCAT 0 55.62 sgRNA 2 GAAGCCTTTACCGACCAGCG 0 67.58 sgRNA 3 GGCATCCCGGATCAGCAGGC 0 48.à4 PCR primers Primer Sequences (5’-3’)   sgRNA 1/2 Forward ATGGACAGGTTGCATTCGTC   sgRNA 1/2 Reverse CGACTGGTGATGGACGAAAG   sgRNA3 Forward TCTGATCACGGACGGAGAG   sgRNA 3 Reverse CGAACTGCCTGTGCATGAAG    79 water and was stored at -20°C until used. Final sgRNA and the CRISPRsg universal reverse primer are detailed in Appendix 3.1.    Generation of a hcn4 mutant zebrafish line 3.3.5.1 Construct injection in one-cell stage zebrafish embryos  Wild-type zebrafish were bred, and the fertilized embryos were collected immediately after being laid. sgRNAs (30 pg), Cas9 protein (250 pg), and H2B-Cerulean3 (200 pg) were microinjected into the yolks of single-cell stage embryos. Borosilicate micropipettes (Sutter Instruments; BF-150-110-10, Novato, CA, USA), made using a one stage Sutter P-97 microelectrode puller (Sutter instruments), were used for the microinjection. The tip of the pipette was broken off with forceps prior to the injection, resulting in 50 pL of the construct-containing solution being released with each injection. Fluorescence was used to identify successfully injected embryos at 24 h post-fertilization. Embryos were raised at 28°C in E3 standard medium until 5 days post-fertilization before being transferred to the University holding facility and raised to adulthood.   3.3.5.2 Identification of INDELS in founder generation (F0) fish  Fish were raised for eight weeks before a tissue sample was taken from the caudal fin. DNA was extracted for screening of allele disruption by PCR followed by agarose gel electrophoresis. Briefly, 100 μl NaOH (50 mM) was added to the tissue sample and was incubated at 95°C for 20 mins. 40 μl Tris-HCl (1 M) was then added followed by a brief centrifugation. Finally, 1 μl of this extract was added to the PCR reaction following the cycling protocol (94°C for 3 mins, 35 cycles between 94°C for 45 s and 55°C for 30 s, 72°C for 90 s) using MyTaq red mix (Bioline; BIO25043). The primers used for the screening PCR are listed in Table 3.1. The PCR product was separated by 3% agarose gel electrophoresis where smears  80 indicated allelic disruption. Fish presenting allelic disruption were reared to sexual maturity and formed the F0 generation, which were subsequently outcrossed with the sqET33Mi59B reporter line.   3.3.5.3 F0 outcross and first-generation (F1) incross First-generation (F1) fish were generated by outcrossing fish F0 carrying a deletion in hcn4 with the SqEt33Mi59B reporter line. The offspring were raised to adulthood and propagation of the disrupted allele was verified by DNA extraction, PCR and gel electrophoresis. Sanger sequencing was performed for heterozygote F1 fish to identify frameshift mutations in the mutated allele. Fish with gene disruption were incrossed and each second generation (F2) germline was screened for homozygous hcn4 mutations. Mutations were identified in individual fish by PCR and gel electrophoresis and high-resolution PCR (QIAxcel DNA Fast Analysis Kit) and Sanger sequencing was performed on fish presenting homozygous mutations. An overview of the CRISPR-mediated generation of a hcn4 knockout zebrafish line workflow is shown in Figure 3.1.  81   Figure 3.1. Schematic overview of the CRISPR Cas9 workflow for the generation of a hcn4 knockout zebrafish.  82  Knockout validation by Western blot  Total protein was extracted from 100 wild-type 5-days post-fertilization (dpf) embryos and 100 F2 hcn4 mutant 5 dpf embryos sacrificed by 500 mg L-1 MS-222 overdose. Tissues were lysed in radioimmunoprecipitation assay (RIPA) buffer [50 mM Tris-HCl, 150 mM NaCl, 0.1% sodium dodecyl sulfate (SDS), 1% NP-40, 1 mM phenylmethanesulfonyl fluoride (PMSF) and 0.5% N,N’-dicylohexylcabodiimide (DCC)] supplemented with protease inhibitor cocktail (G6521; Promega). Equal amounts of protein were loaded onto precast 10% Bis-Tris SDS-polyacrylamide gels (XP0301BOX; Invitrogen). Proteins were then transferred to nitrocellulose membranes using the iBlot dry blotting system (IB21001; Invitrogen) following the manufacturer’s instructions. Membranes were incubated in TBST (20 mM Tris, 150 mM NaCl, 0.1% tween, pH 7.5) containing 3% BSA for 1 h at room temperature, followed by incubation with a polyclonal anti-HCN4 antibody (1:200; APC-052, Alomone labs) overnight at 4°C. This antibody has previously been used for detection of hcn4 in the sinoatrial valve region of the goldfish and the zebrafish (Newton et al., 2014; Stoyek et al., 2015). Subsequently, membranes were incubated with donkey anti-rabbit IgG HRP conjugate (GE Healthcare, NA9340-1ML; 1/10 000 dilution) for 1 h at room temperature. Proteins were visualized using the Bio-Rad ChemiDoc XRS imaging system (1708265; Bio-Rad) after adding ECL substrate (1705061; Bio-Rad). Rabbit anti-ERK1/2 (9102; Cell signaling technology) was used as the loading control.    83  Real-time quantitative PCR (RT-qPCR) Primers for real-time (RT)-qPCR were designed using the webtool “Primer3web” (http://primer3.ut.ee) and are listed in Table 3.2. Hearts were removed from six adult zebrafish before being separated into different chambers (SAN, atrium, and ventricle) and stored in QIAzol (79306; QIAGEN). The samples were subsequently pooled by cardiac chamber for RNA extraction. Samples were homogenized using Precellys tissue homogenizer and Precellys homogenization tubes (Bertin instruments). RNA extraction was completed using the RNeasy mini kit (74104; QIAGEN) following the manufacturer’s instructions and included a DNase step. First-strand cDNA synthesis was performed using the ProtoScript II First Strand cDNA Synthesis Kit (E6560; New England BioLabs). Reactions were prepared using the SensiFAST SYBR qPCR kit (35053; Bioline) according to the manufacturer’s instructions and run on an Mx3000/Mx3005P Real-Time PCR System (Agilent) in triplicate. Cycling conditions for qPCR-reactions were denaturation at 95°C for 20 s, followed by 50 cycles at 95°C for 30 s, and 57°C for 45 s and 72°C for 45 s. β-actin was used as the housekeeping gene and the primers are listed in Table 3.2.        84  Table 3.2. A list of primer pairs used for qPCR. ENSDARG identifier numbers are listed for each of the genes. qPCR primers Primer Sequences (5’-3’) hcn1    (ENSDARG00000104480.2)  Forward  CTTTCACGCCTCATCAGATACATC Reverse GATTAAAGATTCTTACCACCGCACTC hcn2b (ENSDARG00000061665)   Forward  AGTATCAGGAGAAGTACAAGCAAGTT Reverse TTCCCTGGTATCTGTGCTCATAATAG hcn3 (ENSDARG00000027192)  Forward  CCACTAAAAGAGGAGATCGTGAACTA Reverse TGAAAGACCTCGAAACGTAACTTTG hcn4 ENSDARG00000061685  Forward  AGTATCAGGAGAAGTATAAGCAGGTG Reverse CAGAATGCTCTCTTCATCAAACATCT β-actin ENSDARG00000037746 Forward TGCGTCTGGATCTAGCTGG Reverse TCCCATCTCCTGCTCGAAG  85  Critical thermal maximum (CTmax) trials  Three mixed trials, using fifteen wild-type and fifteen Hcn4 mutant fish per trial (n=49 and n=43 for wild-type and mutant fish, respectively) were performed using a glass aquarium filled with 10 L of 28°C aerated water (oxygen maintained >80% air saturation). Wild-type and Hcn4 mutant fish were separated by meshing and water could freely circulate between these arena sections. Two small water pumps circulated water past a heater/chiller-controlled steel heat exchanger coil, which ensured constant mixing of water, maintaining homogenous water temperature (<0.1°C) throughout the fish arena, which was separated from the heater and pumps by a mesh. Pumps were positioned to reduce water flow to a minimum, and the temperature was monitored using a temperature sensor (accuracy ± 0.1°C) coupled to an oxygen and temperature sensing unit (Fibox 3 trace, PreSens). Fish were held at 28°C for 30 min before the temperature was increased by 0.3°C per minute until the occurrence of a 2 s loss of equilibrium (Becker and Genoway, 1979) upon which fish were removed from the tank and culled by lethal overdose of MS-222. Hearts were excised and weighed to compare relative ventricular mass between groups. CTmax trials were used as the temperature at which locomotion becomes disorganized and the fish loses its ability to maintain its upright position, it loses its ability to escape lethal conditions (Beitinger et al., 2000) and is thus an incipient lethal temperature. However, the exact mechanisms by which loss of equilibrium occurs remains elusive (Jutfelt et al., 2019).     86  Swimming trials Fish (n=12 and n=11 for wild-type and mutant fish respectively, all males) were individually placed in a swim tunnel (SW10100; Loligo systems) fitted with an 800 mL cylindric swim flume and filled with dechlorinated water maintained at 28°C and >80% air saturation. Fish were held in the swimming flume for 30 min with no current. Current velocity was then set to 16 cm s-1 and maintained for 30 min before starting the swimming trial. The water velocity was then increased incrementally by 3 cm s-1 every 5 min until the fish voluntarily ceased to swim and were pushed against the back grid meshing for more than 2 s. The flow was then stopped, and the fish were removed from the swimming apparatus for full recovery in 28°C oxygenated water. Maximum speed, total swimming time and time spent at the fastest swimming speed were recorded. Critical swimming speed (Ucrit) was calculated as:  𝑈𝑐𝑟𝑖𝑡 = 𝑈𝑓 + [𝑈𝑖(𝑇𝑓 ∗ 𝑇𝑖−1)]  Where Uf is the highest sustained velocity completed by the fish, Ui is the increment in water velocity, Tf is the time the fish endured at the final swimming speed, and Ti is the time interval of each swimming speed (Brett, 1964).    87  Respirometry Intermittent-flow respirometry was used to measure ṀO2 as a proxy for metabolic rate in four fish simultaneously (wild-type n=12, Hcn4 knockout n=11). Each fish was housed individually in a 100 mL, custom-built, plexiglass respirometry chamber (Steffensen, 1989). The four chambers were submerged in a 30 L water reservoir that was continuously aerated and temperature-controlled (28 ± 0.3°C, ITC-308; Inkbird Tech. Co., Ltd., Shenzhen, China) and continuously mixed to maintain stable environmental conditions throughout the trial. Each respirometry chamber contained a recirculating loop that used a pump to ensure mixing of the water within the chamber and a constant flow over the oxygen sensor (PyroScience, Aachen, Germany) (Svendsen et al., 2016) and dissolved oxygen was recorded using a FireSting oxygen sensor (FS02-4; PyroScience). Bacterial ṀO2 measurements were performed for a minimum of 30 min before and after fish ṀO2 measurements. To achieve a maximum metabolic rate measurement, fish were individually chased to exhaustion for 3 min before being weighed for a standardized 30 s period and then transferred to their respirometry chamber. Continuous ṀO2 measurements were taken for a minimum of 14 h, during which fish were left undisturbed and in the dark (Chabot et al., 2016) with air saturation inside the chamber never decreasing below 80%. AquaResp 3.0 (Denmark, Aquaresp.com) was used to automate each respirometry cycle, which comprised a 75 s flushing phase, 120 s wait period, and a 300 s ṀO2 measurement phase. A scheme of the respiratory setup can be found in Figure 3.2. The ṀO2 data were analyzed in R 3.6.0 (R Foundation for Statistical Computing, Vienna, Austria) using the package FishResp 1.0.4 (Morozov et al., 2019). Oxygen consumption measurements were corrected for microbial respiration by removing the mean ṀO2 of the pre- and post-test background measurements from the total oxygen consumption rate. Only slopes with an  88 R2 > 0.95 were used to calculate metabolic rate. Standard metabolic rate (SMR) was estimated using the 10% of slopes with the shallowest gradient and maximum metabolic rate (MMR) was estimated as the single slope with the highest gradient, and absolute aerobic scope (AAS) was calculated by subtracting the value of SMR from the value of MMR (Fry, 1947; Fry, 1971).        89    Figure 3.2. Schematic of the respirometry setup. (1) oxygenation pump (2) zebrafish in closed 100 mL plexiglass respirometry chamber (3) recirculation pump (4) outflow circuit (5) fiber optic oxygen probe (6) FireSting oxygen sensor (7) mixing pump (8) flush pump (9) temperature probe (10) heater (11) temperature controller (12) computer. 90  Electrocardiogram (ECG) recordings ECG recordings were taken as previously described in chapter 2 and as described in published works (Casselman et al., 2012; Marchant and Farrell, 2019; Sidhu et al., 2014). Briefly, maximum fH (fHmax) was measured in anesthetized fish (80 mg L-1 MS-222 buffered with 50 mg L–1 sodium bicarbonate) maintained at 28°C while the gills were continuously irrigated with aerated water (>80% saturation) containing a maintenance dose of buffered anesthetic (80 mg L–1) MS-222. Intraperitoneal injections of atropine sulfate (1.2 μg g–1) and isoproterenol HCl (7.8 ng g–1) were used to achieve fHmax. A custom-made ECG probe (30 AGW copper wire inside a glass pipette) was positioned on the ventral midline of the heart, and its position was adjusted to achieve a clear ECG signal with a reference electrode placed distally to the recording electrode on the peduncle of the fish. The ECG signal was amplified (Grass P55 AC, Astro Nova Inc.) and recorded using Powerlab 8/35 (ADInstruments Inc.). The temperature was controlled using two heater/chillers, and the water temperature was recorded using a temperature sensor (accuracy ± 0.1°C; Fibox 3 trace, PreSens). After 30 min of stable ECG recording at 28°C, the water temperature was incrementally increased by 1°C every 5 min until signs of cardiac arrhythmia and cardiac failure which developed at high temperature upon which fish were removed from the experimental tank and euthanized by lethal overdose of MS-222 and severing the spinal cord. Analysis was performed off-line using LabChart 8 (ADInstruments, Colorado Springs, CO, USA) to determine the peak fHmax before decline (peak fHmax), the temperature at which the fish reached peak fHmax (Tmax), and the temperature at which a cardiac arrhythmia (Tarr) was first observed (n=5 wild-type; n=3 hcn4 mutant fish). For each fish, fHmax was determined using the R-R interval at each experimental temperature over a 1 min period of the ECG recording (LabChart 8.0). Peak  91 fHmax was defined as the highest fHmax recorded during the warming protocol, and Tmax was assigned at the temperature of peak fHmax. To assess the role of hcn4 in cardiac pacemaking in zebrafish, Hcn channels were pharmacologically blocked with zatebradine, a non-specific Hcn channel blocker (Goethals et al., 1993). Following injection of atropine and isoproterenol, fish were given an intraperitoneal injection of zatebradine  (4 μg g–1) (n=6 wild-type; n=7 hcn4 mutant fish) to block cardiac Hcn channels (Altimiras, 2004) or ryanodine (50 ng g–1)  (n=5 wild-type; n=6 for hcn4 mutant fish) combined with thapsigargin (1.3 μg g–1) to block RyRs and SERCA pumps, respectively, before being placed in the recording chamber for 30 min and until a stable fH was attained (Marchant and Farrell, 2019). All chemicals were obtained from Sigma-Aldrich (Gillingham, UK), with the exception of zatebradine (Cayman Chemicals; Cambridge, UK). Stock solutions of zatebradine and ryanodine were made in dimethyl sulphoxide (DMSO); isoproterenol and atropine directly in saline; thapsigargin was prepared in ethanol. All drug stock solutions were diluted with saline solution (mM: NaCl 124.2, KCl 5.1, Na2HPO4, MgSO4 1.9, CaCl2 1.4, NaHCO3 11.9; pH 7.2) to injectable concentrations and administered by intraperitoneal injection with a final volume ≤ 20 μL (equivalent to 5% of the fish’s body mass).        92  Statistical methods Values are given as means ± standard error of the mean (SEM). Ventricular mass, MMR, SMR and AAS were compared using Student’s t-test. Hcn mRNA expression levels were compared using a two-way ANOVA followed by Sidak’s multiple comparisons test. Statistical comparison of ECG data was performed using a multiple t-test using Holm-Sidak multiple comparisons method for comparisons of wild-type vs. hcn4 mutant fish. When untreated control fish were compared with drug-treated fish, statistical comparison was performed by fitting a mixed model with Geisser-Greenhouse correction, followed by a Sidak post hoc test, using P ≤ 0.05 as the level of significance.  All statistical analyses were performed using GraphPad Prism 8.0 or SPSS version 27 using P ≤ 0.05 as the level of significance.             93 3.4 Results  Introduction of a premature stop codon into zebrafish hcn4 abrogates protein expression  Sanger sequencing was used to confirm the frameshift mutation generated by double-stranded CRISPR-Cas9 targeted cleavage in exon 1 of the hcn4 gene. The non-homologous end-joining repair of the CRISPR-Cas9-mediated DNA breaks led to a 21 bp deletion and a 6 bp insertion, resulting in a 15 bp deletion downstream of the protospacer adjacent motif (PAM) site of the sgRNA. Importantly, a premature stop codon was introduced at position 470-473 of the first exon, which was verified in all F2 hcn4 mutant fish by genotyping and Sanger sequencing (Figure 3.3). The early position of the premature stop codon in the coding sequence resulted in a non-functional truncated protein (Appendix 3.2, Appendix 3.3). The screening results from each successive generation to arrive at the hcn4 mutant population, carrying a premature stop codon in exon 1 are detailed below.         94  Figure 3.3. Schematic diagram of CRISPR hcn4 knockout in zebrafish. Schematic diagram of the hcn4 gene (ENSDARG00000061685) showing the targeting position of the single guide RNA (sgRNA) in the first exon of hcn4. The sgRNA (shown as a green line and text, central panel) was designed including a crRNA complementary to a region within exon 1 of the hcn4 gene (blue boxes, green boxes, lines, and arrowhead indicate untranslated regions, introns, exons, sgRNA target site; top panel) and tracrRNA. The crRNA sequence is indicated in green, the wildtype sequence in black, the PAM site in blue and the site of the double-stranded break is indicated by the orange arrows. The Cas9 protein is in grey. The bottom panel shows the deletion and insertion mutations, respectively, with the novel deletion indicated by hyphens. The novel insertion is indicated in bold with the novel stop codon underlined and in red. The PAM site is underlined in the novel deletion and insertion sequences as a point of reference.   95  Identification of INDELS in founder generation (F0) fish The presence of double bands or smearing in the DNA electrophoresis of DNA isolated from F0 fish indicated the presence of genetic disruption (Figure 3.4). Fish presenting smearing of the electrophoresis indicate allelic mosaicism and were selected for outcross with the sqET33Mi59B reporter line to select for a single mutation.     Figure 3.4. Agarose gel electrophoresis of DNA isolated from fin tissue of eight-week-old zebrafish. (A) Electrophoresis of fish injected with sgRNA1 at the one-cell stage. (B) Electrophoresis of fish injected with sgRNA3 at the one-cell stage. Smears on the gel indicate allele disruption. Wild-type DNA can be found in the first lane on the left and is indicated by WT. Allelic disruption is indicated by red boxes.        96  F0 outcross and first-generation (F1) incross of the mutant zebrafish F0 Fish outcrossed with the sqET33Mi59B reporter line resulted in double bands on the gel electrophoresis (Figure 3.5) and Sanger sequencing revealed a mixed read, confirming the gene disruption. F1 fish presenting gene disruption were incrossed to obtain homozygous F2 individuals.  Figure 3.5.  (A) Agarose gel electrophoresis of DNA isolated from fin tissue of zebrafish. Wild-type DNA band (WT) indicates the size of the wild-type allele. Double bands indicated by red boxes show heterozygous individuals carrying both wild-type and mutant alleles. (B) Sanger sequencing pherogram. Wild-type genotype in the top row and the mixed sequence carrying both the wild type and the mutated genotype in the bottom row. The mixed sequence in the bottom row is identical to the wild-type running 5’ to 3’ until the mutated region (indicated by the red arrow) downstream of the PAM site (TGG; dashed red box). Red boxes indicate the first 3 base pair differences between wild type and mixed sequences genotypes. Multiple peaks in the mixed sequence genotype indicate mutations at the Cas9 cleavage site.  97   Generation of homozygous hcn4 knockout zebrafish in F2    F2 homozygous fish presented a single discrete band, lower than the wild-type fish on the gel electrophoresis (Figure 3.6. A). A 15 bp deletion was identified through high-resolution electrophoresis and was confirmed with Sanger sequencing (Figure 3.6. B, C). The details of the genetic mutation are detailed in Figure 3.3. No difference in early mortality was observed between the wild-type fish and the hcn4 knockout fish, nor was there any difference in mortality beyond 5 dpf when the heart becomes necessary for blood circulation. Lastly, there was no mortality difference observed in the adult fish.        98  Figure 3.6. Validation of hcn4 knockout zebrafish line. (A) Agarose gel electrophoresis of DNA isolated from fin tissue of zebrafish. Wild-type DNA band (WT) indicates the size of the wild-type allele. Smears or double bands indicate heterozygous individuals. Discrete single bands are homozygous individuals for the mutant allele (15 base pair deletion) indicated by red boxes. 100 bp hyperladder (B) Sanger sequencing pherogram. Wild-type genotype in the top row and the mutant sequence carrying only the mutated genotype in the bottom row. The mutated sequence in the bottom row is identical to the wild-type running 5’ to 3’ until it arrives at the mutated region (indicated by the red arrow) downstream of the PAM site (TGG; dashed red box). The red dashed  99 line on the wild-type genotype indicates the primer region containing the deletion of the mutant allele. The green dashed line on the mutant sequence indicates inserted base pairs (TATTAA). The red box indicates the in-frame premature stop codon (TTA). (C) Genotyping knockout validation using high-resolution electrophoresis. *, wild-type sample; ‡, negative control; the remaining lanes contain mutant DNA carrying the 15 bp deletion.                    100  Validation of hcn4 knockout zebrafish line – Western blot and RT-qPCR Western blot analysis showed a clear loss of Hcn4 protein in the hcn4 mutant fish (Figure 3.7. A). In the SAN, hcn2, hcn3, and hcn4 mRNA levels were lower in hcn4 mutant fish compared with those in wild-type fish, while hcn1 mRNA levels were low in both hcn4 mutant and wild-type fish. Hcn2 was downregulated in the ventricle and the atrium of hcn4 mutant fish, while the mRNA levels of all other Hcn isoforms did not differ significantly between hcn4 mutant and wild-type fish in the ventricle (Figure 3.7. B-D).     Figure 3.7. Validation of hcn4 knockout in zebrafish. (A) Representative western blot analysis (performed in triplicate) for Hcn4 protein levels in wild-type and hcn4 mutant 5 dpf zebrafish embryos. B-D. Hcn1, hcn2, hcn3, and hcn4 mRNA levels in the hearts of 5 pooled adult wild-type and hcn4 mutant zebrafish, as determined by RT-qPCR. Hcn mRNA levels in (B) sinoatrial node, (C) atrium, and (D) ventricle were assessed. Two-way ANOVA followed by a Sidak’s multiple comparisons post hoc test;*, P = 0.037; ****, P ≤ 0.0001.   101  Hcn4 knockout zebrafish are indistinguishable from wild-type fish  Hcn4 mutant zebrafish were morphologically indistinguishable from the wild-type sibling animals (Figure 3.8. Α), displaying no difference in mean mass or general body shape (Figure 3.9. C). Although the ventricular mass was significantly larger in hcn4 mutant fish, when corrected for body weight the relative ventricular mass of hcn4 mutant zebrafish was not significantly different from that of wild-type fish as the mass of the Hcn4 fish was significantly larger than the wild-type fish (Figure 3.8. B-C).    Figure 3.8. Morphological appearance and ventricular mass of wild type and hcn4 mutant zebrafish. (A) Image of wild-type (WT; top) and hcn4 mutant (MT; bottom) fish (B) Ventricular mass of wild type and hcn4 mutant fish. ****, P ≤ 0.0001. (C) Mass of the wild-type and Hcn4 mutant fish, P = 0.0009. (D) Relative ventricular mass of wild type and hcn4 mutant fish ((heart mass / body mass) * 100). Wild-type n=10, Hcn4 mutant n=13. Values are mean ± SEM.  Statistical significance was determined using a Student t-test P ≤ 0.05.    102  Loss-of-function of hcn4 mutant increases critical thermal maximum (CTmax) resulting in heat-tolerant zebrafish I assessed critical thermal maximum (CTmax), the point at which locomotion becomes disorganized and the righting reflex is first lost during acute thermal ramping as a measure of thermal tolerance. The mean CTmax of hcn4 mutant zebrafish (40.5 ± 0.1°C) was significantly higher than in wild-type fish (39.7 ± 1.2°C; P = 0.0005; Table 3.3) although this may not have any biological relevance given the relatively small difference in temperature between the CTmax of wild-type and hcn4 knockout fish. Thus, mutant zebrafish tolerated higher temperatures for longer. Individual mutant zebrafish CTmax values ranged from 39.0°C to 42.1°C and the time-to-CTmax ranged from 48.5 to 70.2 min, while for the wild-type they ranged from 36.7 to 40.9°C and from 36.0 to 60.1 min (Figure 3.9 A-B). The mass of the fish used for the CTmax trials was not significantly different between the hcn4 knockout fish and the wild-type fish (Figure 3.9 C) and there was no difference between CTmax and the mass of the fish (appendix 3.4). Taken together, fish with a loss of function mutation in hcn4 were more resistant to heat-induced stress than wild-type controls.    103  Figure 3.9. Critical thermal maximum (CTmax) of wild-type and hcn4 mutant zebrafish. (A) Comparison of CTmax of wild-type (n=47) and hcn4 (n=43) mutant fish shown as violin plots with the median shown as a solid horizontal bar and the quartiles shown as dashed bars. Individual fish are represented as dots. ****, P ≤ 0.0001; Mann-Whitney U test. (B) CTmax of wild-type (n=47) and hcn4 (n=43) mutant fish across the incremental warming profile. Individual fish are represented as dots. ****, P ≤ 0.0001; Mann-Whitney U test. (C) Average weight (mean ± SEM) of wild-type and Hcn4 mutant fish used in CTmax trials. P = 0.675.        104  hcn4 mutant zebrafish maintained maximum respiratory capacity and swimming performance No significant difference in respiratory capacity was observed between the wild-type and the hcn4 mutant- zebrafish (SMR: 480.0 ± 28.2, mg kg-1 h-1, 523.8 ± 31.1 mg kg-1 h-1;P = 0.309; MMR: 930.1 ± 73.2 mg kg-1 h-1, 1010.8 ± 71.3 mg kg-1 h-1;P = 0.096; and AAS: 450.0 ± 62.7 mg kg-1 h-1, 585.7 ± 66.4 mg kg-1 h-1; P = 0.134; for wild-type (n=12) and hcn4 mutant (n=10) zebrafish respectively) (Figure 3.10. A-C; Table 3.3). No significant difference was observed for either Ucrit or the time spent at the final swimming velocity between wild-type (n=12) and hcn4 mutants (n=11) zebrafish (52.2 ± 0.9 cm s-1; 3.57 ± 0.61 min and 49.9 ± 1.6 cm s-1; 3.14 ± 0.72 min, respectively; Figure 3.10. D).      105  Figure 3.10. Comparison of swimming performance and respirometry of wild-type and hcn4 mutant zebrafish at 28°C. Comparison of respirometry parameters between wild-type (n=12) and hcn4 mutant fish (n=10). Violin plots show data distribution and data points show frequency distribution of individuals (median, solid line; 25 and 75 quartiles, dashed line). (A) Standard metabolic rate (SMR), (B) maximum metabolic rate (MMR), and (C) absolute aerobic scope. (D) Comparison of swimming performance (Ucrit) of wild-type (n=12) and hcn4 mutant fish (n=11).  Statistical significance for all parameters was determined using a Student t-test P ≤ 0.05.        106  Hcn4 mutant fish maintained heart rate during acute warming Cardiac performance was evaluated from ECG recordings during acute warming. At 28°C, before incremental warming had begun, there was no significant difference in mean fHmax between wild-type (188 ± 9 beats min-1; n=5) and hcn4 mutant (198 ± 11 beats min-1; n=5) (Figure 3.11). Incremental warming by 1°C progressively increased fHmax of both control and mutant fish until a peak fHmax was reached. With incremental warming, no significant difference in fHmax at common test temperatures was observed between wild-type (n=5) and hcn4 mutant (n=5) zebrafish across the entire warming profile, including the values for Tmax (35.7 ± 1.5°C and 33.2 ± 1.7°C, respectively; P = 0.786) and mean peak fHmax (238 ± 21 beats min-1 and 242 ± 12 beats min-1, respectively; P = 0.863; Table 3.3). Also, peak fHmax similarly ranged from 206 to 268 beats min-1 in individual hcn4 mutant fish and from 203 to 276 beats min-1 in wild-type fish. Further warming resulted in a decline in fHmax (Figure 3.11), with further warming leading to arrhythmia and, ultimately, asystole (Table 3.3). Despite these similarities, all hcn4 knockout fish maintained a rhythmic heartbeat up to 35°C, with only one fish persisting above 36°C and the heart became arrhythmic at 36.6°C. Of the wild-type fish, the first fish exhibited arrhythmia at 36.6°C and arrhythmia occurred in 80% of control fish at a mean temperature of 37.1°C ± 1.0°C; three fish had a rhythmic heartbeat up to the termination of the heating protocol. Tmax of wild-type was significantly lower than CTmax by 4.1°C (P = 0.0001) and even more so (7.3°C) for hcn4 mutant fish (P = 0. 0001). Tarr was significantly lower than CTmax by 2.6°C in wild-type fish (P = 0.007) and by 3.9°C in hcn4 mutant fish that presented arrhythmia.    107  Figure 3.11. Comparison of maximum heart rate (fHmax) of anesthetized wild-type (n=5) and hcn4 knockout fish (n=5). The mean fHmax (± SEM) of wild-type (n=5) and hcn4 knockout fish (n=5) during acute warming in 1°C increments. Dots are connected if no change in the number of individuals, numbers above the disconnected lines indicate the number of individuals. Statistical significance was determined using a mixed model with SPSS P ≤ 0.05.            108 Table 3.3. A comparison of cardiorespiratory and swimming performance of wild-type and hcn4 mutant zebrafish. Data are presented as mean ± SEM. *Indicates a significant difference at P ≤ 0.05; Students t-test.  AAS, Absolute aerobic scope; CTmax, Critical thermal maximum; fHmax, Maximum heart rate; MMR, Maximum metabolic rate; SMR, Standard metabolic rate; Tarr, Temperature of first cardiac arrhythmia; Tmax, Maximum temperature; Ucrit, Maximal critical swimming speed. Parameter n Wild-type zebrafish n Hcn4 mutant zebrafish CTmax (°C) 49 39.73 ± 1.16 43 40.52 ± 0.10 *      Peak fHmax (beats min-1) 5 238 ± 12 5 242 ± 21 Tmax (°C) 5 35.7 ± 1.54 5 33.20 ± 1.66 Missing QRS Temperature (°C) 5 36.99 ± 1.09 2 34.80 ± 0.75 Tarr (°C) 4 37.10 ± 1.04 1 36.64      SMR (mg O2 kg-1 h-1) 12 480.0 ± 28.2 10 523.8 ± 31.1 MMR (mg O2 kg-1 h-1) 12 930.1 ± 73.2 10 1110.8 ± 71.3 AAS (mg O2 kg-1 h-1) 12 450.7 ± 62.7 10 585.7 ± 66.4 Ucrit (cm s-1) 12 52.64 ± 1.40 11 54.07 ± 0.94   109  Effect of pharmacological blockade of Hcn channels and SR-Ca2+ cycling on fHmax Consistent with my previous findings (Marchant and Farrell, 2019), fHmax at 28°C in wild-type zebrafish decreased after pharmacologically blocking either Hcn channels or SR-Ca2+ cycling (Figure 3.12 A, B). Hcn channels were examined with zatebradine (4 μg g–1) which significantly reduced mean fHmax in hcn4  mutant zebrafish (P = 0.006) to the same degree as in wild-type fish (33.9% vs 35.9%, respectively). Incremental 1°C warming increased fHmax in all individuals between 28°C and 33°C before fHmax started to decline at 33°C in hcn4 knockout and at 34°C in wild-type fish (Figure 3.12. C). These results suggest that either Hcn4 does not contribute to cardiac pacemaking in the zebrafish, or that other Hcn channel isoforms had compensated for its function in hcn4 mutant zebrafish (Figure 3.12. C). Either way, peak fHmax was unchanged in the knockout fish, suggesting that Hcn4 is not required to attain maximal beating frequency in zebrafish and that the residual, or compensatory, Hcn channel may have similar kinetics to that of Hcn4 (Stieber et al., 2003b). SR-Ca2+ cycling was assessed by blocking RyRs and SERCA pumps with ryanodine and thapsigargin, respectively. At 28°C, fHmax of wild-type and hcn4 mutant zebrafish was reduced by 33.8% and 38.1%, respectively. Incremental warming by 1°C progressively increased fHmax in wild-type and hcn4 mutant zebrafish, until fHmax reached a peak value. No significant difference in fHmax was observed between wild-type and hcn4 mutant zebrafish between 28°C and 33°C (Figure 3.12. C). Mean fHmax of wild-type zebrafish started to decline at 33°C and was significantly lower than the hcn4 mutant zebrafish by 34°C, which continued to increase fHmax until 34°C (Figure 3.12 B, D). Cardiac arrhythmia and asystole occurred in all wild-type zebrafish by 35°C, whereas some hcn4 mutant zebrafish showed arrhythmia and asystole at a considerably higher temperature (~38°C) than in wild-type fish, suggesting a difference in thermal tolerance. Taken together, these  110 results suggest that SR-Ca2+ cycling plays a greater role in pacemaking in wild-type fish than hcn4 knockout fish at high temperature and that Hcn channels, other than Hcn4, took a more prominent role in the upper thermal tolerance of cardiac performance after hcn4 knockout. In addition, Hcn4 may serve as a depolarization reserve that has low thermal tolerance, and the activation voltage and conductance of other Hcn channels in the zebrafish heart may be close to those of Hcn4.           111  Figure 3.12. A comparison of maximum heart rate (fHmax) in wild-type and hcn4 mutant anesthetized zebrafish during acute 1°C incremental warming. (A) The mean (± SEM) fHmax in untreated wild-type fish (n=5) and wild-type fish treated with zatebradine (n=6) or ryanodine- and thapsigargin (n=5). * indicates significant differences between untreated vs. zatebradine-treated and ryanodine and thapsigargin-treated wild-type fish (P ≤ 0.05). ** indicates significant differences between untreated wild-type fish and zatebradine-treated wild-type fish (P ≤ 0.033). (B) The mean (± SEM) fHmax in untreated hcn4 knockout fish (n=5) and hcn4 mutant fish treated with zatebradine (n=7) or ryanodine and thapsigargin (n=6). * indicates significant differences between hcn4 knockout fish and zatebradine treated hcn4 knockout fish (P ≤ 0.05). ** indicates significant differences between hcn4 knockout fish and both zatebradine-treated and ryanodine- and thapsigargin-treated hcn4 knockout fish (P ≤ 0.033). § indicates significant differences between untreated hcn4 knockout fish and zatebradine-treated hcn4 knockout fish (P ≤ 0.05). (C) The mean (± SEM) fHmax in zatebradine-treated wild-type (n=6) and hcn4 knockout fish (n=7). (D)  112 The mean (± SEM) fHmax in ryanodine- and thapsigargin-treated wild-type (n=5) and hcn4 knockout fish (n=6). ** indicates significant differences between wild-type and hcn4 knockout fish (P = 0.0003).                    113 3.5 Discussion  This chapter assessed the role of Hcn4 in cardiac pacemaking and thermal tolerance of zebrafish by using CRISPR-Cas9 gene editing and tested the hypothesis that hcn4 knockout would result in reduced fHmax, reduced swimming capacity, and reduced thermal tolerance. In contrast to this hypothesis, hcn4 mutant zebrafish maintained fHmax, peak maximum fHmax, as well as oxygen uptake and swimming capacity. Intriguingly, hcn4 depletion improved upper thermal tolerance. These findings suggest that Hcn4 is not vital for cardiac pacemaking, and the function of Hcn4 in cardiac pacemaking is completely compensated, likely because of functional redundancy in Hcn channel isoforms, or due to the activation of yet unknown compensatory pacemaking mechanisms in hcn4 mutant zebrafish. Regardless of the Hcn4 compensation mechanisms, compensation contributed to improving upper thermal tolerance.  Cardiac pacemaking is maintained in hcn4 mutant fish Cardiac performance was assessed by elevating fHmax in anesthetized zebrafish to a peak by incrementally increasing the environmental temperature. Routine fH was not measured because vagal tone was blocked and β-adrenoceptors were maximally stimulated. Hence the major autonomic influences on the ECG were either blocked or standardized. This is because autonomic regulation can modulate acute cardiac thermal tolerance (Gilbert et al., 2019), possibly by increasing Ca2+ influx through T-type calcium channels, the sodium-calcium exchanger (NCX), as well as through adrenergic G-protein-coupled receptor stimulation of HCN channels (Monfredi et al., 2013; Xie et al., 2008), which means autonomic control may differ as a result of Hcn4 depletion. Regardless, hcn4 knockout did not affect fHmax over a wide temperature range preceding cardiac failure.   114   Hcn4 knockout induced no change to the calcium clock The mean reduction of fHmax induced by ryanodine and thapsigargin for the wild-type and the Hcn4 knockout zebrafish at 28°C was not different, indicating no compensatory increase in SR Ca2+ cycling.  Quantitatively, the decrease (between 28 and 40%) is in line with my previous finding  (Marchant and Farrell, 2019). Furthermore, after pharmacological block of the membrane clock with zatebradine, pacemaking driven by the SR alone shows no significant difference between the wild-type and the Hcn4 knockout fish. This may suggest that the SR is not significantly affected by the Hcn4 knockout and does not provide a compensatory depolarization mechanism for pacemaking.    The effect of Hcn4 knockout on the membrane clock Also, zatebradine block of Hcn channels reduced fHmax by ~30% in both wild-type and hcn4 mutant fish, indicating Hcn channels were important for pacemaking, and that the membrane clock is generated by multiple Hcn channels as zatebradine could still reduce fHmax of the hcn4 knockout fish. In addition, ryanodine and thapsigargin block of RyR and SERCA pumps reduced fHmax by up to 40% in both the wild-type and the hcn4 mutant fish, indicating an equal role of the calcium clock in the wild-type and hcn4 mutant fish that was unchanged by the hcn4 knockout. Taken together, these data suggest that the hcn4 mutant zebrafish express other Hcn isoforms to compensate for Hcn4 depletion. However, hcn4 knockout fish showed lower mRNA expression of other Hcn channels than the wild-type fish, suggesting that compensation by Hcn channels may involve increased trafficking of Hcn proteins to the membrane and increased heteromeric channel formation between other Hcn channels, namely Hcn1 and Hcn2, which have gating properties  115 characteristic of neither Hcn protein when forming separate heteromeric channels (Zhang et al., 2009). Although zatebradine has been reported to inhibit the rapidly activating delayed rectifier potassium current (Ikr) which resulted in the prolongation of the AP (Carmen et al., 1996), the effect on potassium channels at concentrations <5 ug L-1 is limited (<20% current reduction) compared to If current reduction (>90% reduction) (Van Bogaert and Pittoors, 2003). Given the low concentration (4 μg.g-1) used, the inhibitory effect of zatebradine on Ikr is limited and was not expected to significantly impact the QRS complex duration. In contrast to the data presented here for zebrafish, disruption of Hcn4 in mammals can lead to deep bradycardia, sinus arrhythmia, AV block, and asystole (Bucchi et al., 2012; Herrmann et al., 2007; Stieber et al., 2003b).  HCN4 is highly expressed in the mammalian SAN and plays a vital role in cardiac pacemaking (Chandler et al., 2009; Knaus et al., 2007). Indeed, Hcn4 mutant mouse embryos had a 75-90% reduced yet regular heart rate; the HCN channel blocker ZD7288 further reduced heart rate, although no arrhythmia was observed (Stieber et al., 2003b). Moreover, cilobradine, another HCN channel blocker, induced bradycardia in Hcn4 knockout mice and accentuated sinus pause durations of up to 10 s, further highlighting the importance of HCN channels and HCN4, in particular, in mammalian cardiac pacemaking for maintaining stable cardiac rhythm  (Herrmann et al., 2007). By rejecting my prediction that hcn4 knockout would lead to reduced fH in zebrafish I show that fish and mammals differ in their involvement of Hcn4 and other Hcn channels in pacemaking mechanisms.  One potential reason for the contradictory findings in fish and mammals is that the HCN blockers are not as effective in blocking all Hcn isoforms in fish as in mammals or may not result in a total block of the channels, regardless of the concentration. Alternatively, Hcn channels could play a reduced role in cardiac pacemaking in fish. Pharmacological block of Hcn channels in the  116 rainbow trout (Oncorhynchus mykiss) reduced fH by 50% (Gamperl et al., 2011) and similarly by 40-60% in the zebrafish, depending upon the test temperature (Marchant and Farrell, 2019). Furthermore, the If current recorded in pacemaker cells of the brown trout was very small (-1.2 ± 0.37 pA pF-1) and was deemed likely insufficient to initiate an AP (Hassinen et al., 2017). Therefore the role of Hcn channels in fish cardiac pacemaking is species-dependent and could vary from playing a limited or no role as seen in the brown trout, to being the only cardiac pacemaking mechanism as in the Pacific hagfish (Hassinen et al., 2017; Wilson et al., 2013). Nevertheless, the brown trout may possess a calcium clock-driven pacemaking system with little to no dependence on the Hcn-driven membrane clock because pharmacological blockade of the calcium clock components (ryanodine; RyR and thapsigargin; SERCA) in isolated pacemaker cells of the rainbow trout reduces AP rate by 44% in 18°C-acclimated fish (Hassinen et al., 2017; Haverinen and Vornanen, 2007). Further studies are needed to identify which specific Hcn channel subtypes are capable of producing the If current in zebrafish, as well as to determine the kinetics of these channels and their relative contribution to native If.   Alternative Hcn channels have a more prominent role in hcn4 mutant knockout zebrafish My data suggest a functional compensatory change to the pacemaking mechanisms in hcn4 knockout fish. After SR Ca2+ cycling was pharmacologically blocked, wild-type zebrafish had a reduced pacemaking capacity at high temperature that led to cardiac collapse at 33°C, whereas hcn4 knockout fish continued to increase fHmax until 36°C. Pacemaking in the hcn4 mutant fish could then be determined by alternative, kinetically rapid Hcn proteins capable of conducting If, likely Hcn2 (Ludwig et al., 1999; Mistrik et al., 2005), especially since Hcn1 had a low expression  117 in the zebrafish heart and Hcn3 has previously been shown to not produce an If current when stimulated across the voltage range of If in the brown trout (Hassinen et al., 2017). My data showed that in the SAN, Hcn2 is the most highly expressed isoform in both the wild-type and hcn4 mutant zebrafish and thus offers an alternative Hcn channel to drive pacemaking.  While my data suggest that zebrafish compensate for the knockout of hcn4, in mammals, knockout of Hcn4 has severe cardiac effects as HCN4 is widely established as the major HCN isoform driving If (Moosmang et al., 2001; Qu et al., 2002; Shi et al., 1999). For example, embryonic HCN4 knockout mice have a reduced (50% reduction) fH in the absence of functional If, but the heart does not stop beating until embryonic days 9.5 to 11.5 (Stieber et al., 2003b). Conditional HCN4 knockout in mice resulted in no difference in fH, but the mice developed sinus pauses, arrhythmia, and SAN cells had a reduced If current that had characteristics of currents generated by both HCN2 and HCN1 channels (Herrmann et al., 2007; Kozasa et al., 2018). Therefore, HCN4 may be important in embryonic development, including cardiac development in mice, but its role in cardiac pacemaking in adults may be compensated by other HCN channels as HCN4 is not the unique If current carrier in adult mice. Human HCN4 mutation led to bradycardia in a Moroccan Jewish population demonstrating the importance of HCN4 in pacemaking by showing the direct link between HCN4 and fH (Laish-Farkash et al., 2010). Furthermore, in humans, it has been shown that exercise can downregulate HCN4 channel expression reducing the resting fH (D’souza et al., 2014). A recent study in mice revealed 575 differentially expressed proteins in the SAN compared to the adjacent atrial tissue, some of which belonged to the membrane clock, but no proteins belonging to the calcium clock were found to be differentially expressed (Linscheid et al., 2019).   118 Previous studies in the zebrafish have shown the important role played by Hcn channels in cardiac pacemaking. The defective pacemaker current in the slow mo mutant zebrafish led to a 37% lower heart rate compared with that of control fish, linked to a reduced rapid component of the pacemaker current, likely produced by Hcn1/Hcn2, although the genetic origins of the mutation remain unidentified. Although Hcn4 is believed to be the major channels carrying the pacemaking current, these and my data, demonstrate the potential importance of other Hcn channels in pacemaking (Warren et al., 2001). The bradycardia linked to the depletion of the rapid component of If in slow mo zebrafish reduces over time, but fH does not reach the fH of wild type fish, likely because the fast If component cannot be compensated for by the slower kinetics of Hcn4 (Stainier et al., 1996). Investigating the genetic origins of the slow mo mutation could provide great insight into cardiac pacemaking using a naturally occurring mutation that dramatically reduces the If current and reduces fH in an age-dependent manner.  In a recent study, hcn4 knockout resulted in a significant increase in sinoatrial pauses in 2 and 5 dpf zebrafish embryos accompanied with sinoatrial arrest (von der Heyde et al., 2020). Despite these cardiac abnormalities, fH of the homozygous hcn4 knockout fish at 5 dpf had higher fH and lower fH variability than non-mutant zebrafish which may suggest that alternative Hcn channels possessing faster gating properties than Hcn4 are driving the membrane clock component of cardiac pacemaking in the Hcn4 knockout fish. When these knockout fish were exposed to ivabradine, heart rate variability increased in a dose-dependent manner and decreased fH (von der Heyde et al., 2020). Taken together, these data suggest that Hcn4 plays a more prominent role during early zebrafish development, whereas the zebrafish heart is less dependent on Hcn4 for cardiac pacemaking in later development. The findings are commensurate with my findings in the adult zebrafish heart which is less dependent on Hcn4 and alternative HCN subtypes can  119 compensate for the loss of functional Hcn4 channels, conserving beating frequency and increased thermal tolerance. As I did not evaluate the effects of the Hcn4 knockout during embryogenesis and through early development, I cannot compare my data with those of this closely related study. However, I can speculate that my hcn4 knockout fish may also have experienced cardiac events such as sinoatrial pauses during the early developmental stages which were not evident in the adult fish tested in my thesis work as compensatory mechanisms may completely compensate for the hcn4 knockout. Providing the increase in heart rate seen in this study on 5 dpf hcn4 knockout zebrafish embryos, it is likely that Hcn2 provides a Hcn-based compensatory pacemaking mechanism in the hcn4 knockout zebrafish and may be caused by Hcn RNA and protein regulatory mechanisms. Taken together, these studies along with mine, suggest greater plasticity of the zebrafish pacemaking mechanism than that of mammals with regard to Hcns which may occur due to compensatory mechanisms between Hcn channels. Functional expression studies have shown that HCN1, HCN2, and HCN4 form heteromeric channel formations (Zhang et al., 2009). Moreover, overexpression of HCN4 in rat ventricular myocytes resulted in increased levels of HCN3 mRNA and HCN2 knockdown in neonatal rat ventricular myocytes resulted in down-regulation of HCN4 mRNA (Zhang et al., 2009). In line with these findings, Hcn4 knockout fish exhibited a significant downregulation of Hcn2 and Hcn3 in the ventricle, and of Hcn2 in the atrium and the SAN, reflecting co-dependency of Hcn expression. More research is needed, including expression and functional studies of Hcn isoforms and electrophysiological characterization of wild-type and knockout zebrafish to resolve the importance of Hcn channels in pacemaking and the possibility that other Hcn isoforms compensate in the absence of Hcn4. Compensation (and/or increased activity of other Hcn subtypes) may have cascaded to improve whole animal thermal tolerance and may also be associated with changes in  120 SR density of Ca2+ cycling dynamics. Such a line of inquiry may require a multiple hcn isoform knockout due to functional redundancy of isoforms, caused by the three rounds of whole-genome duplication of zebrafish and other teleosts (Amores et al., 1998; Meyer and Van De Peer, 2005; Van de Peer et al., 2003), which may in part account for functional redundancy of Hcns in fish (McCluskey and Braasch, 2020). Furthermore, protein expression levels in hcn4 knockout fish and in multiple hcn knockout fish need to be quantified and future investigations should quantify calcium transients along with SR Ca2+ cycling as changes in the membrane clock may lead to changes in SR Ca2+ calcium handling or cell SR content. Moreover, kinetics and thermal tolerance of Hcn isoforms likely differ, and failure of Hcn4 at a lower temperature than other Hcn isoforms may cause pacemaking failure under normal conditions. However, pharmacological induction of fHmax for ECG experiments could increase peak fH and protect from heat-induced arrhythmia, due to the cardioprotective properties of β-adrenergic stimulation (Gilbert et al., 2019).    Hcn4 knockout induced no change to whole-animal aerobic respiration and swimming performance  No difference was found in swimming performance and whole animal respiration indices between wild-type and hcn4 knockout zebrafish. My Ucrit values are in line with previous findings of Ucrit between 50 and 65 cm s-1 for adult zebrafish swimming individually to exhaustion (Conradsen et al., 2016; Palstra et al., 2010; Plaut, 2000). My estimates of SMR (wild-type fish; 480.0 ± 28.2 mg O2 kg-1 h-1 and hcn4 knockout fish; 523.8 ± 31.1 mg O2 kg-1 h-1) are comparable to previous reports (Gerger et al., 2015; Thomas et al., 2013). Although my MMR estimates (wild-type fish; 930.1 ± 73.2 mg O2 kg-1 h-1 and hcn4 knockout fish; 1110.8 ± 71.3  mg O2 kg-1 h-1 ) were consistent with reported values using a similar exercise method (Lucas et al., 2016) they were  121 considerably lower than previous MMR estimates (~2400 – 2700 mg O2 kg-1 h-1) using protocols measuring MMR during exercise (Gerger et al., 2014; Thomas et al., 2013). Thus, the method that I and others have used may underestimate AAS, especially since my estimates of factorial aerobic scope (2) are lower than values reported for the zebrafish (~3-6) where MMR was obtained during swimming  (Gerger et al., 2015; Thomas et al., 2013). Alternative methods of exhaustive exercise, possibly reducing the time from the end of exercise to the start of MMR measurements (Zhang et al., 2020), may help resolve the MMR and AAR measurements in the zebrafish.   Role of Hcn4 in cardiac thermal tolerance of the zebrafish The higher thermal tolerance exhibited by the Hcn4 mutant zebrafish compared with the wild-type fish potentially indicates a low thermal tolerance of the Hcn4 protein compared with the putative compensatory pacemaking mechanisms. Low thermal tolerance of Hcn4 may ultimately cause a temperature-dependent decrease in pacemaking activity in the absence of β-adrenergic stimulation. In the rainbow trout, cardiac upper thermal tolerance has recently been suggested to be caused by depression of ventricular excitability due to increased leak current via potassium channels and failure of the sodium channel, potentially resulting in atrioventricular-conduction failure (Haverinen and Vornanen, 2020a). While this may hold true in the rainbow trout, which inhabits environments of temperatures ranging typically up to 20°C (Hokanson et al., 1977), it may not be true for the zebrafish, which inhabit waters reaching up to 38.6°C (López-Olmeda and Sánchez-Vázquez, 2011) a temperature well beyond the cardiac thermal maximum (25.3°C) reported for the rainbow trout and the failure of INa (20.9°C) (Haverinen and Vornanen, 2020a). Further research is needed on the mechanisms of thermal tolerance, temperature-dependent deterioration of cardiac pacemaking, and propagation of AP, as these are likely species-specific  122 and dependent on the thermal history of individuals and ion channel isoform expression (Abramochkin et al., 2019).   Limitations and conclusions In two independent studies, I have shown that zatebradine decreased fHmax in zebrafish by 40-60% (this chapter; Marchant and Farrell, 2019). I can exclude the possibility that zatebradine block of Hcn channels may have been incomplete in zebrafish. Furthermore, zatebradine non-specificity could have reduced fH by also inhibiting IKr. Also, the effects of ryanodine and thapsigargin are not pacemaker-specific because RyR and SERCA pumps are expressed throughout the myocardium. Inhibition of SR-calcium-induced calcium-release in cardiomyocytes could have reduced fH by interfering with AP conduction or propagation in either the atrium and/or ventricle. Lastly, β-adrenergic stimulation may have improved cardiac thermal tolerance via cAMP stimulation of Hcn channels in only the wild-type fish that possess Hcn4 channels. Caution is also needed in the interpretation of my respirometry results until the best method for measuring MMR in zebrafish is resolved.  I conclude that Hcn4 plays a reduced role in cardiac pacemaking compared to mammals and is not essential for cardiac pacemaking in the zebrafish heart, and may instead of providing the primary depolarizing pacemaking current, provide a depolarization reserve under autonomic control. In addition, pacemaking supported by Hcn4 may have a lower thermal tolerance than pacemaking supported by other Hcn channels. However, due to whole-genome duplication in teleost fishes, the zebrafish may possess an hcn4 orthologue encoding a functional protein capable of performing the same function as Hcn4. This hcn4l orthologue has previously been targeted alongside hcn4 in CRISPR-Cas9 knockout experiments in zebrafish embryos (up to 5 dpf) and sinus pauses and arrest were observed in 10.3% of embryos at 2 dpf and only 2.7% of embryos  123 showed sinoatrial pause at 5 dpf, however, no investigation was performed in adult fish (von der Heyde et al., 2020). Furthermore, when hcn4l was targeted in conjunction with hcn4, qRT-PCR data suggested a compensatory upregulation of Hcn1 and Hcn2 orthologues. The upregulation of transcripts of a similar sequence has been shown to be a typical response when targeting proximal sites for mutagenesis using the CRISPR-Cas9 system (El-Brolosy and Stainier, 2017). Therefore, hcn4 knockout may have led to an increase in hcn4l expression in my hcn4 knockout fish which was not quantified as hcn4l was believed to be a mislabelled hcn3 due to the gene synonym and relatively high sequence similarity of hcn4l with hcn3 (70.91% and 78.02% consensus cDNA similarity and protein sequence similarity respectively (Appendix 3.5). Furthermore, the protein sequence identity between the zebrafish Hcn4 and Hcn4l is relatively low (66.87%), lower than that of Hcn3 (79.86%) with large segments between nucleotides 677 and 1118 that are missing from the Hcn4l protein sequence compared with the Hcn4 protein sequence and the protein similarity score is also unexpectedly low for paralogues at 68.20% (Appendix 3.6; EMBOSS global alignment using Needleman-Wunsch algorithm). In addition, paralogues are known to have less functional overlap than with orthologues and thus sequence and protein similarity may not necessarily translate to similar function (Peterson et al., 2009) Therefore, further efforts are needed to verify function redundancy between Hcn4l with Hcn4 which would require functional expression analysis of the electrophysiological profile of Hcn4l in an expression vector to confirm the functional redundancy with Hcn4. The missing expression hcn4l data is a considerable limitation to my study and should be investigated. However, the hcn4 knockout line of zebrafish provides an excellent tool for investigating the functional redundancy of hcn4 in the zebrafish as a second knockout of hcn4l (or morpholino downregulation of hcn4l) would provide definitive  124 evidence for the functional redundancy of hcn4 and hcn4l and will allow the investigation of compensatory mechanisms among other Hcn isoforms, notably Hcn1 and Hcn2. Other limitations to the study include the confirmation of the knockout line by western blot analysis that only used a single antibody to confirm the reduction of Hcn4. Additionally, the current density of the If current was not recorded and would have provided electrophysiological evidence of the effect of the knockout which would have direct implications on pacemaking. Additionally, the knockout was confirmed by western blot where a single polyclonal antibody was used and two distinct bands were identified. Although both bands were absent in the knockout fish, I did not investigate further the origin of the second band, which may be a non-dominant splice variant of Hcn4 that is also absent in the hcn4 knockout fish.  Another limitation of this study is the limited sample size in some of the experiments performed (ECG recordings, respirometry, and swimming trials), affecting the ability of statistical tests to effectively detect significant differences between datasets. For example, mean MMR of the mutant fish is within the 75th percentile of the distribution of control data, suggesting a possible difference that may not have been detected due to the limited sample size. A retrospective power analysis revealed the required sample size was 29 fish per condition.  Although I have likely shown compensation among Hcn channels when Hcn4 is knocked out,  the Hcn isoforms that likely compensate for Hcn4 loss remain to be identified. Future research efforts should focus on Hcn characterization and functional expression analysis of zebrafish Hcn channels. In order to determine the compensatory mechanism involved in Hcn4 knockout fish, the first line of investigation should quantify Hcn2 protein and its involvement in calcium transients recorded in pacemaker cells.  Further, there is the need for a reduced sino-atrial preparation that is non-contracting to facilitate these cellular measurements. Lastly, changes in SR-Ca2+ cycling  125 cannot be excluded as another compensatory mechanism and may involve an increase in Ca2+ stores or its participation to calcium transients. I predict that cardiac pacemaking in fishes will show considerable species- and Hcn isoform-dependent variability, which is what appears to be emerging so far with very limited studies.  Chapters 2 and 3 have both provided novel insight into the mechanisms of cardiac pacemaking and have revealed that both the membrane and the calcium clock generate cardiac pacemaking. However, direct recordings of pacemaker cell activity are missing which would provide the most direct evidence of the relative importance of the membrane and calcium clocks to cardiac pacemaking. Several challenges, however, exist for recording AP from pacemaker cells directly, the greatest challenge being the physical movement associated with the contraction of the cardiac tissue. Chapter 4, therefore, provides a methodology for recording pacemaker APs using blebbistatin to effectively uncouple the excitation-contraction whilst preserving the excitation properties of the pacemaker cells and the AP waveform.     126  The effect of blebbistatin on the pacemaker action potential 4.1 Synopsis As an emerging model system of cardiac electrophysiology and human arrhythmias, cardiac pacemaking in zebrafish, which apparently involves two major pacemaking mechanisms, requires investigation beyond studies performed at the level of the whole heart, as in chapters 2 and 3. One line of research would be to investigate cardiac pacemaking directly in situ, which presents technical challenges, the greatest being physical movements associated with the contraction of the heart. Therefore, chapter 4 determined the effect on the zebrafish pacemaker AP of a known excitation-contraction uncoupling agent, blebbistatin, that has little effect on the cardiac AP in other species and assess its suitability as an excitation-contraction uncoupler for future electrophysiology research with zebrafish.           127 4.2 Introduction  The zebrafish has become a well-established vertebrate model of cardiac development (Arnaout et al., 2007; Bakkers, 2011; Bournele and Beis, 2016; Genge et al., 2016; Jensen et al., 2013; Liu and Stainier, 2012; Zon and Peterson, 2005) and has emerged as a model for cardiac electrophysiology, drug screening, and human cardiac diseases, including cardiac arrhythmias and their electrophysiological basis (Alday et al., 2014; Arnaout et al., 2007; Bakkers, 2011; Briggs, 2002; Chi et al., 2008; Howe et al., 2013; Milan et al., 2009; Verkerk and Remme, 2012; Vornanen and Hassinen, 2016; Vornanen et al., 2018). The rapid generation times, large number of offspring, ease of genetic manipulation, and its fully sequenced genome make the zebrafish an ideal model for exploring integrative physiology at a genomic scale. With approximately 71% of human genes having at least one orthologue in the zebrafish genome (Howe et al., 2013), including multiple genes encoding ion channels involved in cardiac physiology, such as NaV1.5 (Na+ channel), CaV1.2 (Ca2+ channel), and the KV4.3, KV7.1, and ERG  (K+ channels) (Hassel et al., 2008; Langheinrich et al., 2003; Novak et al., 2006; Rottbauer et al., 2001; Sanhueza et al., 2009), there is considerable genetic overlap between humans and zebrafish. Indeed, 47% of genes have a one-to-one homologue between the human and zebrafish genome, providing a platform for the investigation of human genetic diseases using targeted mutagenesis to generate disease models (Arnaout et al., 2007). Consequently, many features of higher vertebrate complexity are evident in the zebrafish including the embryonic formation of the heart (Chen et al., 1996; Jensen et al., 2013; Lieschke and Currie, 2007; Moorman and Christoffels, 2003; Moorman et al., 2007; Shih et al., 2015). Although the zebrafish heart is two-chambered, there are numerous functional similarities between zebrafish and human hearts including similar heart rate, AP duration, and AP morphology (Arnaout et al., 2007; Brette et al., 2008; Nemtsas et al., 2010) and its basic electrical properties  128 (Bakkers, 2011; Vornanen and Hassinen, 2016; Vornanen et al., 2018). For example, four of the five phases (0-4), of the mammalian AP are similar, the exception being phase 1 generated by the rapid transient outward potassium current (Ito), which is absent (or almost absent) in the fish heart (Alday et al., 2014; Vornanen et al., 2018). Furthermore, cardiac pacemaking and the cell-to-cell propagation of the AP through the atrium and ventricle of the zebrafish heart closely resemble that of the adult mammalian heart (Stoyek et al., 2016).  As in all vertebrates, pacemaking in the zebrafish heart is initiated in pacemaker cells of the SAN, a discrete ring of cells located at the base of the valves at the junction between the sinus venosus and the atrium (Saito, 1973; Stoyek et al., 2015; Tessadori et al., 2012; Vornanen et al., 2010). The AP is propagated as a voltage wave from the SAN throughout the myocardium, first to the atrium and then to the ventricle via cell-to-cell conduction, initiating coordinated and independent contractions of both cardiac chambers (Vornanen, 2016). Intercellular communication via gap-junctions that establish the electrical continuity between cells is of significant importance for the initiation of APs to establish the beating rate of cardiac myocytes (Bakker et al., 2010; Masahito et al., 1994; Shiels, 2017). Furthermore, pacemaker cells have significantly fewer intercellular coupling to allow AP initiation and delay impulse propagation, essential for rhythmic AP generation (Bakker et al., 2010). Therefore, preservation of cell interconnectivity of the SAN and atrial tissue is paramount for pacemaker AP investigation, presenting a technical challenge for recording the electrical activity from a single pacemaker cell in intact, contracting cardiac tissue. Consequently, such recordings are currently missing from the literature in part due to the difficulty in recording intracellular activity from spontaneously beating tissue.   129 High-resolution optical recordings using either fluorescent dyes or constitutively fluorescent calcium proteins are easy methods for recording calcium transients in pacemaker cells and they can be coupled with the intracellular recording of electrophysiological activity (Arnaout et al., 2007; Chi et al., 2008; Milan et al., 2009). However, both methods are sensitive to motion artifacts, with microelectrodes damaging the cell membrane upon cardiomyocyte contraction, causing depolarization of the membrane potential (ion leak) or cell membrane rupture. Contractions can be blocked with excitation-contraction uncouplers thereby removing this technical problem. Blebbistatin, 2,3-butanedione monoxime and cytochalasin D, are all excitation-contraction uncouplers, but the latter two alter Ca2+ handling, ion channel kinetics and modify the characteristics of the AP in a species-dependent manner (Jou et al., 2010; Kettlewell et al., 2004; Liu et al., 1993; Rueckschloss and Isenberg, 2001; Watanabe et al., 2001). Blebbistatin has been used in optical mapping studies in multiple species including zebrafish (Jou et al., 2010) mouse (Dou et al., 2007), rat (Farman et al., 2008; Fedorov et al., 2007), rabbit (Brack et al., 2013; Fedorov et al., 2007), as well as horse (Fenton et al., 2008), dog (Kong et al., 2009), and human (Fedorov et al., 2011) without significantly modifying the AP waveform. In the zebrafish embryo, blebbistatin has been shown to induce cardia bifida between 12 and 20 hpf when treated with concentrations as low as 5 μM blebbistatin (Wang et al., 2015) and totally inhibits the formation of mature furrows in early embryogenesis (40 min post-fertilization) inhibiting cell division (Gupta et al., 2017) as previously reported (Urven et al., 2006). Furthermore, application of 13.8 μM blebbistatin reduced cable constriction tension, leading to distortion of the embryo during epiboly (Chai et al., 2015). Internalization of the neural plate was also disrupted with 50 μM of blebbistatin as neural plate convergence was inhibited along with neural keel formation (Araya et al., 2019). In cardiac development studies, blebbistatin up to 10  130 μM L-1 induced cessation of contraction-induced cardiomyocyte enlargement, but did not significantly modify heart rate (Yang et al., 2014) and was also shown to dramatically decrease the contraction amplitude of the heart muscle in vivo, and thus stroke volume in embryos exposed to 20 μM, but did not modify heart rate (Várkuti et al., 2016). Finally, blebbistatin (10 μM) has previously been shown in 4-8 hpf zebrafish embryos to not significantly alter AP morphology or alter the spontaneous generation of APs (Jou et al., 2010). However, there has been no investigation into the effect of blebbistatin on cardiomyocyte AP in adult zebrafish and no studies, either in embryos of adults on the effect of blebbistatin on the pacemaker AP. Therefore, my primary goal was to use blebbistatin to uncouple excitation-contraction in a reduced adult zebrafish heart preparation and validate its use for intracellular recording of pacemaker cell electrical activity. Further, by comparing AP characteristics with and without blebbistatin at different temperatures, I open up the opportunity to study pacemaker electrophysiology at more than one temperature. Using a reduced preparation, I hypothesized that blebbistatin would stop cardiomyocyte contractions without significantly modifying the characteristic features of the pacemaker AP at any of my test temperatures.  Blebbistatin is a highly specific myosin II inhibitor that readily crosses the cell membrane and is a poor inhibitor of other members of the myosin family (Allingham et al., 2005; Limouze et al., 2004; Straight et al., 2003). It binds to the actin-binding region of the myosin motor domain, located in the large cleft motor domain (50-kDa cleft) in close proximity to the γ-phosphate-binding pocket of the ATPase active site (Allingham et al., 2005) where it exerts an allosteric effect and prevents the formation of strong actomyosin interactions (Limouze et al., 2004; Rauscher et al., 2018). The close proximity of the blebbistatin-binding site to the γ-phosphate-binding pocket results in the stabilization of the ADP-Pi complex, lowering myosin II affinity to  131 actin (Rauscher et al., 2018; Swift et al., 2012). Therefore, myosin II is blocked in an actin-detached state (Kovács et al., 2004). Blebbistatin is suspected not to affect the cardiac AP, [Ca2+]i transients (Efimov et al., 2004) or cardiac electrical activity, including ECG parameters, atrial and ventricular activation patterns, and refractory periods of rat and rabbit hearts (Fedorov et al., 2007; Lou et al., 2012). The effects of blebbistatin, however, on the electrical properties of pacemaking cells remain unknown.   4.3 Materials and methods  Animals I used 31 adult (12-18 months post-fertilization) AB strain zebrafish, obtained either through in-house breeding at Dalhousie University (n=15), bred from a line obtained from the zebrafish international resource center, Eugene, OR, USA), or from a local pet store (Noah’s arc, Vancouver) for use at The University of British Columbia (n=16). Fish were kept under standard laboratory conditions; held in recirculating aquaria at 28°C, 14:10 light:dark photoperiod and fed daily.    Heart isolation and tissue preparation All experiments used a reduced cardiac preparation, consisting of the duct of Cuvier, the SAN, and the atrium. This reduced cardiac preparation offered the following benefits i) isolation from neuronal and hormonal input to modulate the intrinsic firing rate of pacemaker cells (in the absence of spontaneous neuronal activation) and their response to temperature, ii) easy access to the pacemaker cells, and iii) preservation of cell-to-cell connectivity which would otherwise be lost in isolated pacemaker cells.   132 Zebrafish were sacrificed by lethal overdose of buffered MS-222, followed by severing of the spine and pithing of the brain. Fish were transferred to a dissection dish at room temperature (RT) and submerged in extracellular fluid (ECF) solution containing (in mM): 124.1 NaCl, 5.1 KCl, 2.9 Na2HPO4, 1.9 MgSO4-7H2O, 1.4 CaCl2-2H2O, 11.9 NaHCO3; pH 7.2 (Stoyek et al., 2015; Stoyek et al., 2017). The heart was exposed through a ventral midline incision and the ventricle, atrium, sinus venosus, and ducts of Cuvier were removed and transferred to a recording chamber containing RT ECF solution. The bulbus arteriosus, ventricle, and the majority of the atrium are removed from the preparation, leaving only the SAN, part of the atrium, and the duct of Cuvier (Figure 4.1). One side of the atrial tissue and the sinus venosus was pinned to the bottom of the recording chamber and the remaining free tissue was folded back over the tissue to expose the interior of the SAN, providing access to the interior of the cardiac tissue and was pinned to the bottom of the Sylgard-coated recording chamber using dissection pins.   Figure 4.1. Schematic diagram of the sagittal view of the zebrafish heart dissection. Insert shows the reduced heart preparation retained for intracellular microelectrode recordings. The approximate location of the pacemaker cells in the sinoatrial node is indicated in red.    133 Forceful cardiac contractions of regular rhythm resumed during a 5 min recovery after this procedure. Control APs were then recorded from the pacemaker cells by impaling a microelectrode into the cell. The bath solution was then replaced with fresh saline containing 10 μM blebbistatin. The stock solution of blebbistatin (13013; Cayman chemicals, Burlington, ON, Canada) was made in DMSO to 10 mM and 5 μL aliquots were wrapped in foil and stored at -20°C until used. The preparation was left undisturbed in the stop bath (i.e. no perfusion of ECF solution) until complete cession of contraction. Microelectrodes could then be impaled into the pacemaker cells without causing an immediate depolarization of the baseline linked to the rupture of the cell membrane and subsequent ion leak. APs could easily be recorded continuously for over 1 min without signal decay and on one occasion for 20 min. Throughout, the saline containing blebbistatin was protected from direct light to avoid photoinactivation and, whenever possible, kept in the dark by covering the tissue with an aluminum dome.    Recording of the action potentials Microelectrodes, with a mean resistance of 35 MΩ when filled with 3 M KCl, were used to impale SAN cells and record the electrical activity using an Axopatch 200B amplifier and a Digidata 1320 digitizer (Axon Instruments; San Jose, CA, USA). The microelectrodes were pulled from borosilicate glass capillaries (Sutter Instruments; BF-150-110-7.5, Novato, CA, USA) using a micropipette puller (Sutter Instruments; P-97) and were back-filled with the KCl solution using a MicroFil micropipette filling needle (WPI, Sarasota, FL, USA). A 0.25 mm chloride-coated silver electrode was inserted into the microelectrode and was submerged in the pipette solution and provided continuity of electrical conductivity to the headstage and amplifier. Micromanipulation of the microelectrode was performed using a Sutter micromanipulator (MPC- 134 200; Sutter Instrument Novato, CA, USA). A silver-chloride pellet was immersed in the bath solution and served as the reference electrode. Pacemaker APs were identified by a slow diastolic depolarization at phase 4 of the AP. Once a signal was obtained, the pipette was left undisturbed for the duration of the recording. APs were deemed stable when the maximum hyperpolarization voltage and AP threshold potential were consistent over consecutive beats without drifting towards 0 mV, and the amplitude of the APs did not differ. Stable APs were recorded for a minimum of 20 s before the micropipette was removed from the cell.  APs from pacemaker cells were recorded at room temperature (20°C-23°C). For the UBC group, APs were then recorded over a temperature range (20°C to 33°C) by progressively increasing the temperature of the stop bath in discrete temperature steps (by 1°C every 5 min) using an in-line heater attached to a water jacket (Warner Instruments, Hamden, CT, USA). APs were periodically recorded from different SAN cells during warming, noting the exact temperature when APs were successfully recorded from a pacemaker cell.     Data analysis APs were analyzed offline using Clampfit (Axon instruments). AP quality was first verified; APs with a spontaneous firing rate lower than 50 beats min-1 or that had a depolarizing baseline over time were excluded from the analysis. Six consecutive AP were analyzed per recording and 16 parameters were extracted from each AP (see Figure 4.2). Data recorded at Dalhousie University and the University of British Columbia were analyzed as two separate data sets, referred to as DU and UBC, respectively. Data for the temperature bins were pooled from at least 3 fish and a minimum of 6 APs were used for the data extraction from recordings of each fish  135 (Table 4.1). In all cases, with the exception of heart rate (fH; beats min-1), “rate” refers to the change of voltage per unit of time (mV ms-1).  136 Table 4.1. Table of the number of fish used and action potentials recorded for each treatment and temperature grouping.   Control  Blebbistatin   20°C–22.9°C 23°C–25.9°C 26°C–28.9°C 29°C–33°C  20°C–22.9°C 23°C–25.9°C 26°C–28.9°C 29°C–33°C DU  Fish 8 - - -  7 - - -  AP 22 - - -  22 - - - UBC  Fish 5 5 4 3  5 6 2 3  AP 12 9 6 9  12 9 7 6  137  Figure 4.2. A schematic protocol of action potential parameter extraction from the raw traces recorded from pacemaker cells. The points used for data extraction are indicated in green and the parameters extracted are numbered in black from 1 to 16. The action potential parameters are listed under the action potential and the points used for data extraction are in green in parentheses.         138  Statistical analysis The effect of blebbistatin on the AP parameters at RT (20°C to 22.9°C) was determined by comparing non-treated and blebbistatin-treated reduced heart preparations for both group DU and group UBC. Data were tested for normality using the Kolmogorov-Smirnov test and statistical significance was determined using either a t-test or a Mann-Whitney U test. To determine the effect of blebbistatin on the AP parameters at different temperatures, a comparison of temperature bins was implemented by performing a repeated measures ANCOVA followed by a Bonferroni multiple comparisons post hoc test using SPSS version 27. Data bins were tested for normality using the Kolmogorov-Smirnov and Anderson-Darling methods. The relatively large temperature bins (~3°C) increase the risk of type 2 statistical error; therefore a linear regression analysis was also performed to evaluate the effect of blebbistatin across the whole thermal range of the warming profile. The linear regression analyses were performed using the least-squares method and by using an analysis of covariance to test whether the slopes and intercepts of the untreated and the blebbistatin treated groups were significantly different. Statistical difference in the mean values for each temperature grouping was then determined using a one-way-ANOVA with a Bonferroni multiple comparison test. To determine the overall effects of blebbistatin and temperature on the AP, principal components analyses were performed using JMP 15 software (Cary, NC, USA) using the restricted maximum likelihood estimate to compare APs recorded at room temperature and APs recorded between 27°C and 33°C. All statistical analyses unless otherwise stated were carried out using GraphPad prism 8. P ≤ 0.05 was the level of statistical significance for all statistical analyses. Power analysis was performed using G*Power version 3.1. (Erdfelder et al., 2009).   139 4.4 Results The application of blebbistatin resulted in a progressive reduction, followed by complete cessation, of cardiomyocyte contraction after 5 to 20 min, after which AP recordings were more easily obtained, substantially more stable, and could be recorded reliably and for longer periods of time than without blebbistatin.  Most AP parameters (10 of 16) under the untreated control conditions, including mean fH (DU, 71 ± 4 beats min-1; UBC, 71 ± 4 beats min-1), were not significantly different between the DU and UBC groups. However, six descriptors differed significantly (Table 4.3). The AP amplitude, overshoot potential and the AP depolarization potential were all significantly higher in the DU group than in the UBC group, despite a similar maximum hyperpolarization potential and the AP threshold potential (Table 4.3). Further, the DU group had a greater depolarization potential and a shorter AP depolarization duration. Consequently, the group DU and group UBC pacemaker cells were considered two distinct populations and the effect of blebbistatin on each of these populations were analyzed separately.    The effect of blebbistatin on the pacemaker action potential characteristics from the SAN region of zebrafish Consecutive pulses of APs, recorded within a short duration of one another (∼20 s), showed very little variability in the AP waveform because two control APs could be easily overlaid, as could those recorded in the presence of 10 μM blebbistatin (Figure 4.3).  Moreover, none of the 16 APs features differed between the control and blebbistatin-treated trials for either the DU group (Table 4.2) or the UBC group (Table 4.3). For example, the beat-to-beat period was similar (control vs blebbistatin: 897 ± 44 vs 879 ± 52 for DU (Figure 4.4) and 973  140 ± 53 vs 837 ± 27 for UBC). Importantly, the maximum hyperpolarization potential and the AP threshold potential were unchanged by blebbistatin (Table 4.2 and 4.3), indicating no change to the activation potential of voltage-gated Ca2+ channels of the pacemaker cells and no change to channels involved in generating the pacemaking AP with blebbistatin. Only the overshoot potential approached statistical significance (P = 0.055) and only for the DU group (Figure 4.5; Table 4.3). Additionally, the duration of 50% and 80% repolarization did not differ significantly between control and blebbistatin-treated heart preparations in the DU group at RT (Appendix 4.1; Table 4.2), nor did the overshoot potential, AP amplitude, threshold potential and the maximum hyperpolarization potential (Appendix 4.2; Table 4.2).  Blebbistatin had no overall effect on the AP voltage parameters of APs recorded at RT in the DU group as no separation of control and blebbistatin-treated cells was observed in the PCA of voltage parameters (Appendix 4.6). Further, blebbistatin had no effect on the AP duration parameters as no separation of control and blebbistatin-treated cells was observed in the PCA of AP duration parameters (Appendix 4.7).           141   Figure 4.3. Overlay of representative action potentials recorded by intracellular microelectrode in reduced zebrafish heart preparations under control conditions and with 10 μM blebbistatin.  The control action potentials were recorded from the same pacemaker cell within 20 s. The blebbistatin action potential was recorded from the same heart as the control action potentials.   Figure 4.4. The effect of blebbistatin on (A) heart rate and (B) action potential period of control and blebbistatin treated cells. Data points represent individuals. Data are presented with the SD. Statistical significance was determined using a repeated measures ANCOVA followed by a Bonferroni multiple comparisons post hoc test with P ≤ 0.05 as the level of significance.  142  Figure 4.5.  The effect of blebbistatin on action potential parameters recorded at room temperature from zebrafish heart preparations. (A) action potential repolarization duration (B) the rate of repolarization (C) diastolic depolarization duration (D) diastolic amplitude (E) rate of diastolic depolarization (F) action potential depolarization voltage (G) action potential depolarization duration (H) rate of diastolic depolarization. Data points represent individuals. Data are presented with the SD. Statistical significance was determined using a repeated measures ANCOVA followed by a Bonferroni multiple comparisons post hoc test with P ≤ 0.05 as the level of significance.  143 Table 4.2. Table of action potential parameters with and without blebbistatin in the DU group recorded at room temperature (20°C-23°C). Depolarization (depol), hyperpolarization (hyperpol), repolarization (repol). Statistical significance was determined using a t-test with P ≤ 0.05 as the level of significance.  Treatment fH (beats min-1) AP period (ms) AP amplitude (mV) Overshoot potential (mV) Maximum hyperpol potential (mV) AP threshold potential (mV) AP depol potential (mV) AP depol time (ms) Rate of depol (mV/ms) AP repol time (ms) Rate of repol (mV/ms) AP duration 50% (ms) AP duration 80% (ms) Diastolic depol amplitude (mV) Diastolic depol duration (ms) Rate of diastolic depol (mV/ms) Control 71 897 68 16 -52 -42 58 24 2.74 166 0.42 81 98 9.1 694 0.01 SEM 4 44 3 1 2 2 3 2 0.23 6 0.02 4 4 0.70 50 0.0007 Blebbistatin 65 973 70 20 -50 -40 60 24 2.59 162 0.44 84 100 9.8 786 0.01 SEM 5 54 3 2 2 2 3 1 0.13 6 0.014 5 5 0.48 51 0.0009 P-Value 0.650 0.738 0.825 0.464 0.897 0.877 0.813 0.663 0.910 0.634 0.393 0.605 0.425 0.762 0.554 0.736    Table 4.3. A table of all action potential parameters of zebrafish pacemaker cells recorded at room temperature. Data are presented as mean ± SEM. Statistical significance between the mean values was determined using a t-test with P ≤ 0.05 as the level of significance (Table on the next page).  144 AP parameter Group DU Control Group DU Blebbistatin P-Value Group UBC   Control Group UBC Blebbistatin P-Value Group UD Control /Group UBC Control   P-Value Sample size 22 22  12 12   Heart rate (beats min-1) 71 ± 4 65 ± 5 0.391 71 ± 4 76 ± 6 0.499 0.978 Period (ms) 897 ± 44 973 ± 54 0.276 879 ± 52 837 ± 27 0.599 0.795 AP amplitude (mV) 68 ± 3 70 ± 3 0.598 56 ± 2 51 ± 2 0.211 0.0007 Overshoot potential (mV) 16 ± 1 20 ± 2 0.055 8 ± 2 7 ± 1 0.321 0.0005 Maximum hyperpolarization potential (mV) -52 ± 2 -50 ± 2 0.592 -46 ± 1 -46 ± 1 0.950 0.069 AP threshold potential (mV) -42 ± 2 -40 ± 2 0.498 -36 ± 1 -36 ± 2 0.792 0.129 AP depolarization potential (mV) 58 ± 3 60 ± 3 0.699 44 ± 2 46 ± 3 0.511 0.002 AP depolarization duration (ms) 24 ± 2 24 ± 1 0.846 49 ± 2 44 ± 3 0.219 <0.0001 Rate of depolarization (mV/ms) 2.74 ± 0.23 2.59 ± 0.13 0.593 0.92 ± 0.06 2.53 ± 1.50 0.274 <0.0001 AP repolarization duration (ms) 166 ± 6 162 ± 6 0.592 169 ± 7 162 ± 10 0.565 0.758 Rate of repolarization (mV/ms) 0.42 ± 0.02 0.44 ± 0.01 0.496 0.84 ± 0.50 0.58 ± 0.29 0.391 0.003 AP duration 50 (ms) 81 ± 4 84 ± 5 0.642 77 ± 6 69 ± 3 0.244 0.556 AP duration 80 (ms) 98 ± 4 100 ± 5 0.806 97 ± 6 88 ± 4 0.257 0.814 Diastolic depolarization amplitude (mV) 9.1± 0.70 9.8 ± 0.48 0.484 6.58 ± 1.39 6.71 ± 0.76 0.478 0.086 Diastolic depolarization duration (ms) 694 ± 50 786 ± 51 0.214 661 ± 54 632 ± 53 0.706 0.447 Rate of diastolic depolarization (mV/ms) 0.01 ± 0.0007 0.0129 ± 0.0009 0.834 0.0097 ± 0.0020 0.0650 ± 0.0545 0.149 0.059  145  The effect of temperature on the pacemaker action potential with and without blebbistatin in the UBC group The effect of temperature on the AP parameters recorded is summarized in Table 4.4. Increasing temperature increased fH of the control heart preparations and shortened the AP duration as expected (Haverinen and Vornanen, 2007; Vornanen et al., 2014). Further, the duration between APs and the period of spontaneous diastolic depolarization, also decreased significantly with temperature (Figure 4.6; Table 4.4). Blebbistatin had no significant effect on the pacemaker rate as a function of temperature. For example, similar to control heart preparations and with a complete overlap of their 95% confidence intervals (CI), linear regression analysis showed that fH increased progressively with increasing temperature and beat-to-beat period decreased progressively for blebbistatin-treated pacemaker cells (Fig 4.7. A, C). Similarly, discrete temperature bins for control and blebbistatin-treated pacemaker cells did not differ significantly (Fig 4.7. A, C).   Figure 4.6. Representative action potentials recorded by intracellular microelectrode in a reduced zebrafish heart preparation from different SAN cells in the presence of  10 μM blebbistatin.    146  Figure 4.7. The effect of blebbistatin and temperature on the heart rate and beat-to-beat period. (A) A linear regression analysis of heart rate with increasing temperature of control and blebbistatin treated cells. (B) Pairwise comparison of heart rate of control and blebbistatin treated cells at different temperature intervals. (C) A linear regression analysis of beat-to-beat period with increasing temperature of control and blebbistatin treated cells. (D) Pairwise comparison of the beat-to-beat period of control and blebbistatin treated cells at different temperature intervals. Data points represent individuals. Data are presented with the SD. Statistical significance of pairwise comparisons was determined using a repeated measures ANCOVA followed by a Bonferroni multiple comparisons post hoc test with P ≤ 0.05 as the level of significance.     147 Blebbistatin did not significantly affect the AP parameters associated with spontaneous depolarization. For example, the amplitude and the rate of spontaneous diastolic depolarization did not significantly differ with the addition of blebbistatin as the 95% CI for the control and blebbistatin-treated lines completely overlapped (Figure 4.8. A, B; Table 4.4). There was no effect of temperature on the effect of blebbistatin at any of the temperature comparison bins for any of the spontaneous depolarization AP parameters with the single exception of the rate of spontaneous diastolic depolarization between 26°C and 28.9°C (P = 0.027) (Figure 4.8. A-D; Table 4.4; Table 4.5).      148  Figure 4.8. The effect of blebbistatin and temperature on the diastolic depolarization duration, diastolic amplitude, and the rate of diastolic depolarization. (A) A linear regression analysis of diastolic depolarization duration with increasing temperature of control and blebbistatin treated cells. (B) Pairwise comparison of diastolic depolarization duration of control and blebbistatin treated cells at different temperature intervals. (C) A linear regression analysis of diastolic amplitude with increasing temperature of control and blebbistatin treated cells. (D) Pairwise comparison of the diastolic amplitude of control and blebbistatin treated cells at different temperature intervals. (E) A linear regression analysis of the rate of diastolic depolarization with increasing temperature of control and blebbistatin treated cells. (F) A pairwise comparison of the  149 rate of diastolic depolarization of control and blebbistatin treated cells at different temperature intervals. Data points represent individuals. Data are presented with the SD. Statistical significance of pairwise comparisons was determined using a repeated measures ANCOVA followed by a Bonferroni multiple comparisons post hoc test with P ≤ 0.05 as the level of significance.                      150 Blebbistatin did not significantly affect either the AP depolarization voltage, which did not vary with temperature (Figure 4.9. A, B) or the AP depolarization duration and the rate of AP depolarization, which both decreased with temperature (Figure 4.9. C, E; Figure 4.9. D, F). For example, the AP depolarization was not significantly different between control and blebbistatin-treated heart preparations as the 95% CI for the fitted linear regressions completely overlapped (Figure 4.9. A). The temperature bin comparisons of the AP depolarization voltage revealed no significant differences between control and blebbistatin-treated cells at any of the comparison temperatures (Figure 4.9. B; Table 4.5). The same analysis for the AP depolarization duration and the rate of AP depolarization also revealed that the control and blebbistatin-treated heart preparations did not differ significantly (Figure 4.9. C, E; Figure 4.9. D, F; Table 4.5).  Blebbistatin did not significantly affect either of the AP repolarization parameters (the rate of AP repolarization and the action potential repolarization duration), which were all dependent of temperature (Figure 4.10). Similarly, blebbistatin did not significantly affect the AP threshold potential, the overshoot potential, the AP amplitude or the maximum hyperpolarization potential, which were independent of temperature (Appendix 4.3; Appendix 4.4; Table 4.5). Also, AP duration 50% and 80%, which both decreased with temperature, were not significantly affected by blebbistatin using linear regression (Appendix 4.5. A, C). However, the pairwise comparison between the control and blebbistatin-treated pacemaker cells revealed a significantly faster 50% and 80% repolarization with blebbistatin between 26°C and 29°C but at no other temperatures (Appendix 4.5. B, D; Table 4.5).    151  Figure 4.9. The effect of blebbistatin and temperature on the action potential depolarization voltage, action potential depolarization duration, and the action potential duration. (A) A linear regression analysis of action potential depolarization voltage with increasing temperature of control and blebbistatin treated cells. (B) Pairwise comparison of action potential depolarization voltage of control and blebbistatin treated cells at different temperature intervals. (C) A linear regression analysis of depolarization duration with increasing temperature of control and blebbistatin treated cells. (D) Pairwise comparison of depolarization duration of control and  152 blebbistatin treated cells at different temperature intervals. (E) A linear regression analysis of the duration of depolarization with increasing temperature of control and blebbistatin treated cells. (F) A pairwise comparison of the duration of depolarization of control and blebbistatin treated cells at different temperature intervals. Data points represent individuals. Data are presented with the SD. Statistical significance of pairwise comparisons was determined using a repeated measures ANCOVA followed by a Bonferroni multiple comparisons post hoc test with P ≤ 0.05 as the level of significance.                  153   Figure 4.10. The effect of blebbistatin and temperature on the rate of repolarization and action potential repolarization duration, (A) A linear regression analysis of the rate of repolarization with increasing temperature of control and blebbistatin treated cells. (B) Pairwise comparison of AP the rate of repolarization of control and blebbistatin treated cells at different temperature intervals. Data points represent individuals. Data are presented with the SD. Statistical significance of pairwise comparisons was determined using a repeated measures ANCOVA followed by a Bonferroni multiple comparisons post hoc test with P ≤ 0.05 as the level of significance.      154 Table 4.4. Summary table of the effects of temperature on each of the AP parameters at different temperatures in group UBC. Blebbistatin-treated cells at different temperatures are compared to blebbistatin-treated cells at room temperature (20°C to 22.9°C). Depolarization (depol), hyperpolarization (hyperpol), repolarization (repol). Statistical significance was determined using a one-way ANOVA with a Bonferroni multiple comparison test, using P ≤ 0.05 as the level of significance (Table on the next page).  Table 4.5. Pairwise comparison of control and blebbistatin-treated cells with increasing temperature in group UBC. Depolarization (depol), hyperpolarization (hyperpol), repolarization (repol). Statistical significance was determined using a t-test with P ≤ 0.05 as the level of significance (Table on the following page). 155 Table 4.4. Temp  °C Treatment fH (beats min-1) AP Period (ms) AP amplitude (mV) Overshoot potential (mV) Maximum hyperpol potential (mV) AP threshold potential (mV) AP depol potential (mV) AP depol time (ms) Rate of depol (mV/ms) AP repol time (ms) Rate of repol (mV/ms) AP duration 50% (ms) AP duration 80% (ms) Diastolic depol amplitude (mV) Diastolic depol duration (ms) Rate of diastolic depol (mV/ms) 20°C - 22.9°C Blebbistatin 75 860 55 9 -46 -38 47 45 1.08 165 0.34 69 88 8 611 0.02 SEM 5 51 2 1 1 2 2 2 0.07 8 0.02 3 4 1 46 0.004 23°C -25.9°C Blebbistatin 96 562 52 5 -45 -35 40 35 1.19 116 0.44 51 63 11 513 0.02 SEM 12 56 4 1 4 3 4 4 0.07 10 0.03 5 4 2 60 0.004 P-Value 0.404 0.016 >0.999 0.042 >0.999 >0.999 0.261 0.019 >0.999 <0.001 0.053 0.112 0.008 0.308 0.465 >0.999 26°C - 28.9°C Blebbistatin 136 464 45 4 -47 -41 34 29 1.47 96 0.50 36 49 5 348 0.02 SEM 17 58 4 1 5 3 3 2 0.21 5 0.04 9 8 1 58 0.003 P-Value <0.001 <0.001 0.114 0.010 >0.999 >0.999 >0.999 <0.001 0.063 <0.001 0.002 0.003 <0.001 0.424 0.003 >0.999 29°C – 33°C Blebbistatin 136 404 49 6 -42 -37 45 21 1.85 98 0.47 43 54 6 318 0.02 SEM 10 31 3 1 3 5 3 1 0.13 7 0.04 12 11 1 22 0.003 P-Value 0.001 <0.001 0.785 0.203 >0.999 >0.999 >0.999 <0.001 <0.001 <0.001 0.016 0.026 0.002 >0.999 0.002 >0.999    156 Table 4.5. Temp  °C Treatment fH (beats min-1) AP Period (ms) AP amplitude (mV) Overshoot potential (mV) Maximum hyperpol potential (mV) AP threshold potential (mV) AP depol potential (mV) AP depol time (ms) Rate of depol (mV/ms) AP repol time (ms) Rate of repol (mV/ms) AP duration 50% (ms) AP duration 80% (ms) Diastolic depol amplitude (mV) Diastolic depol duration (ms) Rate of diastolic depol (mV/ms) 20°C - 22.9°C Control 73 860 50 8 -43 -36 42 46 0.94 162 0.32 76 94 7 651 0.01 SEM 4 51 3 1 3 1 3 3 0.06 10 0.02 5 6 1 50 0.001 Blebbistatin 75 860 55 9 -46 -38 47 45 1.08 165 0.34 69 88 8 611 0.02 SEM 5 51 2 1 1 2 2 2 0.07 8 0.02 3 4 1 46 0.004 P-Value 0.773 0.572 0.402 0.573 0.584 0.573 0.298 0.966 0.150 0.656 0.614 0.370 0.500 0.782 0.491 0.055 23°C -25.9°C Control 84 741 48 5 -44 -32 39 36 1.19 143 0.38 66 80 11 584 0.01 SEM 6 43 3 1 4 3 4 5 0.13 19 0.04 9 10 2 35 0.003 Blebbistatin 96 562 52 5 -45 -35 40 35 1.19 116 0.44 51 63 11 513 0.02 SEM 12 56 4 1 4 3 4 4 0.07 10 0.03 5 4 2 60 0.004 P-Value 0.400 0.312 0.828 0.826 0.831 0.826 0.910 0.735 0.963 0.216 0.249 0.153 0.121 0.987 0.481 0.172 26°C - 28.9°C Control 142 464 48 6 -38 -31 34 21 1.86 102 0.42 72 80 10 345 0.03 SEM 22 58 7 2 3 3 3 3 0.24 6 0.04 6 6 2 53 0.002 Blebbistatin 136 464 45 4 -47 -41 34 29 1.47 96 0.50 36 49 5 348 0.02 SEM 17 58 4 1 5 3 3 2 0.21 5 0.04 9 8 1 58 0.003 P-Value 0.819 0.812 0.255 0.410 0.143 0.410 0.063 0.087 0.244 0.284 0.186 0.008 0.010 0.086 0.964 0.027 29°C – 33°C Control 165 404 50 8 -46 -37 42 27 1.58 73 0.55 47 52 6 318 0.02 SEM 12 31 4 1 4 5 4 2 0.14 5 0.03 4 4 1 32 0.006 Blebbistatin 136 404 49 6 -42 -37 45 21 1.85 98 0.47 43 54 6 318 0.02 SEM 10 31 3 1 3 5 3 1 0.13 7 0.04 12 11 1 22 0.003 P-Value 0.107 0.417 0.289 0.333 0.530 0.333 0.519 0.384 0.212 0.540 0.152 0.670 0.812 0.742 0.997 0.445  157 4.4.3. The effect of blebbistatin on action potential voltage and duration  Given the similarity of APs from control and blebbistatin-treated pacemaker cells, I decided to do a global analysis of the two groups using PCA analysis to visually reveal interactions between temperature and blebbistatin by comparing the two temperature extremes, RT and 27°C-33°C. Increased temperature clearly had an effect on the time-dependent AP parameters as there was a clear separation between the APs recorded at RT and those recorded at high temperature (Figure 4.11). However, there was no effect of blebbistatin as the APs recorded at RT from control and blebbistatin-treated heart preparations overlapped and those recorded at high temperature also overlapped, regardless of the treatment group (Figure 4.11).   There was no effect of blebbistatin on the voltage-dependent AP parameters as recorded at RT (20°C-23°C) in the UBC group as no separation of APs recorded from untreated cells was observed in the PCA (Figure 4.12). However, the effect of blebbistatin on the voltage parameters of the AP recorded at high temperature (27°C-33°C) revealed a partial separation of the control and blebbistatin-treated cells. The control APs cluster close to the APs recorded at RT whereas the blebbistatin-treated cells tend to cluster away from the untreated preparations. However, the overlap between the clusters and the non-significant differences of all of the individual AP parameters suggest that there is no overall significant effect of blebbistatin at high temperature (Figure 4.12). Although none of the AP parameters differed significantly between control and blebbistatin-treated groups, the summation of any small changes in the AP parameters may drive a significant effect of blebbistatin on the overall AP waveform.   158   Figure 4.11.  Principal component analysis of time-dependent action potential parameters of control and blebbistatin-treated heart preparations recorded at room temperature (20°C-23°C; untreated, solid blue circle; blebbistatin treated, green diamond) and control and blebbistatin-treated cells action potentials recorded at high temperature (27°C-33°C; untreated, solid purple circle; blebbistatin treated, red diamond). APs recorded at room temperature are circled in the solid blue and the AP recorded at high temperature are circled in the dashed purple. Each point represents APs recorded from a single pacemaker cell.    159  Figure 4.12. Principal component analysis of voltage-dependent action potential parameters of control and blebbistatin-treated heart preparations recorded at room temperature (20°C-23°C; untreated, blue circle; blebbistatin treated, green diamond) and control and blebbistatin-treated cells action potentials recorded at high temperature (27°C-33°C; untreated, purple circle; blebbistatin treated, red diamond). APs recorded at room temperature from blebbistatin treated heart preparations are circled in dashed purple and APs recorded at high temperature from blebbistatin-treated heart preparations are circled in solid blue. Each point represents APs recorded from a single pacemaker cell.    160 4.5 Discussion My study investigated the electrophysiologic effects of blebbistatin, a myosin II uncoupler that is increasingly used in cardiac research of active patterns of AP morphology, and intracellular Ca2+ signalling of mammalian hearts for cardiomyopathy investigation. By uncoupling excitation-contraction with blebbistatin, the motion artifacts caused by the contraction of cardiac tissue are eliminated, allowing electrophysiological studies. Other excitation-contraction uncouplers, including 2,3-butanedione monoxime and cytochalasin D, are known to alter Ca2+ handling, ion channel kinetics and modify the characteristics of the AP in a species-dependent manner (Jou et al., 2010; Kettlewell et al., 2004; Liu et al., 1993; Rueckschloss and Isenberg, 2001; Watanabe et al., 2001). Blebbistatin has been used in optical mapping studies in multiple species including mouse (Dou et al., 2007), rat (Farman et al., 2008; Fedorov et al., 2007), rabbit (Brack et al., 2013; Fedorov et al., 2007), as well as horse (Fenton et al., 2008), dog (Kong et al., 2009), and human (Fedorov et al., 2011).  Blebbistatin has previously been shown to not significantly affect the ion channel dynamics, calcium handling and the electrophysiological parameters of the heart including ECG parameters, atrial and ventricular effective refractory periods, and atrial and ventricular activation patterns of rats and rabbits (Fedorov et al., 2007; Lou et al., 2012), and in embryonic zebrafish (Jou et al., 2010). However, there is considerable variability between species in the electrophysiological response to blebbistatin reported in the literature, including conflicting data within species (Brack et al., 2013; Fedorov et al., 2007). Further, the effects have mostly been evaluated on the ventricular and atrial cells, with very little focus on cardiac pacemaker cells. Furthermore, the effects of blebbistatin have not been investigated in any adult fish species. Hence, I investigated the effects of blebbistatin on the electrophysiological properties of zebrafish cardiac  161 pacemaker cells in two independent populations to determine the suitability of blebbistatin as an excitation-contraction uncoupler for cardiac and electrophysiology research. I found differences only between populations that were all related to the depolarization phase of the AP and thus may be due to inherent differences in Ca2+ channel expression in the two populations of zebrafish, leading to faster rise times and a larger AP or caused by slight differences in recording conditions between the two populations. Within the two zebrafish populations, I discovered that blebbistatin effectively and rapidly uncouples excitation-contraction and can be used in the zebrafish heart without concern for modifying the AP as blebbistatin did not significantly affect any of the electrophysiological properties of the cardiac pacemaker AP. However, the limited sample size in this study may not be sufficiently large to detect effectively detect non-significant change between the control group and the blebbistatin treated heart perpetrations as the power analysis revealed that a total sample size of 372 fish would be required to achieve 80% analysis power (appendix 4.8).  As many isolated heart and cellular studies use room temperature as the test temperature (Hassel et al., 2008; Hassinen et al., 2015; Zhang et al., 2011), I decided to test the effect of blebbistatin at room temperature before increasing the temperature to test the effect of temperature on the effect of blebbistatin. The temperature bin comparisons revealed no significant differences between control and blebbistatin-treated cells for all AP features measured, with the exception of the 50% and 80% AP duration and the rate of diastolic depolarization at 26°C to 28.9°C for the UBC group. These significant differences were, however, lost at the 29°C to 33°C temperature bin, and may have been caused by the lower AP sample size in the 26°C to 28.9°C temperature bin. Overall, there was no effect of blebbistatin on the response of the pacemaker AP to increasing temperature.  162 Blebbistatin has been previously tested on embryonic (48 hours post-fertilization; hpf) zebrafish hearts (Jou et al., 2010), where blebbistatin (10 μM) effectively uncoupled cardiac excitation-contraction. Regardless of the concentration of blebbistatin used (1, 5 or 10 μM) no significant effects on AP morphology or the generation of spontaneous APs were seen for atrial and ventricular cells. Cycle length, maximum diastolic potential, maximum upstroke velocity, and the AP duration were all unchanged (Jou et al., 2010). Although fH, and AP repolarization rates were faster in the 48 hpf embryos than in the adult zebrafish, and the maximum diastolic potential in the zebrafish embryo was more negative than my adult recordings (-56.2 ± 6.2 mV and -55.7 ± 6.2 mV for control and blebbistatin treated embryonic cells respectively compared to -43 ± 3 mV to -52 ± 2 mV and -46 ± 1 mV to -50 ± 2 mV for control and blebbistatin treated heart preparations respectively), blebbistatin had no effect on the AP waveform in the embryos or the adult zebrafish. Therefore, the effect of blebbistatin on the electrophysiological properties of the zebrafish heart, at both the adult and the early stages of development do not significantly alter heart electrical properties, and my results for the effect of blebbistatin on zebrafish pacemaker cells from the SAN are in line with these earlier findings.  A principal components analysis of my data showed no clear separation of the untreated and the blebbistatin-treated heart preparations, indicating that the overall effects of blebbistatin do not significantly modify the AP waveform which is in line with the individual analysis of the AP features. Therefore, blebbistatin does not significantly alter the AP waveform indicating that blebbistatin is an effective excitation-contraction uncoupling agent and can therefore be used for high-fidelity measurements of electrical activity and calcium transients in zebrafish heart.  Although this study is the first to directly record the pacemaker AP from adult zebrafish pacemaker cells from the intact SAN, this data is in line with previous findings for adult zebrafish  163 heart rate of isolated heart preparations (Stoyek et al., 2016). Therefore, the dissection of the zebrafish heart produces repeatable results for fH, and likely does not produce any changes in the electrophysiological properties of the pacemaker cell AP. Among the few differences observed in AP parameters between my two zebrafish populations, depolarization rate was faster in the DU group, possibly indicating slightly different recording conditions between the laboratories or a difference in the depolarizing Ca2+ channels between the fish populations.  In mice, blebbistatin did not significantly affect the action potential duration, ventricular activation, or conduction velocity (Baudenbacher et al., 2008). Blebbistatin did, however, reduce myosin Ca2+ sensitivity which resulted in a reduction of arrhythmia susceptibility, even in Ca2+-sensitised hearts. Therefore, blebbistatin protects mice hearts against arrhythmia by reducing myofilament sensitivity to Ca2+ (Baudenbacher et al., 2008). In another study, blebbistatin was reported not to have any effect on the electrophysiological properties of the Ca2+ transient or the AP (i.e. amplitude, duration, upstroke velocity, and time of decay) in the rat heart (Fedorov et al., 2007). Furthermore, blebbistatin has been found not to modify Ca2+ handling of isolated rat myocytes (Farman et al., 2008). Finally, blebbistatin has been used to immobilize human hearts, with no reported effect of blebbistatin on the AP (Fedorov et al., 2010; Fedorov et al., 2011; Glukhov et al., 2010). Although some studies have reported that blebbistatin has no significant effect on the electrophysiological properties of the heart, even at the highest blebbistatin concentration used (10μM) (Fedorov et al., 2007; Lou et al., 2012), significant effects of blebbistatin on the electrophysiological properties of the New Zealand white rabbits ventricle have been reported (Brack et al., 2013). Blebbistatin significantly prolonged the left ventricular apical and basal monophasic action potential duration in Langendorff-perfused hearts and increased the maximal  164 slope of restitution whilst significantly reducing the heart’s susceptibility to ventricular fibrillation at a blebbistatin concentration of 5μM (Brack et al., 2013). The effect of blebbistatin remains under debate for this species with a more recent study confirming the prolongation of the AP duration with blebbistatin contraction-inhibited hearts (Kappadan et al., 2020). There was also reduced ventricular fibrillation as observed in the study conducted by Brack et al. The AP duration was also prolonged in Langendorff-perfused pig hearts containing 10 μM of blebbistatin (Lee et al., 2019). However, this study lacks an appropriate time paired control and thus remains an observation. A caveat of all of these studies, and any study comparing contracting tissue with uncoupled excitation-contraction, is the difference in metabolic demand of these tissues that may drive shortening of the AP due to elevated ATP concentration, activating ATP-sensitive potassium channels (Garrott et al., 2017; Lee et al., 2019). Further, in rats, blebbistatin has been reported to disrupt intracellular calcium dynamics as spontaneous excitation and triggered activities were observed with the application of blebbistatin (Kanlop and Sakai, 2010). However, the concentrations used were between 10 and 100 μM, significantly higher than any other study, and may be the origin of the blebbistatin-induced spontaneous excitation and triggered activities. Furthermore, the authors reported no significant difference in the electrophysiological properties of the rat heart when using either 2,3-butanedione monoxime or cytochalasin D, two compounds widely known for inducing shifts in electrophysiological properties of multiple species (Kettlewell et al., 2004; Liu et al., 1993; Rueckschloss and Isenberg, 2001; Watanabe et al., 2001). Therefore, the effects of blebbistatin may be highly species-dependent and should therefore be thoroughly investigated in different species of interest for cardiovascular research. One important application of blebbistatin is the immobilization of cardiac tissue for optical mapping and fluorescent imaging of Ca2+ transients. It has been reported that blebbistatin can  165 increase the intracellular resting fluorescence of the commonly used non-ratiometric fluorescent dyes such as Fluo-3, Fluo-4, or Fluo 5F by up to 39% (Fedorov et al., 2007; Swift et al., 2012), and is a major caveat of blebbistatin. It remains unclear if the increase in fluorescence is due to fluorescence of the blebbistatin molecule or if it is caused by a blebbistatin-mediated increase in [Ca2+]i. Increased [Ca2+ ]i in the presence of blebbistatin may derive from modified ion channel function and requires further investigation.  4.6 Limitations and perspectives  In this two-center study, blebbistatin was shown to effectively inhibit cardiac contractions and does not significantly alter AP parameters regardless of temperature. The two-center study design increases the robustness of the study as the same effect of blebbistatin was found in both datasets. However, due to the genetic heterogeneity of zebrafish, the study design also presents a limitation as the two data sets cannot be directly compared. To overcome this, zebrafish from each study center should have been sent to the other study center and the experiments performed on both populations in each study center.  Inhibition of excitation-contraction of the cardiac tissue enables reliable intracellular recording of pacemaker electrical activity and enables the use of fluorescent-based assays that are sensitive to motion artifacts. Direct intracellular recordings of pacemaker activity coupled with optical mapping of the atrium and the ventricle will enable future research on pacemaking and electrical conduction. In line with my chapter objective, blebbistatin provides a tool that can be used to evaluate the role of the membrane and calcium clocks at the level of the pacemaker cell, thus eliminating the off-target effects of the drugs on the working myocardium. Furthermore, direct recording of pacemaker cells will enable future drug screening research for pacemaker research.  166 One limitation of the study is that it requires an ex vivo preparation which deprives the heart of autonomic control and hormonal signalling. Further, in order to gain access to the pacemaker cells, the heart perpetration is further reduced to gain access to the endocardium. Disruption of the pacemaker ring and the surrounding tissue during removal of the heart and perpetration of the reduced heart tissue preparation may affect the pacemaker cells by severing their interconnectivity. Another limitation of this study is the absence of time-dependent run down of the sample and a temperature-independent control for run down. To effectively estimate the effect of run-down, both a temperature-independent (no change in temperature) and a temperature-dependent control, where the temperature is lowered instead of increased would be required.  Blebbistatin can be inactivated by brief exposure to 488 nm light (Sakamoto et al., 2005), while blue light irradiation may generate free radicals (Fedorov et al., 2007). Recently, para-aminoblebbistatin has been developed which is highly soluble, is non-phototoxic, and is non-fluorescent (Várkuti et al., 2016). However, despite research showing no cardiac cytotoxic effect of para-aminoblebbistatin in zebrafish embryos  (Várkuti et al., 2016), the electrophysiological effects of para-aminoblebbistatin are unknown in the adult zebrafish and should be determined before its widespread use in cardiac electrophysiology research. As such, this thesis chapter may provide a benchmark for such an investigation.  4.7 Conclusion I have demonstrated in this chapter that blebbistatin has no significant effect on any of the AP parameters measured, regardless of the temperature, and has been previously demonstrated in embryonic zebrafish can be used to uncouple excitation-contraction of the adult zebrafish heart. My findings have implications for electrophysiological studies, such as optical mapping of the zebrafish heart, that can now use blebbistatin to uncouple excitation-contraction and reliably  167 record the electrical activity of pacemaker cells without interfering signals of mechanical contraction. The method outlined could provide a platform for future investigation into mechanisms of cardiac pacemaking and thermal tolerance of pacemaking mechanisms at the level of the pacemaker cell, and will provide direct evidence of the effect of temperature and of pharmacological inhibition, or knockout of Hcn channels on pacemaking.    168  General discussion and future directions 5.1 Synopsis The objective of my thesis was to provide functional insight into the mechanism of cardiac pacemaking in the zebrafish. My specific aims were to determine i) the mechanisms that generate the pacemaking current and their respective contribution to determining heart rate (fH) and to determine the thermal tolerance of the major pacemaking mechanisms and, ii) determine the role of Hcn4 in cardiac pacemaking. My reasoning for such aims stems from a century old, yet enigmatic question – that of the origin of the cardiac pacemaker’s spontaneous activity (Keith and Flack, 1907; Keith and Mackenzie, 1910) – a topic that remains the subject of intense debate in the scientific community (DiFrancesco, 2020).  The ever-increasing quantity of studies involving zebrafish, especially those that establish the zebrafish as a model of cardiac disease, made the zebrafish a highly relevant species for my thesis. The genetic and molecular mechanisms of zebrafish cardiac physiology are largely conserved with humans (and other vertebrates) and the AP waveform is almost identical to that of humans as the ion channels and major cardiac currents are quantitively similar between zebrafish and humans (Howe et al., 2013; Vornanen and Hassinen, 2016). Furthermore, the fully sequenced genome of the zebrafish and the rapid and external embryonic development make the zebrafish easily amenable to genome engineering. The zebrafish is thus the ideal model organism to study the mechanisms of cardiac pacemaking and their thermal tolerance. In an era of global change, the origin and thermal tolerance of the cardiac pacemaker is of major importance for determining the capacity of a species to respond to the evolving climate and to determine their capacity to withstand and adapt to the mitigating environmental factors of the present climate, yet the cause of  169 temperature-dependent deterioration of cardiac electrical excitation remains elusive and our understanding of pacemaking in fish is incomplete (Vornanen, 2016; Vornanen, 2020).   In my first two data chapters, I identified two major pacemaking mechanisms in the zebrafish heart (chapter 2) and addressed the specific role of Hcn4 in cardiac pacemaking of the zebrafish heart using a CRISPR knockout model (chapter 3). Pharmacological blockade of either the Hcn channels or RyR and SERCA pumps independently revealed that the membrane and calcium clocks generate the depolarizing current of pacemaker cells by determining the percentage decrease in maximum heart rate (fHmax) induced by inhibiting the pacemaking mechanisms in both of these chapters. I further demonstrated that their relative contribution varies with temperature, with the membrane clock possessing a slight dominancy over the calcium clock mechanism which plays an additional role in setting pacemaker activity that was independent of temperature, whereas the membrane clock was more dependent on temperature. I further demonstrated that zebrafish cardiac pacemaking can be independent of Hcn4 as the fHmax of Hcn4 knockout fish did not differ significantly from wild-type fish, including when the Hcn channels were inhibited with zatebradine, suggesting total compensation for the loss of Hcn4 and that cardiac function is maintained in the Hcn4 knockout fish through alternative Hcn channels. These two data chapters also addressed the role of cardiac pacemaking in the thermal tolerance of the zebrafish heart by heating the fish until signs of cardiac collapse to evaluate the thermal tolerance of the pacemaking mechanisms. Despite the advances made in chapters 2 and 3 for understanding the mechanisms of cardiac pacemaking, the evidence they provide is indirect, as neither chapter provides direct recording of the effect of temperature and pharmacological intervention on the electrical activity of pacemaker cells. Therefore, a method enabling the direct recording of pacemaker AP is needed  170 to enable the field to advance. Thus, I assessed the suitability of blebbistatin as an excitation-contraction uncoupler for electrophysiology studies of the zebrafish heart (chapter 4). I demonstrated that blebbistatin does not significantly alter the pacemaker AP, regardless of the temperature, and is thus a reliable investigative tool for cardiac electrophysiology in the zebrafish.   5.2 Thermal acclimation and resetting of the heart rate Many fish species, but not all, reset fH after thermal acclimation to improve cardiac performance at the new temperature (Drost et al., 2016; Ferreira et al., 2014; Klaiman et al., 2011; Safi et al., 2019; Vornanen, 2016). I hypothesized that the zebrafish heart would reset intrinsic fH with thermal acclimation and that fish acclimated to the warmest temperature would have a higher cardiac thermal tolerance. I further hypothesized that in the zebrafish, warm acclimation would result in a reduced fHmax at all test temperatures but a greater thermal tolerance and higher maximum fH, whereas cold acclimation would result in a higher fHmax at cold test temperatures, a reduced fHmax at high temperatures and a lower maximum fH. Although not all species reset fH, my hypothesis rests on the fact that the zebrafish’s natural habitats have a wide thermal range, ranging from 14°C to 33°C, with extreme temperatures of 6°C in the winter to 38°C in the summer (Spence et al., 2008), and daily temperature variation up to 5.6°C (López-Olmeda and Sánchez-Vázquez, 2011). However, my hypothesis of cardiac pacemaker resetting with thermal acclimation was rejected as fHmax was unchanged after thermal acclimation. Furthermore, low (18°C) and high (28°C) acclimation temperatures limited the cardiac thermal tolerance, and fish acclimated to an intermediate temperature (23°C) had the highest thermal tolerance and a wider thermal optimum range. Many generations of zebrafish domestication may have led to lineages with low acclimation capacity due to constant rearing temperature and the absence of fluctuating temperatures in laboratory acclimation conditions, which has been reported to increase thermal tolerance (Morgan  171 et al., 2019; Schaefer and Ryan, 2006). Indeed, a previous study has demonstrated that zebrafish do not reset fH after acclimation to cold (18°) and warm temperatures (28°C) (Lee et al., 2016). Whether these differences of acclimation capacity are caused by genetic differences or differences in gene expression remain unknown. Future evaluations of the acclimation capacity of the zebrafish could consider wild-caught populations and ultimately investigate the genetic basis of acclimation, which is currently unknown, but that may involve changes in gene expression of membrane and calcium clock genes or changes in isoform expression or splice variants. Also, the temperature during embryonic development can have long-lasting effects on the thermal tolerance and acclimation capacity of the zebrafish and has persistent effects on metabolic enzymes, muscle fibre recruitment and cardiac anatomy (Dimitriadi et al., 2018; Johnston et al., 2009; Schnurr et al., 2014; Scott and Johnston, 2012). Therefore, embryonic acclimation and transgenerational acclimation effects on cardiac physiology and thermal tolerance need to be investigated. The acclimation capacity of fish, even in species known to reset fH, has been recently questioned by a study demonstrating that rainbow trout do not always reset fH with acclimation (Sutcliffe et al., 2020). In addition to this, the acclimation timeframe for this species was thrown into question with the revelation that the acclimation processes can occur within the first hour of acclimation. Furthermore, the distinction between an acute response and an acclimation response becomes difficult to distinguish and may require species-specific investigation in gene expression patterns with acute temperature exposure vs. acclimation.      172 5.3 Mechanisms of cardiac pacemaking My thesis quantified the relative contribution of the two major known pacemaking mechanisms for the first time and identified their relative importance across a warming profile to determine cardiac thermal tolerance. My thesis significantly contributes to the current understanding of pacemaking dynamics in an increasingly popular vertebrate model and provides further directions for future research in the field of cardiac pacemaking in fish. Furthermore, my thesis provides proof-of-principal for a methodology for direct recording of the electrophysiological activity of pacemaker cells in the intact tissue of the SAN.   The membrane clock  As the mammalian pacemaker is believed to be largely dependent on the membrane clock and HCN channels, I hypothesized the same is true for the zebrafish heart and that pharmacological block on these channels would significantly reduce fHmax. This hypothesis was supported by a large reduction (up to 60%) in fHmax when Hcn channels were blocked by zatebradine. Thus, the pacemaker potential of the zebrafish heart is in part generated by the If current generated by Hcn channels.  Zatebradine is not a specific blocker of hyperpolarization-activated channels and is known to reduce outward potassium channels, notably, IKr (Carmen et al., 1996; Van Bogaert and Pittoors, 2003; Wilson et al., 2016) and has been shown to prolong the AP duration in guinea-pig and rabbit hearts (Doerr and Trautwein, 1990; Thollon et al., 1994). Off-target effects of zatebradine constitute a major caveat in my thesis work, as zatebradine is known to inhibit IKr, possibly resulting in a prolongation of the AP. These off-target effects modify the AP and may ultimately change fH. Ivabradine, from which zatebradine is derived, has also been shown to block IKr and  173 prologue the AP duration in fetal mice, (Lees-Miller et al., 2015) but the effect of ivabradine on the adult fish heart has not been assessed to date. However, in a preliminary study, I found that 4 μg g-1 ivabradine significantly reduced fH but could also induce arrhythmia that often resulted in asystole in the zebrafish. The cause of asystole is unclear and may have resulted from a complete block of cardiac pacemaking through non-specific inhibition of ion channels, or reduced activation potential of Hcn channels by inhibiting repolarizing K+ channel IKr (Lees-Miller et al., 2015). Due to the off-target effects of pharmacological agents that are currently available, their pacemaker-specific effects need to be investigated at the level of the pacemaker cell. Only in doing so can the respective roles of the membrane and calcium clocks be fully elucidated. Novel inhibitors with single-channel specificity will greatly improve the accuracy of my results and will definitively determine the origin of the fish pacemaker current.  The role of Hcn channels in fish cardiac pacemaking is already known to be species-dependent. For example, cardiac pacemaking in the Pacific hagfish (Eptatretus stoutii) is only driven by Hcn channels as zatebradine all but stopped cardiac contractions whereas ryanodine had no effect (Wilson and Farrell, 2013). The membrane clock in the brown trout on the other hand appears to play a reduced or no role in cardiac pacemaking (Hassinen et al., 2017). Further, the major Hcn channels expressed in the SAN of the brown trout, produced no current (Hcn3) and a 1.2 pA/pF current density (Hcn4) at -140 mV when expressed in CHO cells, inconsistent with the membrane clock-driven pacemaking model where If plays a major role. Not only do these results raise questions about the relative importance of the calcium and the membrane clock, but also about the relative importance and relative contribution of the different Hcn channels in developing If. All Hcn channel isoforms of the fish SAN may contribute towards If and may have different functional voltages ranges over which they operate to ensure the pacemaking function across a  174 wide range of cellular (e.g. pH, ion concentration) and environmental conditions (e.g. temperature). In fish, only one study has looked at the differentially expressed genes between the SAN and the atrium and found that genes of both the membrane and calcium clocks were differentially expressed and that protein expression of both clocks changed with thermal acclimation. Specifically, Hcn2 and SERCA2 were both downregulated in the SAN of warm acclimated fish (Sutcliffe et al., 2020). However, gene expression patterns between chambers and mechanisms of thermal acclimation remain largely undetermined. It is now clear however that acclimation mechanisms do not follow a simple linear change of gene expression between two temperatures, but that the process of acclimation is highly dynamic and is heavily dependent on the thermal history of the fish (Sutcliffe et al., 2020).    The calcium clock  Details of molecular and ionic mechanisms of cardiac pacemaking in fish are far less established than in mammals, therefore my hypotheses were based largely, but not entirely on mammalian literature. For example, the calcium clock plays an important secondary role compared with the membrane clock in mammalian cardiac pacemaking (Bers, 2002). Thus, I hypothesized that in the zebrafish, the calcium clock would also play a reduced secondary role in the generation of the pacemaker current. This hypothesis was rejected as the calcium clock appeared to play a major role in cardiac pacemaking although its relative contribution remained lower than the membrane clock.  Inhibition of the calcium clock resulted in a ~40% reduction in fHmax, lower than the reduction in fHmax observed with the inhibition of the membrane clock. A caveat in the pharmacological inhibition of the calcium clock resides in the significant role played by SR calcium cycling in excitation-contraction of the working myocardium. By inhibiting SR calcium  175 cycling in pacemaker cells, the calcium transient will be slowed in both its development and its decay. Inhibition of SERCA inhibits Ca2+ uptake to the SR through SERCA, thus slowing the decay of the Ca2+ transient, slowing the relaxation of the myocyte. At high temperature when fH is rapid, a slowed myocyte relaxation will likely reduce the maximum contractile force as sarcomere relaxation is not total, and may limit the maximum beating frequency, thus limiting the relevance of my results for pacemaking. In order to assess the effect of ryanodine and thapsigargin on the Ca2+ transient, single-cell Ca2+ transient recording using fluorescent probes (e.g. Fura-2) would provide insight into the transient dynamics. Further, force transduction of paced ventricular strips will provide insight into the potential reduction in contractile force and the force-frequency relationship, although challenging due to the size of the zebrafish heart. Definitive evidence for the effect of SR Ca2+ cycling inhibition on cardiac pacemaking will need to be provided by direct intracellular electrophysiological recordings of pacemaker cell activity. Consistent with the absence of fHmax resetting with temperature acclimation, no compensatory changes in SR Ca2+ cycling were observed as fHmax was reduced by ~40% regardless of the acclimation temperature. The role of the SR in cardiac resetting, however, may be important for other species such as rainbow trout that do reset intrinsic fH after both warm and cold acclimation (Sutcliffe et al., 2020). Indeed, expression of SERCA2 was significantly downregulated in all three cardiac chambers including the SAN region in rainbow trout that reset fH after three weeks of warm acclimation, indicating a reduced role of SERCA2 after warm acclimation (Sutcliffe et al., 2020). Further, one group of trout unexpectedly did not reset their fH after the same acclimation period, and did not present a change in SERCA2 expression. Furthermore, RYR3 was downregulated in fish that reset fH after three weeks of warm acclimation in the ventricle and the atrium, but not the SAN (Sutcliffe et al., 2020). Therefore, the calcium  176 clock likely plays a variable role in cardiac pacemaking of the rainbow trout for the resetting of fH as a function of acclimation temperature. The role of the SR in cardiac pacemaking merits further investigation. Pharmacological block of RyR and SERCA pumps during intracellular recording of pacemaker cell electrical activity will provide definitive evidence of the role of the SR in pacemaking. My thesis provides indirect evidence for the important role played by the SR due to the effects of the pharmacological blocking agents used on the SR of the working myocardium. A reduced cardiac preparation consisting of the SAN, ducti of Cuvier and the atrium, allows access to the endocardial side of the heart and direct access to the pacemaker cells would provide direct insight into the role of the SR.   A coupled clock mechanism The current and ongoing debate in the mammalian literature concerns the origins of the pacemaking current and the involvement and interplay of the membrane and calcium clocks (DiFrancesco, 2020). My results in the zebrafish support a coupled clock mechanism and have revealed the existence of a third clock-like mechanism that drives a residual fH when the membrane and calcium clocks are pharmacologically blocked. Such a finding to my knowledge has only been hinted towards in a SERCA2 knockout mouse model (Logantha et al., 2016). More evidence is required to determine the exact origin of this third clock, but it may involve NCX as the current has the ability to drive the current in both forward and reverse mode. The very slow ~20 beats min-1 could be driven by transmembrane concentrations of Ca2+ and Na+ that locally activate the NCX and drive the slow depolarization of the membrane potentials. Intracellular recording of pacemaker cell electrophysiological activity in the presence of an NCX blocker ORM-1096 would provide valuable insight into the residual pacemaking mechanism discovered in the zebrafish (Kohajda et  177 al., 2016; Kohajda et al., 2020). Alternatively, this slow residual fH could be driven by residual Hcn channels, not blocked by HCN blockers which may not be as effective in blocking all Hcn isoforms in fish as in mammals. Further research is needed on both the efficiency and specificity of HCN channel blockers in fish and on possible alternative pacemaking mechanisms in fish and mammals.   5.4 The importance of Hcn4 in cardiac pacemaking of the zebrafish As the major pacemaking mechanism in mammals is the membrane clock and the major HCN isoform generating the If current is HCN4, I hypothesized that in the zebrafish, Hcn4 plays a major role in cardiac pacemaking and that a Hcn4 knockout would significantly reduce fH, resulting in chronic bradycardia. I discovered that Hcn4 is not essential to pacemaking of the adult zebrafish as fHmax was unchanged in the Hcn4 knockout fish compared with the wild-type fish, independent of the test temperature that I used. Furthermore, the swimming and respirometry capacity of the knockout zebrafish were not compromised by the Hcn4 knockout. However, zatebradine significantly reduced fH of the knockout fish, by the same degree as in the wild-type fish, indicating total compensation of Hcn4 in pacemaking. Compensation likely occurred through other Hcn isoforms because SR-Ca2+ cycling was not significantly different between wild-type and knockout fish.  Interestingly, sinoatrial pauses accompanied with sinoatrial arrest are reported in 5 dpf Hcn4 knockout zebrafish, and exposure to ivabradine results in a dose-dependent increase in fH variability and decreased fH (from 229 ± 37 beats min-1 to 173 ± 27 beats min-1 with 25 μM ivabradine) (von der Heyde et al., 2020). These results may be accounted for by the fact that hcn4l is a possible paralogue of hcn4 which can compensate for the loss of hcn4 in the knockout fish.  178 However, functional expression analysis needs to be performed to confirm the functional redundancy between the two proteins.  Important homeostatic mechanisms differ between adult and juvenile stages and many do not fully develop until after embryogenesis (Warren et al., 2001). Therefore, gene functions in the embryo are not necessarily the same in the adult, for example, there is a dramatic difference in fH between zebrafish embryos (~170 beats min-1) and adults (~120 beats min-1) (Liu et al., 2016; Pylatiuk et al., 2014). HCN expression in newborn mammals is high but then decreases throughout development up to adulthood (Biel et al., 2009). Furthermore, HCN4 knockout is lethal in embryogenesis in mice, but HCN4 controlled- knockout in adults produces only a mild phenotype, despite a ~80% reduction in If current  (Biel et al., 2009; Herrmann et al., 2007). Having reared my Hcn4 knockout fish to adulthood, perhaps only those that had successfully compensated for the loss of Hcn4 survived. Alternatively, all of the fish achieved compensation over the 3 months rearing period to adulthood prior to conducting the experiments and the rearing phase may have served as a bottleneck-type selection phase for those that had successfully compensated for the knockout of Hcn4. In order to fully understand the mechanisms of compensation that allow the Hcn4 knockout fish to maintain cardiac performance, measurements at regular intervals throughout development should be made, including fH and mRNA expression analysis of membrane and calcium clock-related genes. Indeed, no upregulation of alternative Hcn channels was seen in the adult fish, indicating that compensation does not occur through changes in gene expression at the adult stage. Compensation may occur instead through changes in regulatory RNA or vesicular storage and membrane trafficking of channel proteins that were not investigated in my thesis or may involve mechanisms involving the formation of heteromeric channels between alternative Hcns (Zhang et al., 2009). Given the extremely small size of the SAN, reportedly  179 consisting of a small ring of cells (Haverinen and Vornanen, 2007; Newton et al., 2014; Stoyek et al., 2015; Stoyek et al., 2016), the zebrafish heart posed a significant challenge for resolving the SAN and atrial tissues for mRNA expression analysis. There is therefore some overlap between SAN and atrial tissue used in the qPCR analysis of the mRNA. Resolving these tissues would require isolation and identification of single pacemaker cells followed by single-cell RNA sequencing (Linscheid et al., 2019). The amenability of the zebrafish to genome engineering renders specific fluorescent tagging of pacemaker cells feasible and has already been developed (Poon et al., 2016). Fluorescent labelled cells can be sorted using high throughput methods such as flow cytometry followed by single-cell RNA sequencing or single-cell qPCR and may offer a means by which to develop cell-type-specific mRNA analysis.   The relative importance of Hcn channels in cardiac pacemaking should be further investigated to enhance our understanding of the mechanisms that drive pacemaking in the zebrafish. In order to determine the relative roles of the different Hcn isoforms, knockouts of multiple Hcn isoforms should be generated, where the Hcn4 knockout is coupled with the knockout of Hcn1, Hcn2 or Hcn3. Pharmacological inhibition will not provide the necessary information due to the considerable overlap between Hcn channels and the effects on multiple other ion channels (i.e. potassium channels).   5.5 Thermal tolerance of the zebrafish pacemaker and temperature-dependent deterioration of the cardiac electrical signal Thermal tolerance of the pacemaking mechanism in vivo was determined using surface ECG recordings obtained from anesthetized fish. The advantage of the setup is that the heart and the pericardium are intact and relevant parameters such as filling pressure and volume are  180 conserved as well as physiologically relevant oxygenation of the myocardium. Of concern, MS-222 is known to reduce fH in the anesthetized fish in vivo, underestimating the maximum fH and potentially overestimating cardiac thermal tolerance (Huang et al., 2010). However, the effect of MS-222 is mediated by increased vagal tone, which is inhibited by atropine in the fHmax induced fish. In addition, MS-222  has been reported to inhibit the cardiac INa of the zebrafish by up to 30% when used at 168 mg L-1 and thus may result in a prolongation of the AP upstroke (Haverinen et al., 2018b). However, I used a substantially lower concentration in my thesis work, thus the effects of MS-222 remain negligible as the effect of MS-222 have been reported to produce only minimal alterations to the ECG (Topic Popovic et al., 2012).  I attributed the absence of a QRS signal to the loss of ventricular excitation-contraction and the loss of atrial excitation-contraction would be attributed to the loss of the P wave. Furthermore, loss of a P wave could indicate either the loss of atrial excitation or the loss of pacemaker excitation, or the inability for the signal to propagate from the SAN to the atrium due to the high resistance of the SAN. However, using ECGs there is no metric for the loss of pacemaker cell excitation and thus the pacemaker thermal tolerance cannot be estimated with this method as the measure of pacemaking activity is not direct. My thesis can therefore only provide information on the electrical excitation of the ventricle and the atrium; pacemaking is unlikely to be interrupted at the same temperature as the atrium as the thermal tolerance of isolated pacemaker cells has been shown in the brown trout to be superior to that of the adjacent atrial tissue (Haverinen et al., 2017). Temperature-dependent collapse of cardiac function may thus occur within the working myocardium and could arise from contraction failure or electrical conduction failure. Ventricular contraction failure may occur at high temperature as the ventricle is larger and requires more time to contract and relax. In addition, the ventricle has a longer refractory period than the  181 atrium and thus cannot be stimulated for contraction, whereas the atrium has a shorter refractory period (Vornanen, 2017). On the ECG recording, this may be represented as the P wave merging into the QRS complex. However, reports of temperature-dependent cardiac collapse, including this thesis, have not reported such occurrence, and instead have reported atrioventricular block shown as missing QRS complexes on the ECG with a preserved P wave (Badr et al., 2016; Haverinen and Vornanen, 2020b; Marchant and Farrell, 2019). As the AP is slowed in the atrioventricular region due to a lower expression of fast depolarizing Na+ channels, leading to low INa density (Hassinen et al., 2021), the atrioventricular node represents a potential site for cardiac arrhythmias at high temperature as Na+ channels have been shown to have the lowest temperature tolerance of the cardiac ion channels (Vornanen et al., 2014). A recent study has quantified the expression levels of 41 ion channel genes in different regions of the rainbow trout heart and showed that SCN4Aa, SCN4Aba, SCN4Abb, SCN5LAba, SCN5LAbb, and SCN5LAa, encoding for Nav1.4a Nav1.4ba Nav1.4bb Nav1.5a Nav1.5ba Nav1.5bb proteins respectively, were all more highly expressed at the ventricular side of the atrioventricular node (Hassinen et al., 2021) whilst the total Kir2 mRNA abundance was lower in the ventricular side of the atrioventricular node (Vornanen, 2020). These data suggest a source-sink mismatch hypothesis wherein the depolarizing INa current is insufficient to overcome the larger current density of the repolarizing K+ currents which results in atrioventricular block at the site where expression of the Na+ channels (Vornanen, 2020). The reason for this source-sink mismatch at high temperature occurs due to the lower thermal tolerance of the Na+ channels which has been shown to be 20.9 ± 0.5°C in the brown trout while the thermal tolerance of Ca2+ and K+ channels exceeds 28°C (Vornanen et al., 2014). In addition to this hypothesis and because of higher metabolic rates at high temperatures, the accumulation of lactate (excreted from metabolic active tissues) may lead to acidosis which can hinder the influx and  182 efflux of Ca2+ through VG-ion channels or the NCX and decrease the response of contractile elements to Ca2+ (Schwieterman et al., 2021; Shiels et al., 2010).  Finally, although understudied, connexins that form gap junctions, the non-specific ion channels that electrically couple neighbouring myocytes may also represent a point of conduction failure within the myocardium. This possibility cannot be excluded by simple means of ECG recordings as the point of conduction failure cannot be identified. In the zebrafish, mutation of connexin 43 (Cx43) by removal of amino acids 256-289 resulted in impaired gap junction endocytosis and increased gap junction intracellular communication which has for effect severe heart malformations with disorganized and malformed vasculature, elongated hearts, decreased fH, and impaired blood flow (Hyland et al., 2021). Deletion of Cx43 in mice has previously been shown to severely interfere with communication between the right ventricle and the outflow tract, impairing blood flow and resulting in neonatal death (Reaume et al., 1995).  Future studies on cardiac pacemaking of the zebrafish should aim to better determine the thermal tolerance of the pacemaker cells via the intracellular recording of the pacemaker cell electrical activity. This will however require an ex vivo preparation as intracellular recordings in vivo are not possible.  In my thesis, I, therefore, validated a method that would best allow reliable recording of pacemaker cell electrical activity in the context of interconnected SAN and atrial tissue which will significantly contribute to future advances in fish cardiac pacemaker electrophysiology. Future investigations on the mechanisms of cardiac pacemaking should use intracellular recordings and, using the respective pharmacological blockers used in this thesis, record the effect of blocking the different clocks directly at the level of the pacemaker cell.      183 5.6 Summary Fish cardiac pacemaking remains unresolved due to the complexity of interacting mechanisms. Although clear advances have been made, future research should focus on identifying genes that encode important proteins of the membrane and calcium clocks and determine the interplay between these two mechanisms. My thesis calls for further investigation on HCN channel blockers in fish and leaves room for a third and unidentified pacemaking mechanism, which could in fact further bridge the membrane and calcium clock in the fish heart. This third clock may in fact be caused by residual pacemaker cells that were not fully blocked by the pharmacological agents that were developed for use in mammals and some of the channels, pumps or receptors of the different clock mechanisms of the zebrafish SAN may have different sensitivity to these pharmacological agents. Alternatively, a third clock truly does exist and may be driven by forward and reverse mode NCX. However, this third mechanism would only occur in non-physiological states (blocked membrane and calcium clock) and NCX is already an integral part of the coupled clock system, and thus, alone does not constitute a clock mechanism.  To better advance the field, and until these mechanisms have been resolved, I believe that a model species of cardiac pacemaking needs to be developed. The fully sequenced genome, rapid generation times, and ease of genetic manipulation of the zebrafish enable many physiological and genomic investigations that are not easily accessible in other fish species. Therefore, following the Krogh principle, the zebrafish appears to be an ideal organism for electrophysiological investigation of cardiac pacemaking.   184 Bibliography Abramochkin, D. V., Hassinen, M. and Vornanen, M. (2018). Transcripts of Kv7.1 and MinK channels and slow delayed rectifier K+ current (IKs) are expressed in zebrafish (Danio rerio) heart. Pflugers Arch. Eur. J. Physiol. 470, 1753–1764. Abramochkin, D. V., Haverinen, J., Mitenkov, Y. A. and Vornanen, M. (2019). Temperature and external K+ dependence of electrical excitation in ventricular myocytes of cod-like fishes. J. Exp. Biol. jeb193607. Accili, E. A., Proenza, C., Baruscotti, M. and DiFrancesco, D. (2002). From funny current to HCN channels: 20 Years of excitation. News Physiol. Sci. 17, 32–37. Aho, E. and Vornanen, M. (1998). Ca2+-ATPase activity and Ca2+ uptake by sarcoplasmic reticulum in fish heart: effects of thermal acclimation. J. Exp. Biol. 201, 525–32. Aho, Vornanen, Aho, E. and Vornanen, M. (1999). Contractile properties of atrial and ventricular myocardium of the heart of rainbow trout Oncorhynchus mykiss: effects of thermal acclimation. J. Exp. Biol. 202, 2663–2677. Alday, A., Alonso, H., Gallego, M., Urrutia, J., Letamendia, A., Callol, C. and Casis, O. (2014). Ionic channels underlying the ventricular action potential in zebrafish embryo. Pharmacol. Res. 84, 26–31. Allingham, J. S., Smith, R. and Rayment, I. (2005). The structural basis of blebbistatin inhibition and specificity for myosin II. Nat. Struct. Mol. Biol. 12, 378–379. Altimiras, J. and Axelsson, M. (2004). Intrinsic autoregulation of cardiac output in rainbow trout (Oncorhynchus mykiss) at different heart rates. J. Exp. Biol. 207, 195–201. Altomare, C., Terragni, B., Brioschi, C., Milanesi, R., Pagliuca, C., Viscomi, C., Moroni, A., Baruscotti, M. and DiFrancesco, D. (2003). Heteromeric HCN1-HCN4 channels: A  185 comparison with native pacemaker channels from the rabbit sinoatrial node. J. Physiol. 549, 347–359. Amores, A., Force, A., Yan, Y., Joly, L., Amemiya, C., Ho, R. K., Langeland, J., Prince, V., Wang, Y., Ekker, M., et al. (1998). Zebrafish hox Clusters and Vertebrate Genome Evolution. Science (80-. ). 282, 1711–1714.a Andersson, K. B., Birkeland, J. A. K., Finsen, A. V., Louch, W. E., Sjaastad, I., Wang, Y., Chen, J., Molkentin, J. D., Chien, K. R., Sejersted, O. M., et al. (2009). Moderate heart dysfunction in mice with inducible cardiomyocyte-specific excision of the Serca2 gene. J. Mol. Cell. Cardiol. 47, 180–187. Anttila, K., Casselman, M. T., Schulte, P. M. and Farrell, A. P. (2013). Optimum Temperature in Juvenile Salmonids: Connecting Subcellular Indicators to Tissue Function and Whole-Organism Thermal Optimum. Physiol. Biochem. Zool. 86, 245–256. Anttila, K., Couturier, C., Øverli, Ø., Johnsen, A., Marthinsen, G., Nilsson, G. . and Farrell, A. . (2014). Atlantic salmon show capability for cardiac acclimation to warm temperatures. Nat. Commun. 5, 4252. Araya, C., Häkkinen, H. M., Carcamo, L., Cerda, M., Savy, T., Rookyard, C., Peyriéras, N. and Clarke, J. D. W. (2019). Cdh2 coordinates Myosin-II dependent internalisation of the zebrafish neural plate. Sci. Rep. 9, 1–13. Arnaout, R., Ferrer, T., Huisken, J., Spitzer, K., Stainier, D. Y. R., Tristani-Firouzi, M. and Chi, N. C. (2007). Zebrafish model for human long QT syndrome. Proc. Natl. Acad. Sci. 104, 11316–11321. Azene, E. M., Xue, T., Marbán, E., Tomaselli, G. F. and Li, R. A. (2005). Non-equilibrium behavior of HCN channels: Insights into the role of HCN channels in native and engineered  186 pacemakers. Cardiovasc. Res. 67, 263–273. Baartscheer, A., Schumacher, C. A., Coronel, R. and Fiolet, J. W. T. (2011). The driving force of the Na+/Ca2+-exchanger during: Metabolic inhibition. Front. Physiol. MAR, 1–7. Badr, A., El-Sayed, M. F. and Vornanen, M. (2016).  Effects of seasonal acclimatization on temperature dependence of cardiac excitability in the roach, Rutilus rutilus . J. Exp. Biol. 219, 1495–1504. Badr, A., Hassinen, M., El-Sayed, M. F. and Vornanen, M. (2017). Effects of seasonal acclimatization on action potentials and sarcolemmal K+ currents in roach (Rutilus rutilus) cardiac myocytes. Comp. Biochem. Physiol. -Part A  Mol. Integr. Physiol. 205, 15–27. Badr, A., Korajoki, H., Abu-Amra, E. S., El-Sayed, M. F. and Vornanen, M. (2018). Effects of seasonal acclimatization on thermal tolerance of inward currents in roach (Rutilus rutilus) cardiac myocytes. J. Comp. Physiol. B Biochem. Syst. Environ. Physiol. 188, 255–269. Baker, K., Warren, K. S., Yellen, G. and Fishman, M. C. (1997). Defective “pacemaker” current (Ih) in a zebrafish mutant with a slow heart rate. Proc. Natl. Acad. Sci. 94, 4554–4559. Bakker, M. L., Christoffels, V. M. and Moorman, A. F. M. (2010). The cardiac pacemaker and conduction system develops from embryonic myocardium that retains its primitive phenotype. J. Cardiovasc. Pharmacol. 56, 6–15. Bakkers, J. (2011). Zebrafish as a model to study cardiac development. Cardiovasc. Res. 91, 279–288. Balami, S., Sharma, A. and Karn, R. (2019). Significance of nutritional value of fish for human health. Malaysian J. Halal Res. 2, 32–34. Bang, A., Grønkjær, P. and Malte, H. (2004). Individual variation in the rate of oxygen  187 consumption by zebrafish embryos. J. Fish Biol. 64, 1285–1296. Baruscotti, M., Bucchi, A. and DiFrancesco, D. (2005). Physiology and pharmacology of the cardiac pacemaker (“funny”) current. Pharmacol. Ther. 107, 59–79. Baruscotti, M., Bucchi, A., Viscomi, C., Mandelli, G., Consalez, G., Gnecchi-Rusconi, T., Montano, N., Casali, K. R., Micheloni, S., Barbuti, A., et al. (2011). Deep bradycardia and heart block caused by inducible cardiac-specific knockout of the pacemaker channel gene Hcn4. Proc. Natl. Acad. Sci. 108, 1705–1710. Baruscotti, M., Bucchi, A., Milanesi, R., Paina, M., Barbuti, A., Gnecchi-Ruscone, T., Bianco, E., Vitali-Serdoz, L., Cappato, R. and DiFrancesco, D. (2017). A gain-of-function mutation in the cardiac pacemaker HCN4 channel increasing cAMP sensitivity is associated with familial Inappropriate Sinus Tachycardia. Eur. Heart J. 38, 280–288. Baudenbacher, F., Schober, T., Pinto, J. R., Sidorov, V. Y., Hilliard, F., Solaro, R. J., Potter, J. D. and Knollmann, B. C. (2008). Myofilament Ca2+ sensitization causes susceptibility to cardiac arrhythmia in mice. J. Clin. Invest. 118, 3893–3903. Becker, C. D. and Genoway, R. G. (1979). Evaluation of the critical thermal maximum for determining thermal tolerance of freshwater fish. Environ. Biol. Fishes 4, 245–256. Beitinger, T. L. and Bennett, W. A. (2000). Quantification of the role of acclimation temperature in temperature tolerance of fishes. Environ. Biol. Fishes 58, 277–288. Beitinger, T. L., Bennett, W. A. and Mccauley, R. W. (2000). Temperature tolerances of North American freshwater fishes exposed to dynamic changes in temperature. Environ. Biol. ofFishes 58, 237–275. Bers, D. M. (1993). Excitation-Contraction Coupling. In Excitation-Contraction Coupling and Cardiac Contractile Force, pp. 119–148. Dordrecht: Springer Netherlands.  188 Bers, D. M. (2001). Excitation-contraction coupling and cardiac contractile force. (ed. Publishers, K. A.) Dordrecht, The Netherlands: Springer Science & Business Media. Bers, D. M. (2002). Cardiac excitation–contraction coupling. Nature 415, 198–205. Bezanilla, F. (2005). Voltage-gated ion channels. IEEE Trans. Nanobioscience 4, 34–48. Biel, M., Wahl-Schott, C., Michalakis, S. and Zong, X. (2009). Hyperpolarization-Activated Cation Channels: From Genes to Function. Physiol. Rev. 89, 847–885. Bogdanov, K. Y., Vinogradova, T. M. and Lakatta, E. G. (2001). Sinoatrial Nodal Cell Ryanodine Receptor and Na+-Ca2+ Exchanger Molecular: Partners in Pacemaker Regulation. Circ. Res. 88, 1254–1258. Bournele, D. and Beis, D. (2016). Zebrafish models of cardiovascular disease. Heart Fail. Rev. 21, 803–813. Bovo, E., Dvornikov, A. V., Mazurek, S. R., De Tombe, P. P. and Zima, A. V. (2013). Mechanisms of Ca2+ handling in zebrafish ventricular myocytes. Pflugers Arch. Eur. J. Physiol. 465, 1775–1784. Brack, K. E., Narang, R., Winter, J. and Ng, G. A. (2013). The mechanical uncoupler blebbistatin is associated with significant electrophysiological effects in the isolated rabbit heart. Exp. Physiol. 98, 1009–1027. Brett, J. R. (1964). The Respiratory Metabolism and Swimming Performance of Young Sockeye Salmon. J. Fish. Res. Board Canada 21, 1183–1226. Brett, J. R. (1971). Energetic Responses of Salmon to Temperature. A Study of Some Thermal Relations in the Physiology and Freshwater Ecology of Sockeye Salmon (Oncorhynchus nerka). Fish. Res. 11, 99–113. Brette, F., Luxan, G., Cros, C., Dixey, H., Wilson, C. and Shiels, H. A. (2008).  189 Characterization of isolated ventricular myocytes from adult zebrafish (Danio rerio). Biochem. Biophys. Res. Commun. 374, 143–146. Briggs, J. P. (2002). The zebrafish: a new model organism for integrative physiology. Am J Physiol Regul. Integr. Comp Physiol 282, R3–R9. Brioschi, C., Micheloni, S., Tellez, J. O., Pisoni, G., Longhi, R., Moroni, P., Billeter, R., Barbuti, A., Dobrzynski, H., Boyett, M. R., et al. (2009). Distribution of the pacemaker HCN4 channel mRNA and protein in the rabbit sinoatrial node. J. Mol. Cell. Cardiol. 47, 221–227. Brown, B. Y. H. and DiFrancesco, D. (1980). Voltage-clamp investigations of membrane currents underlying pacemaker activity in rabbit sino-atrial node. J. Physiol. 308, 331–351. Brown, H. F., DiFrancesco, D. and Noble, S. J. (1979). How does adrenaline accelarete the heart? Nature 280, 235. Bucchi, A., Barbuti, A., DiFrancesco, D. and Baruscotti, M. (2012). Funny current and cardiac rhythm: Insights from HCN knockout and transgenic mouse models. Front. Physiol. 3, 1–10. Burkhard, S., Eif, V. V, Garric, L., Christoffels, V. and Bakkers, J. (2017). On the Evolution of the Cardiac Pacemaker. J. Cardiovasc. Dev. Dis. 4, 4. Carmen, V., Eva, D., Laura, F., Pilar, G., Onésima, P., Juan, T., J., S. D., Valenzuela, C., Delpo´n, E., Franqueza, L., et al. (1996). Class III Antiarrhythmic Effects of Zatebradine. Circulation 94, 562–570. Casselman, M. T., Anttila, K. and Farrell, A. P. (2012). Using maximum heart rate as a rapid screening tool to determine optimum temperature for aerobic scope in Pacific salmon Oncorhynchus spp. J. Fish Biol. 80, 358–377.  190 Chabot, D., Steffensen, J. F. and Farrell, A. P. (2016). The determination of standard metabolic rate in fishes. J. Fish Biol. 88, 81–121. Chai, J., Hamilton, A. L., Krieg, M., Buckley, C. D., Riedel-Kruse, I. H. and Dunn, A. R. (2015). A Force Balance Can Explain Local and Global Cell Movements during Early Zebrafish Development. Biophys. J. 109, 407–414. Chandler, N. J., Greener, I. D., Tellez, J. O., Inada, S., Musa, H., Molenaar, P., DiFrancesco, D., Baruscotti, M., Longhi, R., Anderson, R. H., et al. (2009). Molecular architecture of the human sinus node insights into the function of the cardiac pacemaker. Circulation 119, 1562–1575. Chen, J. N., Haffter, P., Odenthal, J., Vogelsang, E., Brand, M., Van Eeden, F. J. M., Furutani-Seiki, M., Granato, M., Hammerschmidt, M., Heisenberg, C. P., et al. (1996). Mutations affecting the cardiovascular system and other internal organs in zebrafish. Development 123, 293–302. Cheng, H., Lederer, W. J. and Cannell, M. B. (1993). Calcium Sparks: Elementary Events Underlying Excitation-Contraction Coupling in Heart Muscle. Science (80-. ). 262, 740–744. Chi, N. C., Shaw, R. M., Jungblut, B., Huisken, J., Ferrer, T., Arnaout, R., Scott, I., Beis, D., Xiao, T., Baier, H., et al. (2008). Genetic and physiologic dissection of the vertebrate cardiac conduction system. PLoS Biol. 6, 1006–1019. Chopra, S. S., Watanabe, H., Zhong, T. P. and Roden, D. M. (2007). Molecular cloning and analysis of zebrafish voltage-gated sodium channel beta subunit genes: Implications for the evolution of electrical signaling in vertebrates. BMC Evol. Biol. 7,. Clark, T. D., Sandblom, E., Cox, G. K., Hinch, S. G. and Farrell, A. P. (2008). Circulatory  191 limits to oxygen supply during an acute temperature increase in the Chinook salmon (Oncorhynchus tshawytscha). Am. J. Physiol. Integr. Comp. Physiol. 295, R1631–R1639. Clark, T. D., Jeffries, K. M., Hinch, S. G. and Farrell, A. P. (2011). Exceptional aerobic scope and cardiovascular performance of pink salmon (Oncorhynchus gorbuscha) may underlie resilience in a warming climate. J. Exp. Biol. 214, 3074–3081. Conradsen, C., Walker, J. A., Perna, C. and McGuigan, K. (2016). Repeatability of locomotor performance and morphology-locomotor performance relationships. J. Exp. Biol. 219, 2888–2897. Cortemeglia, C. and Beitinger, T. L. (2005). Temperature Tolerances of Wild-Type and Red Transgenic Zebra Danios. Trans. Am. Fish. Soc. 134, 1431–1437. Cros, C., Sallé, L., Warren, D. E., Shiels, H. A. and Brette, F. (2014). The calcium stored in the sarcoplasmic reticulum acts as a safety mechanism in rainbow trout heart. Am J Physiol Regul Integr Comp Physiol 307, 1491–1501. D’souza, A., Bucchi, A., Johnsen, A. B., Logantha, S. J. R. J., Monfredi, O., Yanni, J., Prehar, S., Hart, G., Cartwright, E., Wisloff, U., et al. (2014). Exercise training reduces resting heart rate via downregulation of the funny channel HCN4. Nat. Commun. 5,. Denvir, M. A., Tucker, C. S. and Mullins, J. J. (2008). Systolic and diastolic ventricular function in zebrafish embryos: Influence of norepenephrine, MS-222 and temperature. BMC Biotechnol. 8, 1–8. Dickson, K. A. and Graham, J. B. (2004). Evolution and Consequences of Endothermy in Fishes. Physiol. Biochem. Zool. 77, 998–1018. DiFrancesco, D. (1981). A study of the ionic nature of the pace‐maker current in calf Purkinje fibres. J. Physiol. 314, 377–393.  192 DiFrancesco, D. (1991). The contribution of the ‘pacemaker’ current (If) to generation of spontaneous activity in rabbit sino‐atrial node myocytes. J. Physiol. 434, 23–40. DiFrancesco, D. (1993). Pacemaker Mechanisms in Cardiac Tissue. Annu. Rev. Physiol. 55, 455–472. DiFrancesco, D. (2010). The role of the funny current in pacemaker activity. Circ. Res. 106, 434–446. DiFrancesco, D. (2020). A Brief History of Pacemaking. Front. Physiol. 10, 1–7. DiFrancesco, D. and Noble, D. (2012). The funny current has a major pacemaking role in the sinus node. Hear. Rhythm 9, 299–301. DiFrancesco, D., Ferroni, A., Mazzanti, M. and Tromba, C. (1986). Properties of the hyperpolarizing-activated current (If) in cells isolated from the rabbit sino-atrial node. J. Physiol. 377, 61–88. Dimitriadi, A., Beis, D., Arvanitidis, C., Adriaens, D. and Koumoundouros, G. (2018). Developmental temperature has persistent, sexually dimorphic effects on zebrafish cardiac anatomy. Sci. Rep. 8, 1–10. Doench, J. G., Hartenian, E., Graham, D. B., Tothova, Z., Hegde, M., Smith, I., Sullender, M., Ebert, B. L., Xavier, R. J. and Root, D. E. (2014). Rational design of highly active sgRNAs for CRISPR-Cas9-mediated gene inactivation. Nat. Biotechnol. 32, 1262–1267. Doerr, T. and Trautwein, W. (1990). On the mechanism of the “specific bradycardic action” of the verapamil derivative UL-FS 49. Naunyn. Schmiedebergs. Arch. Pharmacol. 341, 331–340. Dou, Y., Arlock, P. and Arner, A. (2007). Blebbistatin specifically inhibits actin-myosin interaction in mouse cardiac muscle. Am. J. Physiol. - Cell Physiol. 293, 1148–1153.  193 Drost, H. E., Carmack, E. C. and Farrell, A. P. (2014). Upper thermal limits of cardiac function for Arctic cod Boreogadus saida , a key food web fish species in the Arctic. J. Fish Biol. 6, 1781–1792. Drost, H. E., Lo, M., Carmack, E. C. and Farrell, A. P. (2016). Acclimation potential of Arctic cod ( Boreogadus saida ) from the rapidly warming Arctic Ocean. J. Exp. Biol. 219, 3114–3125. Efimov, I. R., Nikolski, V. P. and Salama, G. (2004). Optical imaging of the heart. Circ. Res. 94, 21–33. Ekström, A., Brijs, X. J., Clark, T. D., Gräns, A., Jutfelt, F. and Sandblom, E. (2016a). Cardiac oxygen limitation during an acute thermal challenge in the European perch: effects of chronic environmental warming and experimental hyperoxia. 311, 440–449. Ekström, A., Hellgren, K., Gräns, A., Pichaud, N. and Sandblom, E. (2016b). Dynamic changes in scope for heart rate and cardiac autonomic control during warm acclimation in rainbow trout. J. Exp. Biol. 219, 1106–1109. El-Brolosy, M. A. and Stainier, D. Y. R. (2017). Genetic compensation: A phenomenon in search of mechanisms. PLoS Genet. 13, 1–17. Eliason, E. J., Clark, T. D., Hague, M. J., Hanson, L. M., Gallagher, Z. S., Jeffries, K. M., Gale, M. K., Patterson, D. A., Hinch, S. G. and Farrell, A. P. (2011). Differences in thermal tolerance among sockeye salmon populations. Science (80-. ). 332, 109–112. Eliason, E. J., Clark, T. D., Hinch, S. G. and Farrell, A. P. (2013). Cardiorespiratory collapse at high temperature in swimming adult sockeye salmon. Conserv. Physiol. 1, 1–19. Erdfelder, E., FAul, F., Buchner, A. and Lang, A. G. (2009). Statistical power analyses using G*Power 3.1: Tests for correlation and regression analyses. Behav. Res. Methods 41, 1149– 194 1160. Fangue, N. A., Osborne, E. J., Todgham, A. E. and Schulte, P. M. (2011). The onset temperature of the heat-shock response and whole-organism thermal tolerance are tightly correlated in both laboratory-acclimated and field-acclimatized tidepool sculpins (Oligocottus maculosus). Physiol. Biochem. Zool. 84, 341–352. Farman, G. P., Tachampa, K., Mateja, R., Cazorla, O., Lacampagne, A. and De Tombe, P. P. (2008). Blebbistatin: Use as inhibitor of muscle contraction. Pflugers Arch. Eur. J. Physiol. 455, 995–1005. Farrell, A. P. (1991). From Hagfish to Tuna: A Perspective on Cardiac Function in Fish. Source Physiol. Zool. 64, 1137–1164. Farrell, A. P. (2009). Environment, antecedents and climate change: lessons from the study of temperature physiology and river migration of salmonids. J. Exp. Biol. 212, 3771–3780. Farrell, A. P. and Jones, D. R. (1992). The heart. In Fish physiology (ed. Hoar, W. S.), Randall, D. J.), and Farrell, A. P.), pp. 1–88. New York: Academic press. Farrell, A. P. and Smith, F. (2017). Cardiac Form, Function and Physiology. In Fish Physiology, pp. 155–264. Elsevier Inc. Farrell, A. P., Johansen, J. . and Graham, M. . (1988). The Role of the Pericardium in Cardiac Performance of the Trout (Salmo gairdneri). Physiol. Zool. 61, 213–221. Farrell, A. ., Gamperl, A. ., Hicks, J. M. ., Shiels, H. . and Jain, K. . (1996). Maximum cardiac performance of rainbow trout (Oncorhynchus mykiss) at temperatures approaching their upper lethal limit. J. Exp. Biol. 199, 663–672. Farrell, A. P., Hinch, S. G., Cooke, S. J., Patterson, D. A., Crossin, G. T., Lapointe, M. and Mathes, M. T. (2008). Pacific Salmon in Hot Water: Applying Aerobic Scope Models and  195 Biotelemetry to Predict the Success of Spawning Migrations. Physiol. Biochem. Zool. 81, 697–709. Farrell, A. P., Eliason, E. J., Sandblom, E. and Clark, T. D. (2009). Fish cardiorespiratory physiology in an era of climate change. Can. J. Zool. 87, 835–851. Fedorov, V. V., Lozinsky, I. T., Sosunov, E. A., Anyukhovsky, E. P., Rosen, M. R., Balke, C. W. and Efimov, I. R. (2007). Application of blebbistatin as an excitation-contraction uncoupler for electrophysiologic study of rat and rabbit hearts. Hear. Rhythm 4, 619–626. Fedorov, V. V., Glukhov, A. V., Chang, R., Kostecki, G., Aferol, H., Hucker, W. J., Wuskell, J. P., Loew, L. M., Schuessler, R. B., Moazami, N., et al. (2010). Optical mapping of the isolated coronary-perfused human sinus node. J. Am. Coll. Cardiol. 56, 1386–1394. Fedorov, V. V., Glukhov, A. V., Ambrosi, C. M., Kostecki, G., Chang, R., Janks, D., Schuessler, R. B., Moazami, N., Nichols, C. G. and Efimov, I. R. (2011). Effects of KATP channel openers diazoxide and pinacidil in coronary-perfused atria and ventricles from failing and non-failing human hearts. J. Mol. Cell. Cardiol. 51, 215–225. Fenton, F. H., Cherry, E. M. and Kornreich, B. G. (2008). Termination of equine atrial fibrillation by quinidine: An optical mapping study. J. Vet. Cardiol. 10, 87–103. Ferreira, E. O., Anttila, K. and Farrell, A. P. (2014). Thermal Optima and Tolerance in the Eurythermic Goldfish ( Carassius auratus ): Relationships between Whole-Animal Aerobic Capacity and Maximum Heart Rate. Physiol. Biochem. Zool. 87, 599–611. Fry, F. E. J. (1947). Effect of the environment on animal activity. Toronto, ON: University of Toronto press. Fry, F. E. J. (1971). The effect of environmental factors on the physiology of fish. In Fish  196 Physiology (ed. Hoar, W. S. & Randall, D. J.), pp. 1–98. New York: Academic Press. Fry, F. . E. . J. . and Hart, J, S. (1948). The Relation of Temperature to Oxygen Consumption in the Goldfish. Biol. Bull. 94, 66–77. Gamperl, A. K. and Farrell, A. . (2004). Cardiac plasticity in fishes: environmental influences and intraspecific differences. J. Exp. Biol. 207, 2539–2550. Gamperl, A. K., Swafford, B. L. and Rodnick, K. J. (2011). Elevated temperature, per se, does not limit the ability of rainbow trout to increase stroke volume. J. Therm. Biol. 36, 7–14. Garcia-Frigola, C., Shi, Y. and Evans, S. M. (2003). Expression of the hyperpolarization-activated cyclic nucleotide-gated cation channel HCN4 during mouse heart development. Gene Expr. Patterns 3, 777–783. Garrott, K., Kuzmiak-Glancy, S., Wengrowski, A., Zhang, H., Rogers, J. and Kay, M. W. (2017). KATP channel inhibition blunts electromechanical decline during hypoxia in left ventricular working rabbit hearts. J. Physiol. 595, 3799–3813. Genge, C. E., Lin, E., Lee, L., Sheng, X., Rayani, K., Gunawan, M., Stevens, C. M., Li, A. Y., Talab, S. S., Claydon, T. W., et al. (2016). The Zebrafish Heart as a Model of Mammalian Cardiac Function. In Reviews of physiology, biochemestry and pharmacology, pp. 99–136. Gerger, C. J., Thomas, J. K., Janz, D. M. and Weber, L. P. (2015). Acute effects of β-naphthoflavone on cardiorespiratory function and metabolism in adult zebrafish (Danio rerio). Fish Physiol. Biochem. 41, 289–298. Giardoglou, P. and Beis, D. (2019). On zebrafish disease models and matters of the heart. Biomedicines 7,. Gilbert, M. J. H., Rani, V., McKenzie, S. M. and Farrell, A. P. (2019). Autonomic cardiac  197 regulation facilitates acute heat tolerance in rainbow trout: In situ and in vivo support. J. Exp. Biol. 222, 1–10. Glukhov, A. V, Fedorov, V. V, Lou, Q., Ravikumar, V. K., Kalish, P. W., Schuessler, R. B., Moazami, N. and Eifmov, I. R. (2010). Transmural dispersion of repolarization in failing and nonfailing human ventricle. Circ. Res. 106, 981–991. Goethals, M., Raes, A. and van Bogaert, P. P. (1993). Use-dependent block of the pacemaker current If in rabbit sinoatrial node cells by zatebradine (UL-FS 49). Circulation 88, 2389–2401. Goldspink, G. (1995). Adaptation of fish to different environmental temperature by qualitative and quantitative changes in gene expression. J. Therm. Biol. 20, 167–174. Gollock, M. J., Currie, S., Petersen, L. H. and Gamperl, A. K. (2006). Cardiovascular and haematological responses of Atlantic cod (Gadus morhua) to acute temperature increase. J. Exp. Biol. 209, 2961–2970. Graham, A. M. S. and Farrell, A. P. (1989). The Effect of Temperature Acclimation and Adrenaline on the Performance of a Perfused Trout Heart. Physiol. Zool. 62, 38–61. Gupta, P., Martin, R., Knölker, H. J., Nihalani, D. and Sinha, D. K. (2017). Myosin-1 inhibition by PClP affects membrane shape, cortical actin distribution and lipid droplet dynamics in early Zebrafish embryos. PLoS One 12, 1–21. Hagiwara, N., Irisawa, H. and Kameyama, M. (1988). Contribution of two types of calcium currents to the pacemaker potentials of rabbit sino-atrial node cells. J. Physiol. 395, 233–253. Hanson, L. M., Obradovich, S., Mouniargi, J. and Farrell, A. P. (2006). The role of adrenergic stimulation in maintaining maximum cardiac performance in rainbow trout  198 (Oncorhynchus mykiss) during hypoxia, hyperkalemia and acidosis at 10°C. J. Exp. Biol. 209, 2442–2451. Harper, A. A., Newton, I. P. and Watt, P. W. (1995). The effect of temperature on spontaneous action potential discharge of the isolated sinus venosus from winter and summer plaice (Pleuronectes platessa). J. Exp. Biol. 198, 137–140. Harzheim, D., Pfeiffer, K. H., Fabritz, L., Kremmer, E., Buch, T., Waisman, A., Kirchhof, P., Kaupp, U. B. and Seifert, R. (2008). Cardiac pacemaker function of HCN4 channels in mice is confined to embryonic development and requires cyclic AMP. EMBO J. 27, 692–703. Hassel, D., Scholz, E. P., Trano, N., Friedrich, O., Just, S., Meder, B., Weiss, D. L., Zitron, E., Marquart, S., Vogel, B., et al. (2008). Deficient zebrafish ether-à-go-go-related gene channel gating causes short-QT syndrome in zebrafish reggae mutants. Circulation 117, 866–875. Hassinen, M., Paajanen, V., Haverinen, J., Eronen, H. and Vornanen, M. (2007). Cloning and expression of cardiac Kir2.1 and Kir2.2 channels in thermally acclimated rainbow trout. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 292, 2328–2339. Hassinen, M., Haverinen, J. and Vornanen, M. (2008). Electrophysiological properties and expression of the delayed rectifier potassium (ERG) channels in the heart of thermally acclimated rainbow trout. AJP Regul. Integr. Comp. Physiol. 295, 297–308. Hassinen, M., Haverinen, J., Hardy, M. E., Shiels, H. A. and Vornanen, M. (2015). Inward rectifier potassium current (IK1) and Kir2 composition of the zebrafish (Danio rerio) heart. Pflugers Arch. Eur. J. Physiol. 467, 2437–2446. Hassinen, M., Haverinen, J., Vornanen, M. and Physiology, C. (2017). Small functional If  199 current in sinoatrial pacemaker cells of the brown trout (Salmo trutta fario) heart despite strong expression of HCN channel transcripts. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 313, 711–722. Hassinen, M., Dzhumaniiazova, I., Abramochkin, D. V. and Vornanen, M. (2021). Ionic basis of atrioventricular conduction: ion channel expression and sarcolemmal ion currents of the atrioventricular canal of the rainbow trout (Oncorhynchus mykiss) heart. J. Comp. Physiol. B 191, 327–346. Haverinen, J. and Vornanen, M. (2007). Temperature acclimation modifies sinoatrial pacemaker mechanism of the rainbow trout heart. Am J Physiol Regul Integr Comp Physiol 292 292, 169–169. Haverinen, J. and Vornanen, M. (2009). Responses of Action Potential and K+ Currents to Temperature Acclimation in Fish Hearts: Phylogeny or Thermal Preferences? Physiol. Biochem. Zool. 82, 468–482. Haverinen, J. and Vornanen, M. (2020a). Reduced ventricular excitability causes atrioventricular block and depression of heart rate in fish at critically high temperatures. J. Exp. Biol. 223,. Haverinen, J. and Vornanen, M. (2020b). Atrioventricular block, due to reduced ventricular excitability, causes the depression of fish heart rate in fish at critically high temperatures. J. Exp. Biol. 223,. Haverinen, J., Abramochkin, D. V., Kamkin, A. and Vornanen, M. (2017). Maximum heart rate in brown trout ( Salmo trutta fario ) is not limited by firing rate of pacemaker cells. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 312, R165–R171. Haverinen, J., Hassinen, M., Dash, S. N. and Vornanen, M. (2018a). Expression of calcium  200 channel transcripts in the zebrafish heart: dominance of T-type channels. J. Exp. Biol. 221,. Haverinen, J., Hassinen, M., Korajoki, H. and Vornanen, M. (2018b). Cardiac voltage-gated sodium channel expression and electrophysiological characterization of the sodium current in the zebrafish (Danio rerio) ventricle. Prog. Biophys. Mol. Biol. 138, 59–68. Heath, A. G. and Hughes, G. M. (1973). Cardiovascular and Respiratory Changes During Heat Stress in Rainbow Trout (Salmo Gairdneri). J. Exp. Biol. 59, 323–338. Herrmann, S., Stieber, J., Stöckl, G., Hofmann, F. and Ludwig, A. (2007). HCN4 provides a “depolarization reserve” and is not required for heart rate acceleration in mice. EMBO 26, 4423–4432. Hille, B. (2001). Ion Channels of Excitable Membranes. 3rd ed. Sunderland, MA: Sinauer Associates. Hochachka, P. W. and Somero, G. N. (2002). Biochemical adaptation : Mechanism and Process in Physiological Evolution. New York: Oxford University Press. Hoegh-guldberg, O. (2010). The Impact of Climate Change on the World’s Marine Ecosystems. 1523, 1523–1529. Hokanson, K. E. F., Kleiner, C. F. and Thorslund, T. W. (1977). Effects of Constant Temperatures and Diel Temperature Fluctuations on Specific Growth and Mortality Rates and Yield of Juvenile Rainbow Trout, Salmo gairdneri. J. Fish. Res. Board Canada 34, 639–648. Houston, B. Y. A. H. and Koss, T. F. (1984). Plasma and Red Cell Ionic Composition in Rainbow Trout Exposed To Progressive Temperature Increases. J. Exp. Biol. 110, 53–67. Hove-Madsen, L. and Tort, L. (1998). L-type Ca2+ current and excitation-contraction coupling in single atrial myocytes from rainbow trout. Am J Physiol 275, 2061–2069.  201 Hove-Madsen, L., Llach, A. and Tort, L. (1999). Quantification of calcium release from the sarcoplasmic reticulum in rainbow trout atrial myocytes. Pflugers Arch. Eur. J. Physiol. 438, 545–552. Howe, K., Clark, M. D., Torroja, C. F., Torrance, J., Berthelot, C., Muffato, M., Collins, J. E., Humphray, S., McLaren, K., Matthews, L., et al. (2013). The zebrafish reference genome sequence and its relationship to the human genome. Nature 496, 498–503. Huang, W. C., Hsieh, Y. S., Chen, I. H., Wang, C. H., Chang, H. W., Yang, C. C., Ku, T. H., Yeh, S. R. and Chuang, Y. J. (2010). Combined use of MS-222 (Tricaine) and isoflurane extends anesthesia time and minimizes cardiac rhythm side effects in adult zebrafish. Zebrafish 7, 297–304. Hüser, J., Blatter, L. A. and Lipsius, S. L. (2000). Intracellular Ca2+ release contributes to automaticity in cat atrial pacemaker cells. J. Physiol. 524, 415–422. Hyland, C., Mfarej, M., Hiotis, G., Lancaster, S., Novak, N., Kathryn Iovine, M. and Falk, M. M. (2021). Impaired Cx43 gap junction endocytosis causes cardiovascular defects in zebrafish. bioRxiv 2021.03.07.434329. Irisawa, H. (1978). Comparative physiology of the cardiac pacemaker mechanism. Physiol. Rev. 58, 461. Ishii, T. M., Takano, M., Xie, L. H., Noma, A. and Ohmori, H. (1999). Molecular characterization of the hyperpolarization-activated cation channel in rabbit heart sinoatrial node. J. Biol. Chem. 274, 12835–12839. Ishii, T. M., Takano, M. and Ohmori, H. (2001). Determinants of activation kinetics in mammalian hyperpolarization-activated cation channels. J. Physiol. 537, 93–100. Jensen, D. (1965). The aneural heart of the hagfish. Ann. New York Acad. Sci. 127, 443–458.  202 Jensen, B., Wang, T., Christoffels, V. M. and Moorman, A. F. M. (2013). Evolution and development of the building plan of the vertebrate heart. Biochim. Biophys. Acta - Mol. Cell Res. 1833, 783–794. Jinek, M., Chylinski, K., Fonfara, I., Hauer, M., Doudna, J. A. and Charpentier, E. (2012). A Programmable Dual-RNA – Guided DNA Endonuclease in Adaptive Bacterial Immunity. Science (80-. ). 337, 816–822. Johnston, I. A., Lee, H. T., Macqueen, D. J., Paranthaman, K., Kawashima, C., Anwar, A., Kinghorn, J. R. and Dalmay, T. (2009). Embryonic temperature affects muscle fibre recruitment in adult zebrafish: Genome-wide changes in gene and microRNA expression associated with the transition from hyperplastic to hypertrophic growth phenotypes. J. Exp. Biol. 212, 1781–1793. Jou, C. J., Spitzer, K. W., Tristani-Firouzi, M. and Jerry Jou, C. (2010). Blebbistatin Effectively Uncouples the Excitation-Contraction Process in Zebrafish Embryonic Heart. Cell. Physiol. Biochem. 25, 419–424. Joung, B., Chen, P. S. and Lin, S. F. (2011). The role of the calcium and the voltage clocks in sinoatrial node dysfunction. Yonsei Med. J. 52, 211–219. Ju, Y. K. and Allen, D. G. (2000). The distribution of calcium in toad cardiac pacemaker cells during spontaneous firing. Pflugers Arch. Eur. J. Physiol. 441, 219–227. Jutfelt, F., Roche, D. G., Clark, T. D., Norin, T., Binning, S. A., Speers-Roesch, B., Amcoff, M., Morgan, R., Andreassen, A. H. and Sundin, J. (2019). Brain cooling marginally increases acute upper thermal tolerance in Atlantic cod. J. Exp. Biol. 222, 1–5. Kanlop, N. and Sakai, T. (2010). Optical mapping study of blebbistatin-induced chaotic electrical activities in isolated rat atrium preparations. J. Physiol. Sci. 60, 109–117.  203 Kappadan, V., Telele, S., Uzelac, I., Fenton, F., Parlitz, U., Luther, S. and Christoph, J. (2020). High-Resolution Optical Measurement of Cardiac Restitution, Contraction, and Fibrillation Dynamics in Beating vs. Blebbistatin-Uncoupled Isolated Rabbit Hearts. Front. Physiol. 11, 464–472. Kaupp, U. B. and Seifert, R. (2001). Molecular diversity of pacemaker ion channels. Crit. Rev. Oral Biol. Med. 63, 235–257. Keen, A. N. and Gamperl, A. K. (2012). Blood oxygenation and cardiorespiratory function in steelhead trout (Oncorhynchus mykiss) challenged with an acute temperature increase and zatebradine-induced bradycardia. J. Therm. Biol. 37, 201–210. Keith, A. and Flack, M. (1907). The Form and Nature of the Muscular Connections between the Primary Divisions of the Vertebrate Heart. J. Anat. Physiol. 41, 172–189. Keith, A. and Mackenzie, I. (1910). Recent Researches on the Anatomy of the Heart. Lancet 175, 101–103. Kettlewell, S., Walker, N. L., Cobbe, S. M., Burton, F. L. and Smith, G. L. (2004). The electrophysiological mechanical effects of 2,3-butane-dione monoxime and cytochalasin-D in the Langendorff perfused rabbit heart. Exp. Physiol. 89, 163–172. Klaiman, J. M., Fenna, A. J., Shiels, H. A., Macri, J. and Gillis, T. E. (2011). Cardiac remodeling in fish: Strategies to maintain heart function during temperature change. PLoS One 6, e24464. Knaus, A., Zong, X., Beetz, N., Jahns, R., Lohse, M. J., Biel, M. and Hein, L. (2007). Direct inhibition of cardiac hyperpolarization-activated cyclic nucleotide-gated pacemaker channels by clonidine. Circulation 115, 872–880. Kohajda, Z., Farkas-Morvay, N., Jost, N., Nagy, N., Geramipour, A., Horváth, A., Varga,  204 R. S., Hornyik, T., Corici, C., Acsai, K., et al. (2016). The effect of a novel highly selective inhibitor of the sodium/calcium exchanger (NCX) on cardiac arrhythmias in in vitro and in vivo experiments. PLoS One 11, e0166041. Kohajda, Z., Tóth, N., Szlovák, J., Loewe, A., Bitay, G., Gazdag, P., Prorok, J., Jost, N., Levijoki, J., Pollesello, P., et al. (2020). Novel Na+/Ca2+ exchanger inhibitor ORM-10962 supports coupled function of funny-current and Na+/ Ca2+ exchanger in pacemaking of rabbit sinus node tissue. Front. Pharmacol. 10, 1632. Kong, W., Ideker, R. E. and Fast, V. G. (2009). Transmural optical measurements of Vm dynamics during long-duration ventricular fibrillation in canine hearts. Hear. Rhythm 6, 796–802. Kosicki, M., Tomberg, K. and Bradley, A. (2018). Repair of double-strand breaks induced by CRISPR–Cas9 leads to large deletions and complex rearrangements. Nat. Biotechnol. 36, 765–771. Kovács, M., Tóth, J., Hetényi, C., Málnási-Csizmadia, A. and Seller, J. R. (2004). Mechanism of blebbistatin inhibition of myosin II. J. Biol. Chem. 279, 35557–35563. Kozasa, Y., Nakashima, N., Ito, M., Ishikawa, T., Kimoto, H., Ushijima, K., Makita, N. and Takano, M. (2018). HCN4 pacemaker channels attenuate the parasympathetic response and stabilize the spontaneous firing of the sinoatrial node. J. Physiol. 596, 809–825. Krueger, C. and Tian, L. (2004). A comparison of the general linear mixed model and repeated measures ANOVA using a dataset with multiple missing data points. Biol. Res. Nurs. 6, 151–157. Labun, K., Montague, T. G., Krause, M., Torres Cleuren, Y. N., Tjeldnes, H. and Valen, E. (2019). CHOPCHOP v3: expanding the CRISPR web toolbox beyond genome editing.  205 Nucleic Acids Res. 47, 171–174. Laish-Farkash, A., Glikson, M., Brass, D., Marek-Yagel, D., Pras, E., Dascal, N., Antzelevitch, C., Nof, E., Reznik, H., Eldar, M., et al. (2010). A novel mutation in the HCN4 gene causes symptomatic sinus bradycardia in Moroccan Jews. J. Cardiovasc. Electrophysiol. 21, 1365–1372. Lakatta, E. G. and DiFrancesco, D. (2009). What keeps us ticking: a funny current, a calcium clock, or both? J. Mol. Cell. Cardiol. 47, 157–170. Lakatta, E. G., Vinogradova, T. M. and Maltsev, V. A. (2008). The missing link in the mystery of normal automaticity of cardiac pacemaker cells. Ann. N. Y. Acad. Sci. 1123, 41–57. Lakatta, E. G., Maltsev, V. A. and Vinogradova, T. M. (2010). A Coupled SYSTEM of intracellular Ca2+ clocks and surface membrane voltage clocks controls the timekeeping mechanism of the heart’s pacemaker. Circ. Res. 106, 434–446. Langheinrich, U., Vacun, G. and Wagner, T. (2003). Zebrafish embryos express an orthologue of HERG and are sensitive toward a range of QT-prolonging drugs inducing severe arrhythmia. Toxicol. Appl. Pharmacol. 193, 370–382. Lee, C. H. and MacKinnon, R. (2017). Structures of the Human HCN1 Hyperpolarization-Activated Channel. Cell 168, 111–120. Lee, L., Genge, C. E., Cua, M., Sheng, X., Rayani, K., Beg, M. F., Sarunic, M. V. and Tibbits, G. F. (2016). Functional assessment of cardiac responses of adult zebrafish (Danio rerio) to acute and chronic temperature change using high-resolution echocardiography. PLoS One 11, e0145163. Lee, P., Quintanilla, J. G., Alfonso-Almazán, J. M., Galán-Arriola, C., Yan, P., Sánchez- 206 González, J., Pérez-Castellano, N., Pérez-Villacastín, J., Ibañez, B., Loew, L. M., et al. (2019). In vivo ratiometric optical mapping enables high-resolution cardiac electrophysiology in pig models. Cardiovasc. Res. 115, 1659–1671. Lees-Miller, J. P., Guo, J., Wang, Y., Perissinotti, L. L., Noskov, S. Y. and Duff, H. J. (2015). Ivabradine prolongs phase 3 of cardiac repolarization and blocks the hERG1 (KCNH2) current over a concentration-range overlapping with that required to block HCN4. J. Mol. Cell. Cardiol. 85, 71–78. Lewis, A. S., Estep, C. M. and Chetkovich, D. M. (2010). The fast and slow ups and downs of HCN channel regulation. Channels 4, 215–231. Lieschke, G. J. and Currie, P. D. (2007). Animal models of human disease: Zebrafish swim into view. Nat. Rev. Genet. 8, 353–367. Limouze, J., Straight, A. F., Mitchison, T. and Sellers, J. R. (2004). Specificity of blebbistatin, an inhibitor of myosin II. J. Muscle Res. Cell Motil. 25, 337–341. Lin, E., Ribeiro, A., Ding, W., Hove-Madsen, L., Sarunic, M. V., Beg, M. F. and Tibbits, G. F. (2014). Optical mapping of the electrical activity of isolated adult zebrafish hearts: acute effects of temperature. Am. J. Physiol. Regul. Integr. Comp. Physiol. 306, 823–836. Linscheid, N., Logantha, S. J. R. J., Poulsen, P. C., Zhang, S., Schrölkamp, M., Egerod, K. L., Thompson, J. J., Kitmitto, A., Galli, G., Humphries, M. J., et al. (2019). Quantitative proteomics and single-nucleus transcriptomics of the sinus node elucidates the foundation of cardiac pacemaking. Nat. Commun. 10, 2889. Liu, J. and Stainier, D. Y. R. (2012). Zebrafish in the study of early cardiac development. Circ. Res. 110, 870–874. Liu, Y., Cabo, C., Salomonsz, R., Delmar, M., Davidenko, J. and Jalife, J. (1993). Effects of  207 diacetyl monoxime on the electrical properties of sheep and guinea pig ventricular muscle. Cardiovasc. Res. 27, 1991–1997. Liu, C. C., Li, L., Lam, Y. W., Siu, C. W. and Cheng, S. H. (2016). Improvement of surface ECG recording in adult zebrafish reveals that the value of this model exceeds our expectation. Sci. Rep. 6, 25073. Llach, A., Molina, C. E., Alvarez-Lacalle, E., Tort, L., Benítez, R. and Hove-Madsen, L. (2011). Detection, properties, and frequency of local calcium release from the sarcoplasmic reticulum in teleost cardiomyocytes. PLoS One 6, 23708. Lochowitz, R. T., Miles, H. M. and Hafemann, D. R. (1974). Anesthetic-induced variations in the cardiac rate of the teleost, Salmo gairdneri. Comp. Gen. Pharmacol. 5, 217–224. Logantha, S. J. R. J., Stokke, M. K., Atkinson, A. J., Kharche, S. R., Parveen, S., Saeed, Y., Sjaastad, I., Sejersted, O. M. and Dobrzynski, H. (2016). Ca2+-clock-dependent pacemaking in the sinus node is impaired in mice with a cardiac specific reduction in SERCA2 abundance. Front. Physiol. 7, 197. López-Olmeda, J. F. and Sánchez-Vázquez, F. J. (2011). Thermal biology of zebrafish (Danio rerio). J. Therm. Biol. 36, 91–104. Lou, Q., Li, W. and Efimov, I. R. (2012). The role of dynamic instability and wavelength in arrhythmia maintenance as revealed by panoramic imaging with blebbistatin vs. 2,3-butanedione monoxime. Am. J. Physiol. - Hear. Circ. Physiol. 302, 262–269. Louch, W. E., Hougen, K., Mørk, H. K., Swift, F., Aronsen, J. M., Sjaastad, I., Reims, H. M., Roald, B., Andersson, K. B., Christensen, G., et al. (2010). Sodium accumulation promotes diastolic dysfunction in end-stage heart failure following Serca2 knockout. J. Physiol. 588, 465–478.  208 Lucas, J., Bonnieux, A., Lyphout, L., Cousin, X., Miramand, P. and Lefrançois, C. (2016). Trophic contamination by pyrolytic polycyclic aromatic hydrocarbons does not affect aerobic metabolic scope in zebrafish Danio rerio. J. Fish Biol. 88, 433–442. Ludwig, A., Zong, X., Stieber, J., Hullin, R., Hofmann, F. and Biel, M. (1999). Two pacemaker channels from human heart with profoundly different activation kinetics. EMBO J. 18, 2323–2329. Luo, C. H. and Rudy, Y. (1991). Original Contributions A Model of the Ventricular Cardiac Action Potential. Circ. Res. 68, 1501–1526. Maltsev, V. A. and Lakatta, E. G. (2007). Normal Heart Rhythm is Initiated and Regulated by an Intracellular Calcium Clock Within Pacemaker Cells. Hear. Lung Circ. 16, 335–348. Maltsev, V. A. and Lakatta, E. G. (2008). Dynamic interactions of an intracellular Ca2+ clock and membrane ion channel clock underlie robust initiation and regulation of cardiac pacemaker function. Cardiovasc. Res. 77, 274–284. Maltsev, V. A. and Lakatta, E. G. (2009). Synergism of coupled subsarcolemmal Ca2+ clocks and sarcolemmal voltage clocks confers robust and flexible pacemaker function in a novel pacemaker cell model. Am. J. Physiol. - Hear. Circ. Physiol. 296, 594–615. Maltsev, V. A., Yaniv, Y., Maltsev, A. V., Stern, M. D. and Lakatta, E. G. (2014). Modern perspectives on numerical modeling of cardiac pacemaker cell. J. Pharmacol. Sci. 125, 6–38. Mangoni, M. E. and Nargeot, J. (2008). Genesis and Regulation of the Heart Automaticity. Physiol. Rev. 88, 919–982. Mangoni, M. E., Traboulsie, A., Leoni, A. L., Couette, B., Marger, L., Le Quang, K., Kupfer, E., Cohen-Solal, A., Vilar, J., Shin, H. S., et al. (2006). Bradycardia and slowing  209 of the atrioventricular conduction in mice lacking CaV3.1/α1G T-type calcium channels. Circ. Res. 98, 1422–1430. Männikkö, R., Elinder, F. and Larsson, H. P. (2002). Voltage-sensing mechanism is conserved among ion channels gated by opposite voltages. Nature 419, 837–841. Marchant, J. L. L. and Farrell, A. P. P. (2019). Membrane and calcium clock mechanisms contribute variably as a function of temperature to setting cardiac pacemaker rate in zebrafish Danio rerio. J. Fish Biol. 95, 1265–1274. Marionneau, C., Couette, B., Liu, J., Li, H., Mangoni, M. E., Nargeot, J., Lei, M., Escande, D. and Demolombe, S. (2005). Specific pattern of ionic channel gene expression associated with pacemaker activity in the mouse heart. J. Physiol. 562, 223–234. Masahito, O., Hisakazu, K., Yumiko, O., Atsushi, M., Hideyo, O. and Michio, M. (1994). The Expression, Phosphorylation, and Localization of Connexin 43 and Gap-Junctional Intercellular Communication during the Establishment of a Synchronized Contraction of Cultured Neonatal Rat Cardiac Myocytes. Exp. Cell Res. 351–358. McCluskey, B. M. and Braasch, I. (2020). Zebrafish Phylogeny and Taxonomy. In The Zebrafish in Biomedical Research (ed. Cartner, S. C.), Farmer, S. C.), Kent, M. L.), Eisen, J. S.), Guillemin, K. J.), and Sanders, G. E.), pp. 15–24. Elsevier Inc. Mendonça, P. C. and Gamperl, A. K. (2010). The effects of acute changes in temperature and oxygen availability on cardiac performance in winter flounder (Pseudopleuronectes americanus). Comp. Biochem. Physiol. - A Mol. Integr. Physiol. 155, 245–252. Meyer, A. and Van De Peer, Y. (2005). From 2R to 3R: Evidence for a fish-specific genome duplication (FSGD). BioEssays 27, 937–945. Milan, D. J., Jones, I. L., Ellinor, P. T. and MacRae, C. A. (2006). In vivo recording of adult  210 zebrafish electrocardiogram and assessment of drug-induced QT prolongation. AJP Hear. Circ. Physiol. 291, 269–273. Milan, D. J., Kim, A. M., Winterfield, J. R., Jones, I. L., Pfeufer, A., Sanna, S., Arking, D. E., Amsterdam, A. H., Sabeh, K. M., Mably, J. D., et al. (2009). Drug-sensitized zebrafish screen identifies multiple genes, including GINS3, as regulators of myocardial repolarization. Circulation 120, 553–559. Milanesi, R., Baruscotti, M., Gnecchi-Ruscone, T. and DiFrancesco, D. (2006). Familial sinus bradycardia associated with a mutation in the cardiac pacemaker channel. N. Engl. J. Med. 354, 151–157. Mistrik, P., Mader, R., Michalakis, S., Weidinger, M., Pfeifer, A. and Biel, M. (2005). The murine HCN3 gene encodes a hyperpolarization-activated cation channel with slow kinetics and unique response to cyclic nucleotides. J. Biol. Chem. 280, 27056–27061. Mittelbach, G. G., Osenberg, C. W. and Wainwright, P. C. (1992). Variation in resource abundance affects diet and feeding morphology in the pumpkinseed sunfish (Lepomis gibbosus). Oecologia 90, 8–13. Mobli, M., Undheim, E. A. B. and Rash, L. D. (2017). Chapter Seven - Modulation of Ion Channels by Cysteine-Rich Peptides: From Sequence to Structure. In Ion Channels DownUnder (ed. Geraghty, D. P.) and Rash, L. D. B. T.-A. in P.), pp. 199–223. Academic Press. Monfredi, O., Maltsev, V. A. and Lakatta, E. G. (2013). Modern concepts concerning the origin of the heartbeat. Physiology 28, 74–92. Moorman, A. F. M. and Christoffels, V. M. (2003). Cardiac chamber formation: Development, genes, and evolution. Physiol. Rev. 83, 1223–1267.  211 Moorman, A. F. M., Christoffels, V. M., Anderson, R. H. and Van Den Hoff, M. J. B. (2007). The heart-forming fields: One or multiple? Philos. Trans. R. Soc. B Biol. Sci. 362, 1257–1265. Moosmang, S., Stieber, J., Zong, X., Biel, M., Hofmann, F., Ludwig, A., Stieber, J., Zong, X. and Biel, M. (2001). Cellular expression and functional characterization of four hyperpolarization-activated pacemaker channels in cardiac and neuronal tissues. Eur. J. Biochem. 268, 1646–1652. Morad, M. and Goldman, Y. (1973). Excitation-contraction coupling in heart muscle: Membrane control of development of tension. Prog. Biophys. Mol. Biol. 27, 257–313. Morgan, R., Sundin, J., Finnøen, M. H., Dresler, G., Vendrell, M. M., Dey, A., Sarkar, K. and Jutfelt, F. (2019). Are model organisms representative for climate change research? Testing thermal tolerance in wild and laboratory zebrafish populations. Conserv. Physiol. 7, 36. Morozov, S., Scott McCairns, R. J. and Merilä, J. (2019). FishResp: R package and GUI application for analysis of aquatic respirometry data. Conserv. Physiol. 7, 3. Much, B., Wahl-Schott, C., Zong, X., Schneider, A., Baumann, L., Moosmang, S., Ludwig, A. and Biel, M. (2003). Role of Subunit Heteromerization and N-Linked Glycosylation in the Formation of Functional Hyperpolarization-activated Cyclic Nucleotide-gated Channels. J. Biol. Chem. 278, 43781–43786. Nemtsas, P., Wettwer, E., Christ, T., Weidinger, G. and Ravens, U. (2010). Adult zebrafish heart as a model for human heart? An electrophysiological study. J. Mol. Cell. Cardiol. 48, 161–171. Nerbonne, J. M. and Kass, R. S. (2005). Molecular physiology of cardiac repolarization.  212 Physiol. Rev. 85, 1205–1253. Newton, C. M., Stoyek, M. R., Croll, R. P. and Smith, F. M. (2014). Regional innervation of the heart in the goldfish, Carassius auratus: A confocal microscopy study. J. Comp. Neurol. 522, 456–478. Novak, A. E., Taylor, A. D., Pineda, R. H., Lasda, E. L., Wright, M. A. and Ribera, A. B. (2006). Embryonic and larval expression of zebrafish voltage-gated sodium channel α-subunit genes. Dev. Dyn. 235, 1962–1973. Nyboer, E. A. and Chapman, L. J. (2018). Cardiac plasticity influences aerobic performance and thermal tolerance in a tropical, freshwater fish at elevated temperatures. J. Exp. Biol. 221, 178087. Palstra, A. P., Tudorache, C., Rovira, M., Brittijn, S. A., Burgerhout, E., van den Thillart, G. E. E. J. M., Spaink, H. P. and Planas, J. V. (2010). Establishing zebrafish as a novel exercise model: Swimming economy, swimming-enhanced growth and muscle growth marker gene expression. PLoS One 5, 14483. Pape, H.-C. (1996). Queer Current and Pacemaker: The Hyperpolarization-Activated Cation Current in Neurons. Annu. Rev. Physiol. 58, 299–327. Payne, A. I. and Temple, S. A. (1996). River and floodplain fisheries in the ganges basin. final report, marine resources assessment group ltd, overseas development administration, r.5485, uk. 154. Peterson, M. E., Chen, F., Saven, J. G., Roos, D. S., Babbitt, P. C. and Sali, A. (2009). Evolutionary constraints on structural similarity in orthologs and paralogs. Protein Sci. 18, 1306–1315. Plaut, I. (2000). Effects of fin size on swimming performance, swimming behaviour and routine  213 activity of zebrafish Danio rerio. J. Exp. Biol. 203, 813–820. Poon, K. L., Liebling, M., Kondrychyn, I., Brand, T. and Korzh, V. (2016). Development of the cardiac conduction system in zebrafish. Gene Expr. Patterns 21, 89–96. Pörtner, H. O., Peck, M. A. and Ecophysiology, I. (2010). Climate change effects on fishes and fisheries: Towards a cause-and-effect understanding. J. Fish Biol. 77, 1745–1779. Pylatiuk, C., Sanchez, D., Mikut, R., Alshut, R., Reischl, M., Hirth, S., Rottbauer, W. and Just, S. (2014). Automatic zebrafish heartbeat detection and analysis for zebrafish embryos. Zebrafish 11, 379–383. Qu, J., Altomare, C., Bucchi, A., DiFrancesco, D. and Robinson, R. B. (2002). Functional comparison of HCN isoforms expressed in ventricular and HEK 293 cells. Pflugers Arch. Eur. J. Physiol. 444, 597–601. Randall, D. J. (1970). 4 The Circulatory System. Fish Physiol. 4, 133–172. Rauscher, A., Gyimesi, M., Kovács, M. and Málnási-Csizmadia, A. (2018). Targeting Myosin by Blebbistatin Derivatives: Optimization and Pharmacological Potential. Trends Biochem. Sci. 43, 700–713. Reaume, A. G., de Sousa, P. A., Kulkarni, S., Langille, B. L., Zhu, D., Davies, T. C., Juneja, S. C., Kidder, G. M. and Rossant, J. (1995). Cardiac malformation in neonatal mice lacking connexin43. Science (80-. ). 267, 1831–1834. Rivaroli, L., Rantin, F. T. and Kalinin, A. L. (2006). Cardiac function of two ecologically distinct Neotropical freshwater fish : Curimbata , Prochilodus lineatus ( Teleostei , Prochilodontidae), and trahira, Hoplias malabaricus (Teleostei, Erythrinidae). Comp. Biochem. Physiol. Part A 145, 322–327. Robinson, R. B. and Siegelbaum, S. A. (2003). Hyperpolarization-Activated Cation Currents:  214 From Molecules to Physiological Function. Annu. Rev. Physiol. 65, 453–480. Rottbauer, W., Baker, K., Wo, Z. G., Mohideen, M. A. P. K., Cantiello, H. F. and Fishman, M. C. (2001). Growth and Function of the Embryonic Heart Depend upon the Cardiac-Specific L-Type Calcium Channel α1 Subunit. Dev. Cell 1, 265–275. Rubenstein, D. S. and Lipsius, S. L. (1989). Mechanisms of automaticity in subsidiary pacemakers from cat right atrium. Circ. Res. 64, 648–657. Rueckschloss, U. and Isenberg, G. (2001). Cytochalasin D reduces Ca2+ currents via cofilin-activated depolymerization of F-actin in guinea-pig cardiomyocytes. J. Physiol. 537, 363–370. Safi, H., Zhang, Y., Schulte, P. M. and Farrell, A. P. (2019). The effect of acute warming and thermal acclimation on maximum heart rate of the common killifish Fundulus heteroclitus. J. Fish Biol. 95, 1441–1446. Saito, T. (1969). Electrophysiological Studies on the Pacemaker of Several Fish Hearts. Zool. Mag. 78, 291-296. Saito, T. (1973). Effects of vagal stimulation on the pacemaker action potentials of carp heart. Comp. Biochem. Physiol. Part A Physiol. 44, 191–199. Saito, T. and Tenma, K. (1976). Effects of left and right vagal stimulation on excitation and conduction of the carp heart (Cyprinus carpio). J. Comp. Physiol. 111, 39–53. Sakamoto, T., Limouze, J., Combs, C. A., Straight, A. F. and Sellers, J. R. (2005). Blebbistatin, a myosin II inhibitor, is photoinactivated by blue light. Biochemistry 44, 584–588. Sampurna, B. P., Audira, G., Juniardi, S., Lai, Y. H. and Hsiao, C. Der (2018). A simple ImageJ-based method to measure cardiac rhythm in zebrafish embryos. Inventions 3, 1–11.  215 Sandblom, E., Clark, T. D., Gräns, A., Ekström, A., Brijs, J., Sundström, L. F., Odelström, A., Adill, A., Aho, T. and Jutfelt, F. (2016). Physiological constraints to climate warming in fish follow principles of plastic floors and concrete ceilings. Nat. Commun. 7, 11447. Sander, J. D. and Joung, J. K. (2014). CRISPR-Cas systems for editing, regulating and targeting genomes. Nat. Biotechnol. 32, 347–350. Sanhueza, D., Montoya, A., Sierralta, J. and Kukuljan, M. (2009). Expression of voltage-activated calcium channels in the early zebrafish embryo. Zygote 17, 131–135. Santoro, B., Liu, D. T., Yao, H., Bartsch, D., Kandel, E. R., Siegelbaum, S. A. and Tibbs, G. R. (1998). Identification of a gene encoding a hyperpolarization-activated pacemaker channel of brain. Cell 93, 717–729. Santoro, B., Chen, S., Lüthi, A., Pavlidis, P., Shumyatsky, G. P., Tibbs, G. R. and Siegelbaum, S. A. (2000). Molecular and functional heterogeneity of hyperpolarization-activated pacemaker channels in the mouse CNS. J. Neurosci. 20, 5264–5275. Santoso, F., Sampurna, B. P., Lai, Y. H., Liang, S. T., Hao, E., Chen, J. R. and Hsiao, C. Der (2019). Development of a simple imagej-based method for dynamic blood flow tracking in Zebrafish embryos and its application in drug toxicity evaluation. Inventions 4,. Sartiani, L., Mannaioni, G., Masi, A., Novella Romanelli, M. and Cerbai, E. (2017). The Hyperpolarization-Activated Cyclic Nucleotide–Gated Channels: from Biophysics to Pharmacology of a Unique Family of Ion Channels. Pharmacol. Rev. 69, 354–395. Satoh, H. (2003). Sino-Atrial Nodal Cells of Mammalian Hearts: Ionic Currents and Gene Expression of Pacemaker Ionic Channels. J. Smooth Muscle Res. 39, 175–193. Schaefer, J. and Ryan, A. (2006). Developmental plasticity in the thermal tolerance of zebrafish Danio rerio. J. Fish Biol. 69, 722–734.  216 Schnurr, M. E., Yin, Y. and Scott, G. R. (2014). Temperature during embryonic development has persistent effects on metabolic enzymes in the muscle of zebrafish. J. Exp. Biol. 217, 1370–1380. Schulte, P. M., Healy, T. M. and Fangue, N. A. (2011). Thermal performance curves, phenotypic plasticity, and the time scales of temperature exposure. Integr. Comp. Biol. 51, 691–702. Schulze-bahr, E., Pongs, O., Isbrandt, D., Schulze-bahr, E., Neu, A., Friederich, P., Kaupp, U. B., Breithardt, G., Pongs, O. and Isbrandt, D. (2003). Pacemaker channel dysfunction in a patient with sinus node disease Find the latest version : with sinus node disease. 111, 1537–1545. Schweizer, P. A., Thomas, D., Zehelein, J., Katus, H. A., Yampolsky, P., Malik, R. and Koenen, M. (2009). Transcription profiling of HCN-channel isotypes throughout mouse cardiac development. Basic Res. Cardiol. 104, 621–629. Schwieterman, G. D., Winchester, M. M., Shiels, H. A., Bushnell, P. G., Bernal, D., Marshall, H. M. and Brill, R. W. (2021). The effects of elevated potassium, acidosis, reduced oxygen levels, and temperature on the functional properties of isolated myocardium from three elasmobranch fishes: clearnose skate (Rostroraja eglanteria), smooth dogfish (Mustelus canis), and sandbar s. J. Comp. Physiol. B Biochem. Syst. Environ. Physiol. 191, 127–141. Scott, G. R. and Johnston, I. A. (2012). Temperature during embryonic development has persistent effects on thermal acclimation capacity in zebrafish. Proc. Natl. Acad. Sci. 109, 14247–14252. Sedmera, D., Reckova, M., DeAlmeida, A., Sedmerova, M., Biermann, M., Volejnik, J.,  217 Sarre, A., Raddatz, E., McCarthy, R. A., Gourdie, R. G., et al. (2003). Functional and morphological evidence for a ventricular conduction system in zebrafish and Xenopus hearts. Am. J. Physiol. - Hear. Circ. Physiol. 284, 1152–1160. Seebacher, F., White, C. R. and Franklin, C. E. (2015). Physiological plasticity increases resilience of ectothermic animals to climate change. Nat. Clim. Chang. 5, 61–66. Sehnert, A. J., Huq, A., Weinstein, B. M., Walker, C., Fishman, M. and Stainier, D. Y. R. (2002). Cardiac troponin T is essential in sarcomere assembly and cardiac contractility. Nat. Genet. 31, 106–110. Shelford, V. . (1931). Some Concepts of Bioecology. Ecology 12, 455–467. Shi, W., Wymore, R. T. R. T., Yu, H., Wu, J., Wymore, R. T. R. T., Pan, Z., Robinson, R. B., Dixon, J. E., McKinnon, D. and Cohen, I. S. (1999). Distribution and prevalence of hyperpolarization-activated cation channel (HCN) mRNA expression in cardiac tissues. Circ. Res. 85, 1–6. Shiels, H. A. (2017). Cardiomyocyte Morphology and Physiology. In Fish Physiology (ed. Gamperl, K. A.), Gillis, T. E.), Farrell, A. P.), and Brauner, C. J.), pp. 55–98. New York: Academic Press. Shiels, H. A. and Farrell, A. P. (2000). The effect of ryanodine on isometric tension development in isolated ventricular trabeculae from Pacific mackerel (Scomber japonicus). Comp. Biochem. Physiol. - A Mol. Integr. Physiol. 125, 331–341. Shiels, H. A. and Galli, G. L. J. J. (2014). The Sarcoplasmic Reticulum and the Evolution of the Vertebrate Heart. Physiology 29, 456–469. Shiels, H. A. and White, E. (2005). Temporal and spatial properties of cellular Ca2+ flux in trout ventricular myocytes. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 288, 1756–1766.  218 Shiels, H. A. H., Stevens, E. D. E. and Farrell, A. P. (1998). Effects of temperature, adrenaline and ryanodine on power production in rainbow trout oncorhynchus mykiss ventricular trabeculae. J. Exp. Biol. 201, 2701–10. Shiels, H. A., Freund, E. V, Farrell, A. P. and Block, B. A. (1999). The sarcoplasmic reticulum plays a major role in isometric contraction in atrial muscle of yellowfin tuna. J. Exp. Biol. 202, 881–890. Shiels, H. A., Blank, J. M., Farrell, A. P. and Block, B. A. (2004). Electrophysiological properties of the L-type Ca2+ current in cardiomyocytes from bluefin tuna and Pacific mackerel. Am. J. Physiol. Integr. Comp. Physiol. 286, 659–668. Shiels, H. A., Santiago, D. A. and Galli, G. L. J. (2010). Hypercapnic acidosis reduces contractile function in the ventricle of the armored catfish, pterygoplichthys pardalis. Physiol. Biochem. Zool. 83, 366–375. Shigekawa, M. and Iwamoto, T. (2001). Cardiac Na+-Ca2+ Exchange: Molecular and Pharmacological Aspects. Circ. Res. 88, 864–876. Shih, Y. H., Zhang, Y., Ding, Y., Ross, C. A., Li, H., Olson, T. M. and Xu, X. (2015). Cardiac Transcriptome and Dilated Cardiomyopathy Genes in Zebrafish. Circ. Cardiovasc. Genet. 8, 261–269. Shinohara, T., Joung, B., Kim, D., Maruyama, M., Luk, H. N., Chen, P. S. and Lin, S. F. (2010). Induction of atrial ectopic beats with calcium release inhibition: Local hierarchy of automaticity in the right atrium. Hear. Rhythm 7, 110–116. Sidhu, R., Anttila, K. and Farrell, A. P. (2014). Upper thermal tolerance of closely related Danio species. J. Fish Biol. 84, 982–995. Somero, G. N. (2005). Linking biogeography to physiology: Evolutionary and acclimatory  219 adjustments of thermal limits. Front. Zool. 2,. Somero, G. N. (2010). The physiology of climate change: how potentials for acclimatization and genetic adaptation will determine “winners” and “losers.” J. Exp. Biol. 213, 912–920. Spence, R., Gerlach, G., Lawrence, C. and Smith, C. (2008). The behaviour and ecology of the zebrafish, Danio rerio. Biol. Rev. 83, 13–34. Stainier, D. Y. R., Fouquet, B., Chen, J. N., Warren, K. S., Weinstein, B. M., Meiler, S. E., Mohideen, M. A. P. K., Neuhauss, S. C. F., Solnica-Krezel, L., Schier, A. F., et al. (1996). Mutations affecting the formation and function of the cardiovascular system in the zebrafish embryo. Development 123, 285–292. Stecyk, J. A. W., Paajanen, V., Farrell, A. P. and Vornanen, M. (2007). Effect of temperature and prolonged anoxia exposure on electrophysiological properties of the turtle (Trachemys scripta) heart. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 293, 421–437. Steffensen, J. F. (1989). Some errors in respirometry of aquatic breathers: How to avoid and correct for them. Fish Physiol. Biochem. 6, 49–59. Steinhausen, M. F., Sandblom, E., Eliason, E. J., Verhille, C. and Farrell, A. P. (2008). The effect of acute temperature increases on the cardiorespiratory performance of resting and swimming sockeye salmon (Oncorhynchus nerka). J. Exp. Biol. 211, 3915–3926. Stieber, J., Thomer, A., Much, B., Schneider, A., Biel, M. and Hofmann, F. (2003a). Molecular Basis for the Different Activation Kinetics of the Pacemaker Channels HCN2 and HCN4. J. Biol. Chem. 278, 33672–33680. Stieber, J., Herrmann, S., Feil, S., Löster, J., Feil, R., Biel, M., Hofmann, F. and Ludwig, A. (2003b). The hyperpolarization-activated channel HCN4 is required for the generation of pacemaker action potentials in the embryonic heart. Proc. Natl. Acad. Sci. U. S. A. 100,  220 15235–15240. Stieber, J., Hofmann, F. and Ludwig, A. (2004). Pacemaker Channels and Sinus Node Arrhythmia. Trends Cardiovasc Med. 14, 23–28. Stieber, J., Wieland, K., St, G., Ludwig, A. and Hofmann, F. (2006). Bradycardic and Proarrhythmic Properties of Sinus Node Inhibitors. Mol. Pharmacol. 69, 1328–1337. Stoyek, M. R. (2016). Autonomic innervation and control of chronotropy in the zebrafish heart. Stoyek, M. R., Croll, R. P. and Smith, F. M. (2015). Intrinsic and extrinsic innervation of the heart in zebrafish (Danio rerio). J. Comp. Neurol. 523, 1683–1700. Stoyek, M. R., Quinn, T. A., Croll, R. P. and Smith, F. M. (2016). Zebrafish heart as a model to study the integrative autonomic control of pacemaker function. Am. J. Physiol. Heart Circ. Physiol. 311, 676–688. Stoyek, M. R., Schmidt, M. K., Wilfart, F. M., Croll, R. P. and Smith, F. M. (2017). The in vitro zebrafish heart as a model to investigate the chronotropic effects of vapor anesthetics. Am. J. Physiol. Integr. Comp. Physiol. 313, 669–679. Straight, A. F., Cheung, A., Limouze, J., Chen, I., Westwood, N. J., Sellers, J. R. and Mitchison, T. J. (2003). Dissecting temporal and spatial control of cytokinesis with a myosin II inhibitor. Science (80-. ). 299, 1743–1747. Strecker, R., Seiler, T. B., Hollert, H. and Braunbeck, T. (2011). Oxygen requirements of zebrafish (Danio rerio) embryos in embryo toxicity tests with environmental samples. Comp. Biochem. Physiol. - C Toxicol. Pharmacol. 153, 318–327. Sun, P., Zhang, Y., Yu, F., Parks, E., Lyman, A., Wu, Q., Ai, L., Hu, C. H., Zhou, Q., Shung, K., et al. (2009a). Micro-electrocardiograms to study post-ventricular amputation of zebrafish heart. Ann. Biomed. Eng. 37, 890–901.  221 Sun, X., Hoage, T., Bai, P., Ding, Y., Chen, Z., Zhang, R., Huang, W., Jahangir, A., Paw, B., Li, Y. G., et al. (2009b). Cardiac hypertrophy involves both myocyte hypertrophy and hyperplasia in anemic zebrafish. PLoS One 4, 6596. Sutcliffe, R. L., Li, S., Gilbert, M. J. H., Schulte, P. M., Miller, K. M. and Farrell, A. P. (2020). A rapid intrinsic heart rate resetting response with thermal acclimation in rainbow trout, Oncorhynchus mykiss. J. Exp. Biol. 223, 215210. Svendsen, M. B. S., Bushnell, P. G. and Steffensen, J. F. (2016). Design and setup of intermittent-flow respirometry system for aquatic organisms. J. Fish Biol. 88, 26–50. Swift, L. M., Asfour, H., Posnack, N. G., Arutunyan, A., Kay, M. W. and Sarvazyan, N. (2012). Properties of blebbistatin for cardiac optical mapping and other imaging applications. Pflugers Arch. Eur. J. Physiol. 464, 503–512. Tacon, A. G. J. and Metian, M. (2013). Fish matters: importance of aquatic foods in human nutrition and global food supply. Rev. Fish. Sci. 21, 22–38. Tallarida, R. J. (2011). Quantitative methods for assessing drug synergism. Genes Cancer 2, 1003–1008. Tellez, J. O., Dobrzynski, H., Greener, I. D., Graham, G. M., Laing, E., Honjo, H., Hubbard, S. J., Boyett, M. R. and Billeter, R. (2006). Differential expression of ion channel transcripts in atrial muscle and sinoatrial node in rabbit. Circ. Res. 99, 1384–1393. Tessadori, F., van Weerd, J. H., Burkhard, S. B., Verkerk, A. O., de Pater, E., Boukens, B. J., Vink, A., Christoffels, V. M. and Bakkers, J. (2012). Identification and Functional Characterization of Cardiac Pacemaker Cells in Zebrafish. PLoS One 7, 47644. Tewksbury, J. J., Huey, R. B. and Deutsch, C. A. (2008). Putting the Heat on Tropical Animals. Science (80-. ). 320, 1296–1297.  222 Thollon, C., Cambarrat, C., Vian, J., Prost, J. ‐F, Peglion, J. L. and Vilaine, J. P. (1994). Electrophysiological effects of S 16257, a novel sino‐atrial node modulator, on rabbit and guinea‐pig cardiac preparations: comparison with UL‐FS 49. Br. J. Pharmacol. 112, 37–42. Thollon, C., Bedut, S., Villeneuve, N., Cogé, F., Piffard, L., Guillaumin, J. P., Brunel-Jacquemin, C., Chomarat, P., Boutin, J. A., Peglion, J. L., et al. (2007). Use-dependent inhibition of hHCN4 by ivabradine and relationship with reduction in pacemaker activity. Br. J. Pharmacol. 150, 37–46. Thomas, J. K., Wiseman, S., Giesy, J. P. and Janz, D. M. (2013). Effects of chronic dietary selenomethionine exposure on repeat swimming performance, aerobic metabolism and methionine catabolism in adult zebrafish (Danio rerio). Aquat. Toxicol. 130–131, 112–122. Tiitu, V. and Vornanen, M. (2002a). Cold adaptation suppresses the contractility of both atrial and ventricular muscle of the crucian carp heart. J. Fish Biol. 59, 141–156. Tiitu, V. and Vornanen, M. (2002b). Regulation of cardiac contractility in a cold stenothermal fish, the burbot Lota lota L. J. Exp. Biol. 205, 1597–606. Topic Popovic, N., Strunjak-Perovic, I., Coz-Rakovac, R., Barisic, J., Jadan, M., Persin Berakovic, A. and Sauerborn Klobucar, R. (2012). Tricaine methane-sulfonate (MS-222) application in fish anaesthesia. J. Appl. Ichthyol. 28, 553–564. Trexler, J. C., Travis, J. and Trexler, M. (1990). Phenotypic plasticity in the sailfin molly, Poecilia latipinna (Pisces: Poeciliidae). II. Laboratory experiment. Evolution (N. Y). 44, 157–167. Tuladhar, R., Yeu, Y., Tyler Piazza, J., Tan, Z., Rene Clemenceau, J., Wu, X., Barrett, Q., Herbert, J., Mathews, D. H., Kim, J., et al. (2019). CRISPR-Cas9-based mutagenesis frequently provokes on-target mRNA misregulation. Nat. Commun. 10, 4056.  223 Urven, L. E., Yabe, T. and Pelegri, F. (2006). A role for non-muscle myosin II function in furrow maturation in the early zebrafish  embryo. J. Cell Sci. 119, 4342–4352. Van Bogaert, P. P. and Pittoors, F. (2003). Use-dependent blockade of cardiac pacemaker current (If) by cilobradine and zatebradine. Eur. J. Pharmacol. 478, 161–171. Van de Peer, Y., Taylor, J. S. and Meyer, A. (2003). Are all fishes ancient polyploids? J. Struct. Funct. Genomics 3, 65–73. Várkuti, B. H., Képiró, M., Horváth, I. Á., Végner, L., Ráti, S., Zsigmond, Á., Hegyi, G., Lenkei, Z., Varga, M. and Málnási-Csizmadia, A. (2016). A highly soluble, non-phototoxic, non-fluorescent blebbistatin derivative. Sci. Rep. 6, 26141. Verhille, C., Anttila, K. and Farrell, A. P. (2013). A heart to heart on temperature: Impaired temperature tolerance of triploid rainbow trout (Oncorhynchus mykiss) due to early onset of cardiac arrhythmia. Comp. Biochem. Physiol. - A Mol. Integr. Physiol. 164, 653–657. Verkerk, A. O. and Remme, C. A. (2012). Zebrafish: A novel research tool for cardiac (patho)electrophysiology and ion channel disorders. Front. Physiol. 3,. Verkerk, A. O. and Wilders, R. (2015). Pacemaker activity of the human sinoatrial node: An update on the effects of mutations in hcn4 on the hyperpolarization-activated current. Int. J. Mol. Sci. 16, 3071–3094. Vinogradova, T. M., Zhou, Y. Y., Maltsev, V., Lyashkov, A., Stern, M. and Lakatta, E. G. (2004). Rhythmic Ryanodine Receptor Ca2+ Releases during Diastolic Depolarization of Sinoatrial Pacemaker Cells Do Not Require Membrane Depolarization. Circ. Res. 94, 802–809. von der Heyde, B., Emmanouilidou, A., Mazzaferro, E., Vicenzi, S., Höijer, I., Klingström, T., Jumaa, S., Dethlefsen, O., Snieder, H., de Geus, E., et al. (2020). Translating GWAS- 224 identified loci for cardiac rhythm and rate using an in vivo image- and CRISPR/Cas9-based approach. Sci. Rep. 10, 11831. Vornanen, M. (1997). Sarcolemmal Ca influx through L-type Ca channels in ventricular myocytes of a teleost fish. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 272, 1432–1440. Vornanen, M. (1998). L-Type Ca2 + current in fish cardiac myocytes: effects of thermal acclimation and β-adrenergic stimulation. Solutions 547, 533–547. Vornanen, M. (2016). The temperature dependence of electrical excitability in fish hearts. J. Exp. Biol. 219, 1941–1952. Vornanen, M. (2017). 3 - Electrical Excitability of the Fish Heart and Its Autonomic Regulation. In The Cardiovascular System (ed. Gamperl, A. K.), Gillis, T. E.), Farrell, A. P.), and Brauner, C. J. B. T.-F. P.), pp. 99–153. Academic Press. Vornanen, M. (2020). Feeling the heat: source-sink mismatch as a mechanism underlying the failure of thermal tolerance. J. Exp. Biol. 223, 225680. Vornanen, M. and Hassinen, M. (2016). Zebrafish heart as a model for human cardiac electrophysiology. Channels 10, 101–110. Vornanen, M., Matikainen, N. and Vornanen, M. (1992). Effect of Season and Temperature Acclimation on the Function of Crucian Carp (Carassius Carassius) Heart. J. Exp. Biol. 167, 203–220. Vornanen, M., Shiels, H. A. and Farrell, A. P. (2002a). Plasticity of excitation – contraction coupling in fish cardiac. Comp. Biochem. Physiol. - part A 132, 827–846. Vornanen, M., Ryökkynen, A. and Nurmi, A. (2002b). Temperature-dependent expression of sarcolemmal K(+) currents in rainbow trout atrial and ventricular myocytes. Am. J. Physiol. Regul. Integr. Comp. Physiol. 282, 1191–1199.  225 Vornanen, M., Hassinen, M., Koskinen, H. and Krasnov, A. (2005). Steady-state effects of temperature acclimation on the transcriptome of the rainbow trout heart. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 289, 1177–1184. Vornanen, M., Hälinen, M. and Haverinen, J. (2010). Sinoatrial tissue of crucian carp heart has only negative contractile responses to autonomic agonists. BMC Physiol. 10,. Vornanen, M., Haverinen, J. and Egginton, S. (2014). Acute heat tolerance of cardiac excitation in the brown trout (Salmo trutta fario). J. Exp. Biol. 217, 299–309. Vornanen, M., Haverinen, J. and Hassinen, M. (2018). Excitation and Excitation-Contraction Coupling of the Zebrafish Heart: Implications for the Zebrafish Model in Drug Screening. In In Recent Advances in Zebrafish Researches, (ed. Bozkurt, Y.), p. 38. Intechopen. Wahl-Schott, C., Fenske, S. and Biel, M. (2014). HCN channels : new roles in sinoatrial node function. Curr. Opin. Pharmacol. 15, 83–90. Wang, X., Chong, M., Wang, X., Wang, H., Zhang, J., Xu, H., Zhang, J. and Liu, D. (2015). Block the function of nonmuscle myosin II by blebbistatin induces zebrafish embryo cardia bifida. Vitr. Cell. Dev. Biol. - Anim. 51, 211–217. Warren, K. S., Baker, K. and Fishman, M. C. (2001). The slow mo mutation reduces pacemaker current and heart rate in adult zebrafish. Am. J. Physiol. Heart Circ. Physiol. 281, 1711–1719. Watanabe, Y., Iwamoto, T., Matsuoka, I., Ohkubo, S., Ono, T., Watano, T., Shigekawa, M. and Kimura, J. (2001). Inhibitory effect of 2,3-butanedione monoxime (BDM) on Na+/Ca2+ exchange current in guinea-pig cardiac ventricular myocytes. Br. J. Pharmacol. 132, 1317–1325. Watters, J. V., Lema, S. C. and Nevitt, G. A. (2003). Phenotype management: A new approach  226 to habitat restoration. Biol. Conserv. 112, 435–445. Whitaker, G. M., Angoli, D., Nazzari, H., Shigemoto, R. and Accili, E. A. (2007). HCN2 and HCN4 isoforms self-assemble and co-assemble with equal preference to form functional pacemaker channels. J. Biol. Chem. 282, 22900–22909. Wilson, C. M. and Farrell, A. P. (2013). Pharmacological characterization of the heartbeat in an extant vertebrate ancestor, the Pacific hagfish, Eptatretus stoutii. Comp. Biochem. Physiol. - A Mol. Integr. Physiol. 164, 258–263. Wilson, C. M., Stecyk, J. A. W., Couturier, C. S., Nilsson, G. E. and Farrell, A. P. (2013). Phylogeny and effects of anoxia on hyperpolarization- activated cyclic nucleotidegated channel gene expression in the heart of a primitive chordate, the pacific hagfish (Eptatretus stoutii). J. Exp. Biol. 216, 4462–4472. Wilson, C. M., Roa, J. N., Cox, G. K., Tresguerres, M. and Farrell, A. P. (2016). Introducing a novel mechanism to control heart rate in the ancestral Pacific hagfish. J. Exp. Biol. 219, 3227–3236. Xie, Y., Ottolia, M., John, S. A., Chen, J. N. and Philipson, K. D. (2008). Conformational changes of a Ca2+-binding domain of the Na+/Ca2+ exchanger monitored by FRET in transgenic zebrafish heart. Am. J. Physiol. - Cell Physiol. 295, 388–393. Yamauchi, A. and Burnstock, G. G. (1968). An electron microscopic study on the innervation of the trout heart. J. Comp. Neurol. 132, 567–587. Yampolsky, P., Koenen, M., Mosqueira, M., Geschwill, P., Nauck, S., Witzenberger, M., Seyler, C., Fink, T., Kruska, M., Bruehl, C., et al. (2019). Augmentation of myocardial If dysregulates calcium homeostasis and causes adverse cardiac remodeling. Nat. Commun. 10, 2395.  227 Yang, J., Hartjes, K. A., Nelson, T. J. and Xu, X. (2014). Cessation of contraction induces cardiomyocyte remodeling during zebrafish cardiogenesis. Am. J. Physiol. - Hear. Circ. Physiol. 306, 382–395. Yaniv, Y., Lakatta, E. G. and Maltsev, V. A. (2015). From two competing oscillators to one coupled-clock pacemaker cell system. Front. Physiol. 6,. Zeng, J., Laurita, K. R., Rosenbaum, D. S. and Rudy, Y. (1995). Two components of the delayed rectifier K+ current in ventricular myocytes of the guinea pig type. Circ. Res. 77, 140–152. Zhang, Q., Huang, A., Lin, Y. C. and Yu, H. G. (2009). Associated changes in HCN2 and HCN4 transcripts and If pacemaker current in myocytes. Biochim. Biophys. Acta - Biomembr. 1788, 1138–1147. Zhang, P.-C., Llach, A., Sheng, X. Y., Hove-Madsen, L. and Tibbits, G. F. (2011). Calcium handling in zebrafish ventricular myocytes. AJP Regul. Integr. Comp. Physiol. 300, R56–R66. Zhang, X. H., Wei, H., Šarić, T., Hescheler, J., Cleemann, L. and Morad, M. (2015). Regionally diverse mitochondrial calcium signaling regulates spontaneous pacing in developing cardiomyocytes. Physiol. Behav. 57, 321–336. Zhang, Y., Gilbert, M. J. H. and Farrell, A. P. (2020). Measuring maximum oxygen uptake with an incremental swimming test and by chasing rainbow trout to exhaustion inside a respirometry chamber yields the same results. J. Fish Biol. 28–38. Zicha, S., Fernández-Velasco, M., Lonardo, G., L’Heureux, N. and Nattel, S. (2005). Sinus node dysfunction and hyperpolarization-activated (HCN) channel subunit remodeling in a canine heart failure model. Cardiovasc. Res. 66, 472–481.  228 Zon, L. I. and Peterson, R. T. (2005). In vivo drug discovery in the zebrafish. Nat. Rev. Drug Discov. 4, 35–44.        229 Appendix   Appendix 2.1. ECG recordings from a 23°C-acclimated fish incrementally heated from 18°C to 34.4°C. (A) The ECG recording at 18°C (B) the ECG recording at 34.4°C. Red arrows indicate missing QRS complexes.        230  Appendix 2.2. Heart rate response to a second injection of either (A) zatebradine (4 μg g-1) or (B) ryanodine (50 ng g-1) in combination with thapsigargin (1.3 μg g-1) . 231 Appendix 3.1. Primers used for sgRNA synthesis. T7 promotor is in red, target sequence is in black and the tracerRNA is underlined. CRISPRsg reverse universal primer. Indicated in bold is the annealing sequence.  sgRNAs Guide sequence (5’-3’)  sgRNA 1  TAATACGACTCACTATAGGGCAGAAAGCGTCCATCATGTTTTAGAGC sgRNA 2 TAATACGACTCACTATAGAAGCCTTTACCGACCAGCGGTTTTAGAGC sgRNA 3 TAATACGACTCACTATAGGCATCCCGGATCAGCAGGCGTTTTAGAGC CRISPRsg universal  reverse primer AAAGCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGCCTTATTTTAACTTGCTATTTCTAGCTCTAAAAC  232  Appendix 3.2. Computational model of wild type and mutant truncated form of hcn4 as mediated by CRISPR-Cas9 insertion of a premature stop codon in position 433-435 (TAA). Swiss model web tool was used to design the models using known HCN models as a template. (A) Zebrafish Hcn4 modeled from the protein sequence (B) Global Quality Estimate of the model (C) local quality estimate of the model (>60 indicates low model confidence). (D) Zebrafish hcn4-/- protein with premature stop codon located at the 432 bp. (E) Global Quality Estimate of the mutant model (F) local quality estimate of the model (>60 indicates low model confidence). Blue indicates high confidence in the model, red indicates low confidence.  233  Appendix 3.3.  (A)  Alignment of hcn4 coding sequence with CRISPR-Cas9 mediated premature stop codon (TAA) indicated by a red arrow and a dashed red line indicating the position of the premature stop codon in the translated protein sequence and associated functional domains. (B) Protein domains of HCN4 as determined by Ensemble. (C) Protein domains, families and functional sites identified by Prosite.    234  Appendix 3.4. The weight and CTmax of individual wild-type (n=47) and hcn4 (n=43) knockout fish.                 0.0 0.2 0.4 0.6 0.83638404244Weight (g)CTMax (°C)Wild-typeHcn4-/-Y = 4.11X+38.02R2 = 0.0941Y = 0.4006X + 40.35R2 = 0.0036P = 0.084 235 Appendix 3.5. The percent similarity (%) between the zebrafish and human cDNA sequences and protein sequences for different HCN isoforms. * signifies no significant difference in BLAST with Megablast (highly similar sequences), the percentage shown was found with blastn (somewhat similar sequences).     Zebrafish (Danio rerio) Human (Homo sapiens)    hcn4  hcn4l  Hcn4  ENSG00000138622  Hcn3  ENSG00000143630  cDNA sequence percent identity (%) Zebrafish hcn4 ENSDARG00000061685   81.71 77.74 Zebrafish hcn4l ENSDARG00000074419 83.37  80.16 75.90* Zebrafish hcn3 ENSDARG00000027192 75.33 70.91 76.26 76.09    Protein sequence percent identity (%)  Zebrafish Hcn4    74.71 76.99 Zebrafish Hcn4l 66.87  84.64 77.93 Zebrafish Hcn3 79.86 78.03 68.71 76.29    236 Appendix 3.6. Sequence similarity between the protein sequences of zebrafish Hcn4 and Hcn4l. Identical nucleotides are linked by a vertical line, highly similar nucleotides are represented by 2 dots, similar nucleotides are represented by a single dot and different dissimilar nucleotides are represented by a blank space between the nucleotides. Obtained using EMBOSS global alignment using the Needleman-Wunsch algorithm.  237  Appendix 4.1. The effect of blebbistatin on (A) action potential duration 50% and (B) action potential duration 80% recorded in reduced heart preparations at room temperature (20°C-22.9°C). Data points represent individuals. Data are presented with the SD. Statistical significance was determined using a t-test with P ≤ 0.05 as the level of significance.        238  Appendix 4.2.  The effect of blebbistatin on (A) overshoot potential (B) AP amplitude (C) Ap threshold potential and (D) maximum hyperpolarization potential of control and blebbistatin treated cells recorded in reduced heart preparations at room temperature (20°C-22.9°C). Data points represent individuals. Data are presented with the SD. Statistical significance was determined using a t-test with P ≤ 0.05 as the level of significance.     239   Appendix 4.3. The effect of blebbistatin and temperature on the action potential threshold potential and the overshoot potential. (A) A linear regression analysis of AP threshold potential with increasing temperature of control and blebbistatin treated cells. (B) A pairwise comparison of AP threshold potential of control and blebbistatin treated cells at different temperatures intervals. (C) A linear regression analysis of overshoot potential with increasing temperature of control and blebbistatin treated cells. (D) A pairwise comparison of overshoot potential of control and blebbistatin treated cells at different temperatures intervals. Data points represent individuals. Data are presented with the SD. Statistical significance of pairwise comparisons was determined using a t-test with P ≤ 0.05 as the level of significance.  240   Appendix 4.4. The effect of blebbistatin and temperature on AP amplitude and the maximum hyperpolarization potential. (A) A linear regression analysis of the AP amplitude with increasing temperature of control and blebbistatin treated cells. (B) A pairwise comparison of AP amplitude of control and blebbistatin treated cells at different temperatures intervals. (C) A linear regression analysis of the maximum hyperpolarization potential with increasing temperature of control and blebbistatin treated cells. (D) A pairwise comparison of the maximum hyperpolarization potential of control and blebbistatin treated cells at different temperatures intervals. Data points represent individuals. Data are presented with the SD. Statistical significance of pairwise comparisons was determined using a t-test with P ≤ 0.05 as the level of significance.     241     Appendix 4.5. A linear regression analysis of the action potential duration 50% and 80%. (A) A linear regression analysis of AP duration at 50% repolarization with increasing temperature of control and blebbistatin treated cells. (B) A pairwise comparison of AP duration at 50% repolarization of control and blebbistatin treated cells at different temperatures intervals. (C) A linear regression analysis of AP duration at 80% repolarization with increasing temperature of control and blebbistatin treated cells. (D) A pairwise comparison of AP duration at 80% repolarization of control and blebbistatin treated cells at different temperatures intervals. Data points represent individuals. Data are presented with the SD. Statistical significance of pairwise comparisons was determined using a t-test with P ≤ 0.05 as the level of significance.   242    Appendix 4.6. Principal component analysis of voltage-dependent action potential parameters of control (blue circle) and blebbistatin-treated (green diamond) heart preparations recorded at room temperature (20°C-23°C). Each point represents APs recorded from a single pacemaker cell.        243  Appendix 4.7. Principal component analysis of time-dependent action potential parameters of control (blue circle) and blebbistatin-treated (green diamond) heart preparations recorded at room temperature (20°C-23°C). Each point represents APs recorded from a single pacemaker cell.    244  Appendix 4.8. Sample power plot of required effect size as performed by G*Power 3.1.  0.6 0.65 0.7 0.75 0.8 0.85 0.9 0.950100200300400500600Total sample size = 0.291421Effect size dt tests -  Means: Difference between two independent means (two groups)Tail(s) = Two. Allocat ion rat io N2/N1 = 1. α err prob = 0.05. Effect size d = 0.291421Power (1-β err prob)

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