CBA-09746; No of Pages 9 Comparative Biochemistry and Physiology, Part A xxx (2014) xxx–xxx

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Review

The beat goes on: Cardiac pacemaking in extreme conditions☆ Christopher M. Wilson a,⁎, Georgina K. Cox a, Anthony P. Farrell a,b a b

Department of Zoology, University of British Columbia, Vancouver, British Columbia V6T 1Z4, Canada Faculty of Land & Food Systems, University of British Columbia, Vancouver, British Columbia V6T 1Z4, Canada

a r t i c l e

i n f o

Article history: Received 29 June 2014 Received in revised form 17 August 2014 Accepted 20 August 2014 Available online xxxx Keywords: Heart rate HCN Hagfish Turtle Carp Salmon Anoxia Hypoxia Temperature

a b s t r a c t In order for an animal to survive, the heart beat must go on in all environmental conditions, or at least restart its beat. This review is about maintaining a rhythmic heartbeat under the extreme conditions of anoxia (or very severe hypoxia) and high temperatures. It starts by considering the primitive versions of the protein channels that are responsible for initiating the heartbeat, HCN channels, divulging recent findings from the ancestral craniate, the Pacific hagfish (Eptatretus stoutii). It then explores how a heartbeat can maintain a rhythm, albeit slower, for hours without any oxygen, and sometimes without autonomic innervation. It closes with a discussion of recent work on fishes, where the cardiac rhythm can become arrhythmic when a fish experiences extreme heat. © 2014 Elsevier Inc. All rights reserved.

Contents 1. Introduction . . . . . . . . . . . . . 2. Pacemaker activity in craniates . . . . . 3. HCN channel isoforms and their evolution 4. Heart rate control in anoxia . . . . . . 5. Fish at supra-optimal water temperatures Acknowledgments . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . .

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1. Introduction Heart rate in craniate animals (chordates with a well-defined head) varies considerably in response to the varying demands placed on the Abbreviations: SL, sarcolemma; IKr, delayed rectifier K+ current; HCN, hyperpolarizationactivated cyclic nucleotide-gated; If, funny current; SR, sarcoplasmic reticulum; RyRs, ryanodine receptors; NCX, sodium/calcium exchanger channel; PO, power output; CTmax, upper thermal tolerance; Tarr, arrhythmia triggering temperature. ☆ Fuelling the Fire of Life: Papers from a symposium celebrating the career of Dr. William K. Milsom, held in Edinburgh, Scotland 24th–27th June, 2014. The symposium was sponsored by the Society of Experimental Biology (SEB). ⁎ Corresponding author. Tel.: +1 778 968 6994. E-mail address: [email protected] (C.M. Wilson).

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heart to deliver blood to tissues, e.g., during times of exercise, stress, digestion and altered body temperature. In extant animals as diverse as the most primitive craniates (hagfishes), through fishes, amphibians and reptiles, to birds and mammals, cardiac output typically can vary at least 3-fold and such changes are often attributed to an altered heart rate (e.g., Shin et al., 1995; Eliason et al., 2013). Here we consider the control of heart rate under two extreme conditions. First, we consider environmental anoxia (or very severe hypoxia), which is when the primary role of the circulatory system to transport O2 and CO2 between tissues and the respiratory organ is lost, or becomes minimal, due to ambient O2 being temporarily unavailable. As O2 levels in arterial blood fall, the tissue's capacity to generate ATP using oxidative phosphorylation becomes progressively limited and

http://dx.doi.org/10.1016/j.cbpa.2014.08.014 1095-6433/© 2014 Elsevier Inc. All rights reserved.

Please cite this article as: Wilson, C.M., et al., The beat goes on: Cardiac pacemaking in extreme conditions, Comp. Biochem. Physiol., A (2014), http://dx.doi.org/10.1016/j.cbpa.2014.08.014

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C.M. Wilson et al. / Comparative Biochemistry and Physiology, Part A xxx (2014) xxx–xxx

even the heart becomes challenged with generating sufficient ATP through glycolytic pathways. Moreover, an anoxic brain may constrain autonomic nervous control (Nilsson, 2001; Nilsson and Lutz, 2004), including that of the cardiac pacemaker. The second environmental condition we consider is supra-optimal temperature in fishes, which is when maximum cardiac performance is in decline due to excessive warming. Cardiac performance is tightly related to heart rate. In order to understand how cardiac performance is altered under these two extreme environments, extrinsic and intrinsic control of pacemaker activity has been explored to understand the modulation of the rhythm of the heartbeat. We start by considering pacemaker activity in an ancestral craniate heart, that of the hagfish, which has no autonomic innervation. As such, this naturally aneural heart offers a unique approach to the study of the initiation and regulation of the craniate heartbeat. 2. Pacemaker activity in craniates

Hyperpolarizaon

Repolarizaon

Depolarizaon

The agnathan branchial heart shares many functional and anatomical characteristics with vertebrate hearts. Like other craniates, the hagfish heart is multi-chambered and contains cardiac muscle cells (cardiomyocytes) that intrinsically respond to increased stretch (increased cardiac filling) by increasing contractile force (increased cardiac stroke volume), which is termed the Frank–Starling mechanism. Cardiac contractions are also myogenic (initiated within the heart itself). Yet, despite the lack of cardiac innervation (Axelsson et al., 1990; Bloom et al., 1963; Farrell, 2007; Ota and Kuratani, 2007), heart rate in the Pacific hagfish (Eptatretus stoutii) can still vary considerably (e.g., fourfold on recovery from anoxia; Cox et al., 2010), albeit much more slowly (it takes approximately 2 h for the changes on recovery from anoxia) than with direct autonomic neural control. Recent discoveries on hagfish heartbeat initiation and cardiac control in the context of prolonged anoxia have provided novel and interesting insights into how the earliest form of the craniate heart is operated. Prior to highlighting these novel and recent discoveries for pacemaking of the hagfish heart, we first provide a brief introduction of the initiation of the heartbeat in vertebrate pacemaker tissues because the initiation of the vertebrate heartbeat is well studied in non-piscine models. Specialized pacemaker cells (termed the sinoatrial node and located in the sinus venosus, or at the sinoatrial border in more derived vertebrates) initiate the heartbeat. These cells self-generate rhythmic action potentials (Fig. 1) that subsequently trigger action potentials in nonpacing contractile cardiomyocytes in a rhythmic manner. Pacemaker

Fig. 1. The major electrical stages of a cardiac pacemaker cell action potential. Upon the reaching of T-type Ca2+ threshold potential, an influx of Ca2+ depolarizes the cell. Efflux of potassium repolarizes the cell, and even hyperpolarizes it, activating the pacemaker potential. This steady influx of Na+ and K+ gently depolarizes the cell back to the threshold for T-type Ca2+. See text for more details.

cells have been identified in all major craniate groups from fish to mammals, including hagfish (Jensen, 1965). In fact, pacemaker potentials with shapes similar to those found in mammalian hearts have been recorded in hagfish hearts using sharp-microelectrodes (Jensen, 1965; Arlock, 1975; Vornanen et al., 2002). However, hagfish cardiomyocytes are considerably depolarized at rest (−41 and −48 mV in the atrium and ventricle, respectively) compared to teleost fishes and mammals (−65 mV and −75 mV in the atrium and ventricle, respectively). The action potential of a pacemaker cell differs from that of a contractile cardiomyocyte by having a pacemaker potential, an unstable resting membrane potential that, over time, depolarizes until it reaches the threshold for the action potential upstroke. Historically, two models have been proposed to explain the pacemaker potential in vertebrate pacemaker cells: the “membrane clock hypothesis” and the “calcium clock hypothesis”. The “membrane clock hypothesis” proposes that all ion currents affecting pacemaker rhythmicity are located in the sarcolemma (SL) of the pacemaker cell. Following the spike of the action potential, an outward delayed rectifier K+ current (IKr) causes the SL membrane to become highly hyperpolarized (negatively charged compared to the threshold of the Ca2 + channels), which activates the hyperpolarization-activated cyclic-nucleotide gated (HCN) channels located in the SL membrane. This then allows a slow influx of Na+ and K+ (termed the funny current, If), which slowly depolarizes the cell membrane (Brown et al., 1979a; Brown and DiFrancesco, 1980; DiFrancesco and Ojeda, 1980; Yanagihara and Irisawa, 1980; Doerr et al., 1989; Baker et al., 1997; DiFrancesco, 2010). This gradual depolarization (the pacemaker potential) occurs until the threshold potential to activate T-type Ca2+ channels is reached. Rapid entry of Ca2+ into the pacemaker cells via these T-type Ca2+ channels creates the upstroke of the action potential and the depolarization again activates IKr, restarting the cycle (Hagiwara et al., 1988; Doerr et al., 1989; Zhou and Lipsius, 1994). The “calcium clock hypothesis” proposes that the ion currents that affect pacemaker rhythmicity are located primarily in the sarcoplasmic reticulum (SR) membrane as well as in the SL (Vinogradova et al., 2004). With this model, depolarization of the cell membrane during the pacemaker potential is initiated by spontaneous release of Ca2+ from the SR stores into the cytoplasm (termed Ca2+ sparks that can be visualized using calcium imaging techniques) through ryanodine receptors (RyRs) in the SR membrane (Rubenstein and Lipsius, 1989; Huser et al., 2010; Lyashkov et al., 2007; Maltsev and Lakatta, 2007). As Ca2+ builds up in the cytoplasm from these Ca2+ sparks, the sodium/calcium exchanger channel (NCX) removes one Ca2+ ion in exchange for 3 Na+ ions, with the result of each exchange being the net movement of 1 positive ion into the cell (Shigekawa and Iwamoto, 2001) and a slow depolarization of the membrane potential. The coupling of SR Ca2+ release with translocation of Ca2+ out of the pacemaker cell via NCX is aided by the co-localization of NCX channels in the SL and RYRs in the SR. Recently, Monfredi et al. (2013) bought the membrane and calcium clock models for vertebrate pacemaker cells together by proposing that the depolarizations caused by both If and SR-Ca2+ release-stimulated NCX reinforce each other to bring about the pacemaker potential. A simple pharmacological approach has been used to test these membrane and calcium clock models in hagfish hearts by applying selective pharmacological channel blockers (Wilson and Farrell, 2013). When isolated hagfish hearts are placed into physiological saline held at 10 °C, they continue to beat rhythmically at about 21 bpm for over 24 h. When the ventricle is separated from the atrium, each chamber continues to beat rhythmically, the atrium at the same rate, but the ventricle at 41% (8 bpm) of the atrial rate (Wilson and Farrell, 2013). HCN channels in the SL can be blocked by zatebradine and ZD7222 and such application would provide support for the membrane clock hypothesis if it reduced heart rate. When 0.05 mM of zatebradine, the HCN blocker, is applied to the whole heart, heart rate is reduced by 61% (8.5 bpm; Wilson and Farrell, 2013). With 5 mM zatebradine, spontaneous atrial contractions ceased completely, while ventricular

Please cite this article as: Wilson, C.M., et al., The beat goes on: Cardiac pacemaking in extreme conditions, Comp. Biochem. Physiol., A (2014), http://dx.doi.org/10.1016/j.cbpa.2014.08.014

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contractions continued at a further reduced rate of 3 bpm, possibly because the atrium was no longer pumping the antagonist into the ventricle (Wilson and Farrell, 2013). Conversely, blocking the ryanodine receptor with ryanodine, which has been shown to reduce the pacemaker potential in feline pacemaker cells (Rubenstein and Lipsius, 1989; Huser et al., 2000; Lyashkov et al., 2007; Maltsev and Lakatta, 2007), would provide support for the calcium clock hypothesis. When ryanodine was applied to the whole heart (from 0.1 μM to 100 μM) neither the atrial nor ventricular contraction rate changed (Wilson and Farrell, 2013). These results for zatebradine and ryanodine applications lend strong support to membrane clock hypothesis rather than the calcium clock. Thus, an HCN-mediated If may be the ancestral pacemaking mechanism for the vertebrate-type heart. Work conducted in vivo with teleost fishes shows that infusion of HCN agonists slows resting and maximal heart rates by 11% and 33%, respectively (Keen and Gamperl, 2012), which is much less of an effect than in hagfish and more in line with the quantitative effect seen in mammalian studies. Yet, a mutated HCN channel results in an embryonically lethal bradycardia in the slow mo zebrafish (Danio rerio) mutation (Baker et al., 1997). While these studies lend support for a membrane clock hypothesis in modern teleosts, parallel and definitive studies with ryanodine are lacking. Instead, while investigating Ca2+-cycling in rainbow trout (Oncorhynchus mykiss) hearts, Haverinen and Vornanen (2007) found that ryanodine and thapsigargin (a SR-Ca2 + re-uptake antagonist) decreased heart rate only at a supra-optimal temperature (18 °C). At 4 °C, when heart rate was three times slower than at 18 °C, neither antagonist altered heart rate. Taken together with reports of more active fishes having a greater reliance on SR-Ca2+ cycling (Vornanen, 1989; Shiels et al., 1998, 1999; Shiels and Farrell, 2000), one interpretation of these results is that the calcium clock may become important in supporting higher heart rates in more active fishes, such as rainbow trout. There is a need of further research in this area. 3. HCN channel isoforms and their evolution If is carried across the SL membrane by HCN channels, which are part of the K+ channel super family (Craven and Zagotta, 2006). In mammals, HCN channels consist of four subunits, each of which can be one of four isoforms, named HCN1-4, joined together as homotetramers or heterotetramer (Whitaker et al., 2007; Chow et al., 2011). Mammalian hearts predominantly express HCN4 in the sinoatrial node, with HCN2 dominating in the ventricle (Shi et al., 1999; Moosmang et al., 2001; Marionneau et al., 2005). In addition, early in ontogeny, there is a switch from HCN3 to HCN4 in the embryonic mouse heart, which is timed to occur as cardiac contractions begin (Qu et al., 2008; Schweizer et al., 2009). Two studies have considered the evolution of HCN channels by studying the sequences and expression of HCN isoforms in urochordate, Ciona (Jackson et al., 2007) and hagfish, E. stoutii (Wilson et al., 2013) hearts. After discovering 3 urochordate specific isoforms (HCNa–c), whose sequences most closely resembled vertebrate HCN3 and likely resulted from lineage-specific duplication events, Jackson et al. (2007) proposed that the four mammalian HCN genes evolved from a single ancestral HCN gene most similar to HCN3. Work with the hagfish heart revealed expression HCN2, HCN3 and HCN4 mRNA, with the HCN3 isoform dominating the mRNA expression in both atrial and ventricular tissues, followed by HCN4 (Wilson et al., 2013). While studies on these proteins are lacking, this finding supports the contention of Jackson et al. (2007; Fig. 2) that HCN3 may be the ancestral HCN isoform. Moreover, hagfishes have mRNA encoding a total of three HCN3 isoforms (HCN3a–c), with HCN3b existing as two paralogs, as well as two HCN2 isoforms (HCN2a and b). Thus, the hagfish cardiac mRNA encoded a total of six HCN genes (Wilson et al., 2013). Although the hagfish data represents mRNA rather than protein expression, it has been proposed that a switch from HCN3 to HCN4 dominance took place at some point in the vertebrate taxa (Wilson et al., 2013), similar to the

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embryonic switch that occurs in mice as heart contractions begin (Qu et al., 2008). This evolutionary switch is likely to have occurred prior to the emergence of the tetrapods, since the turtle, Trachemys scripta, shows HCN4 dominance in the sinus venosus (Jonathan Stecyk, personal communication), though the situation in teleost and other fishes is unknown. To date, protein and mRNA expression studies on nonmammalian vertebrate HCN channels are extremely limited to test this speculation. Instead, genomic sequences have been studied in representatives for the avian, teleost fish and mammalian lineages (Jackson et al., 2007). Interestingly, some teleost fishes have extra copies of some HCN isoforms, for example, the green puffer has multiple copies of HCN4, indicating perhaps that, like in the urochordates and hagfish, HCN duplication events are fairly common (Jackson et al., 2007; Fig. 2). 4. Heart rate control in anoxia Across the vertebrate phylum, very few species can endure O2 deprivation for more than a few minutes, in part because the heart is an aerobic organ and a myocardial O2 supply must be maintained for a lifesustaining, rhythmic heartbeat. The exceptions include the known champions of cardiac anoxia tolerance: hagfishes such as E. stoutii, freshwater turtles such as T. scripta and the crucian carp, Carassius carassius, each of which has been shown to reach new steady states for cardiac activity during prolonged anoxia (Cox et al., 2010; Stecyk et al., 2004a, 2009a). Therefore, during anoxia these animals maintain a rhythmic heartbeat to sustain life in a state of tolerance until ambient O2 is again available and normal activities may resume. The general question posed here, is what happens to heart rate during anoxia when the animal faces a triple challenge: the primary role of the circulatory system to transport gases is relaxed; cardiac contractile activity is fuelled by anaerobically generated ATP; and the anoxic brain may be unable to regulate heart rate. Pacific hagfishes live and feed in hypoxic and anoxic environments. In the laboratory, they are capable of surviving and recovering from up to 36 h of complete anoxia at 10 °C (Cox et al., 2010, 2011; Martini, 1998), but not much longer at this temperature. Anoxia tolerance of hagfish as a whole is attributed to their extremely low routine metabolic rate in combination with fermentable glycogen stores, high blood volume and by a suppression of routine metabolic rate, which appears to be activated after about 6 h of anoxia (Cox et al., 2011; Farrell and Stecyk, 2007). Over a period of 36 h of anoxia, it has been estimated that E. stoutii can decrease routine metabolic rate by at least 50% (Cox et al., 2010, 2011). Accompanying the suppression of routine metabolic rate, heart rate similarly decreases by ~ 50%, from 10 bpm to 5 bpm (Fig. 3; Table 1). Without autonomic innervation, how can the hagfish slow heart rate in anoxia? Shifts in HCN expression can be potentially eliminated because mRNA expression was unchanged in atrial tissue even though HCN3 mRNA expression fell in ventricular tissue after 24 h of anoxia (Wilson et al., 2013). However, this conclusion is made in the absence of protein expression data, and it is possible that changes in protein expression or post-translational expression could still play an important role in controlling heart rate of anoxic hagfish. What is clear, however, is that modulation of HCN channels by cAMP plays a major role in the adjustment of heart rate in the aneural hagfish heart due to intracellular cAMP interactions with HCN channels. The finding that adrenergic antagonists injected in normoxic hagfish decrease heart rate provides the first line of evidence for this mode of heart rate modulation (e.g., Axelsson et al., 1990). This is because catecholamines (adrenaline and noradrenaline), which stimulate βadrenoceptors, are well-established regulators of cytosolic cAMP in cardiomyocytes. These β-adrenoceptors in turn activate membranebound adenylyl cyclase that converts ATP to cAMP, which can then alleviate the inhibition of HCN by the CNBD (Brown et al., 1979a,b; Brown and DiFrancesco, 1980; Kaumann et al., 1982; Waelbroeck et al., 1983; DiFrancesco, 1985, 2010; Ikezono et al., 1987; Lohse et al., 2003; Guo et al., 2004). In the mammalian heart If flow through HCN channels is

Please cite this article as: Wilson, C.M., et al., The beat goes on: Cardiac pacemaking in extreme conditions, Comp. Biochem. Physiol., A (2014), http://dx.doi.org/10.1016/j.cbpa.2014.08.014

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C.M. Wilson et al. / Comparative Biochemistry and Physiology, Part A xxx (2014) xxx–xxx HCN2a Hagfish

HCN1 HCN2

HCN2b Hagfish

HCN3 HCN4 Outgroup HCN2b Green puffer HCN2 Human HCN2b Fugu

HCN2 Rat HCN2 Mouse HCN4 Human HCN4 Mouse HCN4 Rat HCN4 Chimpanzee HCN4b Fugu 21 HCN4 Rabbit HCN4b Green puffer 30 HCN4 Hen 25 HCN4 Hagfish HCN3c Hagfish

HCN4a Green puffer HCN1 Rat

HCN4a Fugu

826

HCN2 Hen

HCN1 Opossum

31

HCN1 Human

141

5 3

972

109 288

31

HCN3a Hagfish

342

HCN1 Rabbit 237

HCN3b Hagfish 80 604 121

HCN3 Green puffer

303

82 95 55 130

851

111 118 118 224

HCN1 Chimpanzee

670 988

HCN3 Fugu

430

442

667 930

HCN1 Mouse

644

HCNb Ciona

829

409

HCN1 Hen

329

HCNa Ciona

974

HCN2 Trout

400

HCNc Ciona

HCN1 Green puffer 442

866

HCN1 Trout 681 300

HCN2a Green puffer

ERG Human CNG Drosophila

HCN3 Opossum

158 91 110

HCN3 Rat

HCN2 Killifish

HCN2 Zebrafish

HCN3 Human

0.01

HCN2a Fugu

HCN3 Mouse

HCN3 Dog

HCN3 Cow

Fig. 2. Bootstrapped Neighbor-Joining phylogeny of HCN genes including hagfish HCN isoforms. Hagfish genes were partially cloned and sequenced from hagfish hearts. Drosophila, Ciona and vertebrate sequences downloaded from GenBank and Ensembl. Sequences aligned in ClustalX, edited and translated to amino acids in GeneDoc, phylogeny compiled using Phylip-3.69 and edited using Adobe Illustrator CS5. Tree run with 1000 bootstrap datasets, low bootstrap values likely a consequence of both the short 37 amino acid sequence used and a high sequence identity as seen in Jackson et al. (2007). HCN, hyperpolarization-activated cyclic nucleotide-gated channel, CNG, cyclic nucleotide gated channel, ERG, Ether-á-go-go related gene. Nomenclature used by Jackson et al. (2007) is used with chicken abbreviated to Hen for clarity. Scale bar represents 1% residue substitutions per site. Taken from Wilson et al. (2013).

inhibited by the cyclic nucleotide binding domain (CNBD) attached to the rest of the protein by a C-linker close to the C-terminus (Craven and Zagotta, 2006; Jackson et al., 2007). When cAMP binds to the CNBD, conformational changes of CNBD and C-linker allow the formation of a gating ring between adjacent subunits of the tetramer, reducing the autoinhibition of the CNBD, and increasing If flow (Chow et al., 2011). Thus, a reduction in cytosolic cAMP will reduce If and, subsequently, heart rate. Adrenergic stimulation of the heart and its modulation of heart rate can involve sympathetic nerve endings in cardiac tissues, including the pacemaker region, that release noradrenaline. In addition, the adrenal gland and chromaffin tissue release adrenaline and noradrenaline into

the blood when stimulated by the sympathetic nervous system (Nilsson, 1983). Hagfish hearts are unusual in that they contain dense staining, catecholamine-filled granules visible under electron microscopy (Johnels and Palmgren, 1960; Östlund et al., 1960; Bloom et al., 1961; Jensen, 1961; von Euler and F nge, 1961; Perry et al., 1993), likely a functional equivalent to chromaffin tissue but without innervation. Indeed, high concentrations of adrenaline and noradrenaline have been measured in both the atrium and ventricle of hagfish (von Euler and F nge, 1961; Bloom et al., 1963; Perry et al., 1993; Farrell, 2007). Thus, autocrine stimulation by catecholamines may provide control of heart rate in hagfishes (Farrell, 2007), which would compensate for the low concentration of circulating catecholamines reported in hagfish blood

Please cite this article as: Wilson, C.M., et al., The beat goes on: Cardiac pacemaking in extreme conditions, Comp. Biochem. Physiol., A (2014), http://dx.doi.org/10.1016/j.cbpa.2014.08.014

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Fig. 3. Representative ventral aortic blood flow recordings from a single Pacific hagfish in normoxia, during 36 h of anoxia and after 1.5 h of normoxic recovery. Each trace is for 1 min and heart rate (fH) is reported as beats per minute. Modified from Cox et al. (2010).

even after stimuli known to cause catecholamine release have been applied (Perry et al., 1993). Moreover, there is excellent evidence that routine heart rate is under a strong tonic catecholamine stimulation via β-adrenergic receptors. Heart rate under normoxic conditions is typically reduced by 50% (from 25 bpm to 14 bpm) when the catecholamine effect is blocked with dihydroergotamine in vitro (Fänge and Östlund, 1954), and with sotalol or propranolol in vivo (Axelsson et al., 1990; Forster et al., 1992). Under anoxia, an overlooked problem with respect to tonic adrenergic stimulation of heart rate in hagfish is the O2 dependence of the ratelimiting step in catecholamine production; 3,4-dihydroxyphenylalanine can no longer be produced from tyrosine (Levitt et al., 1965). Thus, it seems likely that catecholamine production and recycling cease during anoxia and, over the long term, cardiac stores normally used for tonic stimulation could become depleted. This problem may explain the bradycardia to around 5 bpm seen in vivo during which anoxia develops so slowly (Cox et al., 2010). Similarly, when hagfish hearts are isolated and made anoxic, heart rate also falls to 5 bpm over a period of 2 h. Furthermore, under anoxic conditions the anoxic heart is no refractory to adrenergic stimulation. Maximally stimulating β-adrenoceptors with the agonist forskolin does increase heart rate to a routine normoxic levels but not beyond (Wilson, 2014), suggesting that the anoxic hagfish heart is similar to the anoxic crucian carp where isoprenaline, a βadrenergic agonist, failed to increase heart rate (Vornanen et al., 2010). The bradycardia seen in anoxia may also involve extrinsic factors associated with anoxia, such as extracellular acidosis and hyperkalemia, both of which reduce heart rate in anoxic, cold-acclimated turtles (Jackson, 2000; Jackson and Ultsch, 1982; Overgaard et al., 2005; Stecyk and Farrell, 2007). Consequently, a hypothesis worth testing is that the decrease in heart rate during prolonged anoxia in hagfish Table 1 Comparison of routine and anoxic heart rate values in 3 anoxia tolerant species. Species

Temp (°C)

Eptatretus stoutii Carassius carassius Trachemys scripta

10 8 5 21

Heart rate (beats min−1) Routine 10.4 16.94 4.96 21.1

± ± ± ±

Anoxia 1.3 1.50 0.38 1.1

4.7 14.24 1.06 10.1

E. stoutii data from Cox et al. (2010). C. carassius data from Stecyk et al. (2004a,b) and personal communication. T. scripta data from Hicks and Farrell (2000b).

± ± ± ±

1 0.76 0.11 0.6

involves a reduction in catecholamine-stimulated, adenylyl cyclase produced cAMP. Also, in need of explanation is the limited ability of adrenergic stimulation to increase heart rate beyond routine normoxic rates. For example, adrenaline addition to hagfish hearts produces a very modest increase in heart rate, and forskolin fails to increase heart rate in isolated hearts above routine (Axelsson et al., 1990; Wilson, 2014). Yet, during the recovery from anoxia, the heart rate in E. stoutii peaks at almost twice the routine rate (18 bpm) about 1.5 h after O2 is reintroduced into the water (Cox et al., 2010). How the aneural heart is stimulated to beat at the highest rates observed in vivo remains a mystery in need of explanation. Winter survival in ice-covered ponds also requires anoxia tolerance to combat the progressive depletion of O2 by respiration without any appreciable replenishment of O2 through the ice from the atmosphere. The goldfish and crucian carp families, as well as freshwater turtles, display impressive tolerance to anoxia, especially at near freezing temperatures (Clark and Miller, 1973; Farrell and Stecyk, 2007; Herbert and Jackson, 1985a,b; Hicks and Farrell, 2000a,b; Hyvarinen et al., 1985; Lutz and Nilsson, 1997; Stecyk and Farrell, 2002, 2006, 2007; Stecyk et al., 2004b; Warren et al., 2006). Work with the crucian carp and the freshwater turtle has revealed different cardiovascular strategies to survive prolonged anoxia (Clark and Miller, 1973; Farrell and Stecyk, 2007; Hicks and Farrell, 2000a,b; Hyvarinen et al., 1985; Lutz and Nilsson, 1997; Stecyk and Farrell, 2007; Stecyk et al., 2004b; Warren et al., 2006). Similar to the hagfish, some freshwater turtles suppress routine metabolic rate, but more rapidly (within 1–2 h) and to a much greater degree (to between 5 and 10% of the routine rate at 3 °C; Herbert and Jackson, 1985b). Likewise, heart rate and cardiac output in T. scripta are suppressed to a greater degree, 80% (1 bpm) at 5 °C and 52% (10 bpm) at 22 °C in anoxia (Table 1) (Hicks and Farrell, 2000a; Stecyk and Farrell, 2007). As a result, and in conjunction with an arterial hypotension, the ATP requirements of the rhythmic contractions of the heart are easily met by cardiac glycolysis (Farrell and Stecyk, 2007). However, it is clear that temperature plays an important role with regard to maximum glycolytic potential (Q10 of 2–4; Stecyk and Farrell, 2007) and consequently cardiovascular function during anoxia (Q10 of 2.9; Stecyk et al., 2004a; Hicks and Farrell, 2000a,b). Based on the above studies, a reasonable benchmark for maximum glycolytic potential in both turtle and trout hearts at 15 °C appears to be an ATP turnover rate of about 70 nmol ATP·g−1·s−1. This would generate a cardiac power output (PO) of around 0.7 mW·g−1. A pronounced bradycardia is a shared response in both cold- (5 °C) and warm-acclimated (22 °C) T. scripta to reduce the cardiac PO to within the capacity of the

Please cite this article as: Wilson, C.M., et al., The beat goes on: Cardiac pacemaking in extreme conditions, Comp. Biochem. Physiol., A (2014), http://dx.doi.org/10.1016/j.cbpa.2014.08.014

C.M. Wilson et al. / Comparative Biochemistry and Physiology, Part A xxx (2014) xxx–xxx

25 Red-eared slider (5oC) Crucian carp (8oC) Common carp (6oC)

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cardiac glycolytic potential. However, cardiovascular responses to anoxia in cold-acclimated turtles differ from warm-acclimated turtles in several ways. Foremost, cardiac down-regulation due to cold acclimation alone is substantial without any anoxic intervention (Hicks and Farrell, 2000a). For example, normoxic routine PO for T. scripta at 22 °C is 0.66 mW·g−1 and is reduced to 0.044 mW·g−1 at 5 °C, which represents a remarkably high Q10 of 8.8. Similarly, routine PO is 1.05 mW·g− 1 at 21 °C in T. scripta, but only 0.23 mW·g−1 at 5 °C, which represents a Q10 of 2.9 (Stecyk et al., 2004a). Furthermore, warm-acclimated (22 °C) T. scripta retain autonomic control of heart rate during anoxia and respond to acute anoxia with a vagal-mediated bradycardia (Hicks and Farrell, 2000a,b; Stecyk et al., 2004a). In contrast, in cold-acclimated T. scripta, vagal inhibition does not account for the anoxic bradycardia (Hicks and Farrell, 2000b; Stecyk and Farrell, 2007; Stecyk et al., 2004a). Given that cold-acclimated turtles survive for longer periods of time in anoxia when compared to warmacclimated turtles (6 h at 22 °C versus 22 days at 5 °C) it is not easy to delineate the effects of prolonged anoxia and cold acclimation (Herbert and Jackson, 1985a,b; Hicks and Farrell, 2000a,b; Jackson and Ultsch, 1982; Ultsch, 1985; Ultsch and Jackson, 1982a,b). It has been suggested that extrinsic factors triggered by anoxia, such as extracellular acidosis and hyperkalemia, play a role in depressing heart rate and contractility in anoxic, cold-acclimated turtles (Jackson, 2000; Jackson and Ultsch, 1982; Overgaard et al., 2005; Stecyk and Farrell, 2007). Hagfish and T. scripta are decidedly immobile during anoxia. However, the crucian carp (C. carassius) can maintain some activity during anoxia through the up-regulation of glycolysis to meet a decreased ATP demand (Johansson et al., 1995; Nilsson et al., 1993). In addition, lactate and H+ wastes from glycolysis are converted to ethanol and CO2 prior to excretion across the gills (Shoubridge and Hochachka, 1980; Stecyk et al., 2004b). Despite an initial decrease in heart rate and cardiac output at the outset of an anoxic exposure, both return to a stable level that is the same as during normoxia for the remainder of a 5-day anoxic exposure (Stecyk et al., 2004b). It has been suggested that anoxic crucian carp need a fully functional cardiovascular system to transport glycogen and process anaerobic wastes (Stecyk et al., 2004b). Other anoxia- and hypoxia-tolerant members of the carp family are the goldfish (Carassius auratus) and the common carp (Cyprinus carpio). Both species have the ability to survive hours of O2 deprivation at near freezing temperatures and, unlike turtles and hagfish, maintain a reduced level of activity (Lutz and Nilsson, 1997; Stecyk and Farrell, 2002). It has been suggested that the brain of the goldfish operates at a level that maintains control of sensory and locomotory activities during anoxia (Nilsson, 2001; Nilsson and Lutz, 2004), which introduces the possibility of direct autonomic control of heart rate during anoxia. During severe hypoxia, the common carp initially depress heart rate to between 3 and 9 bpm independent of their acclimation temperature (Stecyk and Farrell, 2002). At warm acclimation temperatures (10 °C and 15 °C), heart rate barely attains a new steady state because it is maintained only for 2 h or less before the animals succumb to anoxia (Stecyk and Farrell, 2002). However, when acclimated to 6 °C, common carp can maintain a steady state bradycardia for over 10 h (Stecyk and Farrell, 2002). At all temperatures, intra-arterial atropine injection approximately doubles heart rate indicating that a vagal tone is present and the autonomic control of heart rate remains functional during these relatively short (for champions of anoxia) periods of severe hypoxia (Stecyk and Farrell, 2006). In contrast, when 5 °C-acclimated freshwater turtles were made anoxic, a new myocardial steady state was maintained for 12 days and was evident from the intracellular energy status as well as heart rate, which has led to the suggestion that intracellular energy status and heart rate may be related (Stecyk et al., 2009b). What is interesting is that there appears to be no vagal control involved in maintaining this new steady state at 5 °C because the heart is unresponsive to atropine injections at the cold but not the 21 °C acclimation temperature (Fig. 4; Stecyk et al., 2009b) when intracellular energy status did not reach a new steady state either.

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Time (h) Fig. 4. The effects of anoxia and subsequent atropine injection (open symbols) on the heart rates of red-eared slider freshwater turtle (■), crucian carp (●) and common carp (▲) at cold temperatures (5 °C, 8 °C and 6 °C respectively). An increase in heart rate upon administration shows vagal tone in both the common carp and crucian carp, but the absence of an increase in the turtle indicates that there is no vagal tone in anoxia in this species. Created using data from Stecyk et al. (2009a,b); Stecyk and Farrell (2006); and Stecyk et al. (2004a,b).

5. Fish at supra-optimal water temperatures The second environmental extreme that we considered here is supra-optimal temperatures in fishes. This is relevant for three reasons. Foremost, there is excellent experimental evidence for sockeye salmon (Oncorhynchus nerka) that the scope for heart rate has an optimum temperature very similar to that for aerobic scope and cardiac output (Pörtner and Farrell, 2008; Eliason et al., 2011; Eliason et al., 2013). Second, the responses of cardiac output and metabolic rate to acute warming below the optimum temperature are primarily driven by an increase in heart rate up to a maximum value, because cardiac stroke volume and tissue O2 extraction change relatively little during acute warming. Thus, when a fish is warmed to temperatures that are supra-optimal for aerobic scope, the heart can do little to assist with increasing tissue O2 demands once maximum heart rate is reached. Lastly, evidence is accumulating that the heartbeat becomes irregular (arrhythmic or dysrhythmic) at temperatures around or below CTmax (which is the traditional measure of upper thermal tolerance — the loss of the righting reflex). Thus, if we are to understand how fish can tolerate, acclimate and adapt to elevated temperatures, we need to better understand what sets maximum heart rate, how maximum heart rate varies among species and if maximum heart rate is physiologically adjusted. However, in contrast with the control of heart rate in anoxia, studies that address these important questions are very much in their early days. In vivo studies have acutely warmed a variety of fish species while monitoring routine cardiorespiratory status (Gollock et al., 2006; Clark et al., 2008; Steinhausen et al., 2008; Farrell et al., 2009; Farrell, 2009; Eliason et al., 2011; Penney et al., 2014). For resting fish, heart rate and cardiac output increase with acute warming, but with very little change in cardiac stroke volume. When fishes are warmed while exercising, the primary difference is that increased cardiac stroke volume and increased tissue O2 extraction are exploited to support the tissue O2 demand of the exercise state, and then when the exercising fish is acutely warmed heart rate and cardiac output again increase. However, heart rate of the exercising fish reaches its maximum rate at a lower temperature than in resting fish. Nevertheless, the maximum heart rates reached during warming of resting and exercising salmonids are very similar. Further warming beyond the temperature that generates the maximum heart rate can result in either a plateau at this rate, or a

Please cite this article as: Wilson, C.M., et al., The beat goes on: Cardiac pacemaking in extreme conditions, Comp. Biochem. Physiol., A (2014), http://dx.doi.org/10.1016/j.cbpa.2014.08.014

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decrease in heart rate, but ultimately an irregular heart rate (see, for example, Ferreira et al., in press), which signals a collapse in maximum cardiac output. In fact, Eliason et al. (2013) reported that salmon swimming at a supra-optimal temperature could decrease their heart rate when compared with resting fish at the same temperature. What is still unclear from these in vivo studies is whether the slowing of heart rate and the irregular heartbeats at supra-optimal temperature reflect either increased parasympathetic inhibition via the vagus nerve or a problem intrinsic to cardiac physiology. Valuable insights have accrued from studying the effect of acute warming on maximum heart rate in a growing number of species that include zebrafishes (Sidhu et al., 2014), salmonids (Casselman et al., 2012; Chen et al., 2013; Anttila et al., 2013; Verhille et al., 2013), Arctic cod (Drost et al., 2014) and goldfish (Ferreira et al., in press). The principle behind these assessments is that maximum heart rate can be pharmacologically generated in an anesthetized fish by blocking inhibitory vagal tone to the heart with an injection of atropine, and maximally stimulating β-adrenergic tone with an injection of isoproterenol. A good agreement has been shown for maximum heart rate in vivo and the pharmacologically stimulated rate of anesthetized fish (Casselman et al., 2012). Indeed, maximum heart rate in vivo is remarkably similar to those recorded in situ using pharmacology to maximally stimulate the heart in anesthetized fish (Casselman et al., 2012). The seven species studied so far reveal that maximum heart rate at a given temperature can vary greatly among species. However, in all species studied so far, maximum heart rate is reached at a temperature 3–6 °C below the temperature that triggers the arrhythmic heart rate (Tarr), a temperature that is typically close to but slightly below the CTmax and upper critical temperature (where aerobic scope is close to zero). Thus, these observations suggest that the maximum heart rate reached near the optimum temperature for aerobic scope may be the maximum heart rate that the heart can sustain in any condition. Maximum heart rate is certainly not fixed within a species or between species. Body mass during early juvenile development affects heart rate (e.g., Barrionuevo and Burggren, 1999; Chen et al., 2013). Moreover, warm acclimation resets heart rate to a lower value for a given temperature as well as increasing the upper thermal limit for maximum heart rate, responses that have been observed in vivo and in vitro (Aho and Vornanen, 2001; Keen et al., 1993; Graham and Farrell, 1989; Ferreira et al., in press). Thus, for eurythermal fish species such as goldfish that have good thermal acclimation abilities, the shift to a warmer temperature optimum temperature window with warm acclimation is aided by a lower maximum heart rate at a given temperature as well as a higher absolute maximum heart rate (Ferreira et al., in press). Even Atlantic salmon can increase Tarr by 6 °C when warm acclimated from 10 to 20 °C (Anttila et al., 2013). Other than the obvious requirement for a variable pacemaker potential, we do not understand why the pacemaker and cardiomyocytes operate at a very different heart rate at the same temperature either when species are compared or following thermal acclimation. As noted above, there are indications in some fish that the role of the SR in calcium delivery for the pacemaker potential increases with temperature and in species that experience high water temperatures. Additionally, rainbow trout heart rate has been shown to be sensitive to blockade of IKr with E-4031 sensitivity increasing with temperature (Haverinen and Vornanen, 2007). A larger IKr supports a higher heart rate due to faster repolarization and, therefore, faster activation of HCN channels and the pacemaker potentials. In addition, β-adrenergic modulation of the heart is important. Earlier work with rainbow trout suggested that an increase in SL β-adrenoceptor density and sensitivity increased with cold-acclimation (Keen et al., 1993), but recent work with sockeye salmon (O. nerka) has revealed that SL β-adrenoceptor density can increase with warm acclimation and a sockeye salmon population with a higher upper temperature limit has a very high SL β-adrenoceptor density (Eliason et al., 2011). Finally, recent work suggests that INa is strongly affected by high temperatures and may lead to arrhythmia due to an increasing variability in

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cardiac rhythmicity (Vornanen et al., 2014). Clearly much work lies ahead in this area. In conclusion, we are making significant inroads into our understanding of how the rhythm of the heartbeat is controlled in extreme environments. Perhaps, we should not be surprised to discover anoxia-tolerant animals that can maintain a rhythmic heartbeat without any O2 to generate ATP for cardiac contraction and without autonomic nervous control. Similarly, perhaps we should not be surprised to discover that supraoptimal temperature triggers an arrhythmic heartbeat in all fish tested to date. Lastly, we hope that Bill's heart continues to beat with a strong rhythm for many decades to come. Acknowledgments The authors' research is supported by the Natural Sciences and Engineering Research Council of Canada (10R81993). APF holds a Canada Research Chair. References Aho, A., Vornanen, M., 2001. 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Please cite this article as: Wilson, C.M., et al., The beat goes on: Cardiac pacemaking in extreme conditions, Comp. Biochem. Physiol., A (2014), http://dx.doi.org/10.1016/j.cbpa.2014.08.014