Cardiac activity of the turtle (Trachemys scripta) is greatly depressed with cold acclimation and anoxia. We examined what electrophysiological modifications accompany and perhaps facilitate this depression of cardiac activity. Turtles were first acclimated to 21°C or 5°C and held under either normoxic or anoxic (6 h at 21°C; 14 days at 5°C) conditions. We then measured cardiac action potentials (APs) using spontaneously contracting whole heart preparations and whole cell current densities of sarcolemmal ion channels using isolated ventricular myocytes under appropriate normoxic and anoxic conditions. Compared with 21°C-acclimated turtles, 5°C-acclimated turtles exhibited a less negative resting membrane potential (by 18–29 mV), a 4.7- to 6.8-fold slower AP upstroke rate, and a 4.2- to 4.9-fold greater AP duration. Correspondingly, peak densities of ventricular voltage-gated Na+ (INa) and L-type Ca2+ currents and inward slope conductances of inward rectifier K+ (IK1) channel current were ∼1/7th (Q10 = 3.4), 1/13th (Q10 = 5.0), and one-half (Q10 = 1.4) of those of 21°C-acclimated ventricular myocytes, respectively. With anoxia at 21°C, peak INa density doubled and ventricular AP duration increased by 47%, a change proportional to the reported ∼30% reduction of intrinsic heart rate. In contrast, with anoxia at 5°C, ventricular AP characteristics were unaffected; of the ion currents investigated, only the inward conductance via IK1 changed significantly (reduced by 46%). The present findings indicate that cold temperature, more so than prolonged anoxia, results in substantial modifications of cardiac APs and reduction of ventricular ion current densities. These changes likely prepare cardiac muscle for winter anoxia conditions.
- action potential
- delayed-rectifier potassium channel current
- voltage-gated sodium channel current
- L-type calcium channel current
- inward-rectifier potassium channel current
- red-eared slider turtle
- thermal acclimation
unlike the vast majority of vertebrate species, the red-eared slider freshwater turtle Trachemys scripta is extremely anoxia tolerant. At warm acclimation temperatures (20–25°C), this animal can survive 12–24 h of anoxic submergence; however, when acclimated to 3–5°C, anoxia survival time is extended to ∼45 days (62, 69). During prolonged anoxia exposure at both warm and cold acclimation temperatures, the heart of the turtle continues its role in internal convection, although without oxygen and at a massively reduced rate. Specifically, it is a profound anoxic bradycardia that largely reduces systemic cardiac (Qsys) and power (POsys) outputs by 4.5- to 20-fold after 6 h and 14–21 days of anoxic exposures in warm- and cold-acclimated turtles, respectively (18–21, 57, 59). Heart rate (fH) in turtles decreases from ∼25 to ∼10 beats/min after just 1 h of anoxia at 21–25°C and from a normoxic rate of ∼5 beats/min to <1 beat/min after 24 h of anoxia at 5°C. These reductions in cardiac activity during anoxia reflect and match the reduction of whole-animal metabolic rate and demand for blood flow (18, 24) and also serve as a strategy to ensure that cardiac ATP demand falls well below the cardiac glycolytic capacity to supply ATP (9, 19).
The mechanisms underlying the reduction of cardiac activity during prolonged anoxia are not fully understood. In warm-acclimated turtles, cholinergic cardiac inhibition contributes to ∼36–48% of the anoxic bradycardia (20, 21), but α-adrenergic (58) and adenosinergic (59) cardiac inhibitory mechanisms do not. However, in cold-acclimated turtles, autonomic cardiovascular control is blunted during anoxia and does not account for the anoxic bradycardia (20, 57). Similarly, adenosinergic cardiac inhibition is not involved either (59). Instead, the suggestion has been made that intrinsic electrophysiological changes account for the anoxic depression of cardiac activity in cold-acclimated turtles (20, 41, 56). Certainly, an increased prevalence of atrial-ventricular blocks, a phenomenon in which ventricular contraction rate is less than the atrial contraction rate, in isolated turtle hearts during anoxia exposure (25) suggests a reduced ventricular excitability and/or a delay or blockage of electrical impulses through the atrial-ventricular node. Also, intrinsic electrophysiological modifications are likely involved in the profound depression of cardiac activity with cold acclimation, since cholinergic cardiac inhibition is known to be suppressed (20) and rates of contraction and relaxation are decreased (41).
As a first step to understanding these potential electrophysiological modifications with both cold acclimation and prolonged anoxia exposure, the present study examined hearts from the red-eared slider turtle before and after cold acclimation and before and after prolonged anoxia exposure. Specifically, measurements were made of cardiac action potentials (APs) and current densities of four ventricular sarcolemmal ion channels involved in generating cardiac APs, namely the voltage-gated Na+ (INa), L-type Ca2+ (ICa), inward-rectifier K+ (IK1), and delayed-rectifier K+ (IKr) channel currents (46). Recordings were acquired from four acclimation groups of turtles under conditions similar to those experienced by the turtle. The four acclimation groups were 1) 21°C-acclimated, normoxic; 2) 21°C-acclimated, 6-h anoxia exposure; 3) 5°C-acclimated, normoxic; and 4) 5°C-acclimated, 14-day anoxia exposure. This experimental design allowed comparisons for the effects of temperature alone in normoxic turtles and the effects of temperature alone in anoxic turtles, as well as the effects of anoxia with and across the two temperature acclimation groups. In addition to measuring electrophysiological characteristics under conditions similar to those in which the turtle itself had been acclimated and exposed to, we also made some electrophysiological recordings after an acute change in either the experimental temperature or the perfusate pH. Cardiac APs and ventricular sarcolemmal ion channel currents were rerecorded after an acute temperature change in both 21- and 5°C-acclimated, normoxic hearts. This acute procedure was done to explicitly distinguish the effects of cold acclimation from acute and perhaps direct effects of ambient temperature on the rate of these physiological processes (termed here direct temperature effects). Specifically, the acute temperature change was from either 5 to 21°C or from 21 to 5°C for the AP recordings, and the acute temperature change was to the common temperature of 11°C for the ion channel current recordings. To mimic in vivo blood plasma pH of anoxia-exposed turtles, cardiac APs were rerecorded for anoxia-acclimated hearts after a switch to a combined acidotic and anoxic saline. This was done to investigate the effect of acidosis on turtle cardiac APs given the reported temperature dependency of its negative inotropic and chronotropic effects on the turtle myocardium (41, 49, 56, 71, 73). For sarcolemmal ion channel current recordings, we focused on ventricular myocytes because the ventricle is the power-generating tissue of the turtle heart. Our prediction was that changes in AP shape and duration induced by cold acclimation and/or prolonged anoxia exposure would be reflected in changes in ion current densities. Also, because the duration of cardiac contraction and AP duration (APD) are closely correlated in other ectothermic vertebrate species (e.g., Refs. 43, 55), we reasoned that changes in cardiac APD accompany the known decreases in fH associated with both cold acclimation and with prolonged anoxia exposure.
MATERIALS AND METHODS
Fifty-six red-eared sliders (Trachemys scripta, Gray) with body masses ranging between 124 and 518 g [235 (SD 77) g] were used in this study. Turtles were obtained from Lemberger (Oshkosh, WI) and The Charles D. Sullivan (Nashville, TN) and shipped by air to the University of Joensuu, Joensuu, Finland, or the University of British Columbia, Vancouver, Canada. The exposure design for the turtles was normoxia and anoxia exposure at 21 and 5°C acclimation. In vitro measurements were made at the same temperature as the acclimation temperature (unless stated otherwise; i.e., an acute temperature change) and in the appropriate normoxic or anoxic saline. Turtles studied at 21°C were held indoors in aquaria under a 12:12-h light-dark photoperiod, had free access to basking platforms and diving water, and were fed several times a week with commercial turtle food pellets. The turtles studied at 5°C were kept in aquaria with shallow water (3–4 cm) under a 12:12-h light-dark photoperiod in a temperature-controlled room set to 5°C for 5 wk before experimentation to allow adequate time for cold acclimation (19). Acclimation to 5°C occurred during winter months, and turtles were fasted during the acclimation period. Normoxic turtles were sampled from these conditions. For prolonged anoxia, 21°C turtles were exposed to anoxia for 6 h and 5°C turtles for 14 days. These anoxia durations were utilized to be consistent with previous examinations of the anoxic turtle heart and because anoxic cardiovascular status is relatively stable at these times (19, 20, 56, 57). The anoxic conditions were achieved by individually placing turtles into an enclosed, water-containing plastic chamber that still allowed access to air for 24 h, after which the plastic chamber was completely filled with water, continuously bubbled with N2 (water Po2 of <0.3 kPa), and access to the water surface denied by means of mesh suspended below the surface of the water. Turtles acclimated at 21°C were not comatose after the anoxia exposure, whereas 5°C-acclimated turtles were found to be unresponsive to tactile stimulation.
All experiments were conducted with the consent of the University of British Columbia or University of Joensuu committee for animal experimentation.
AP Recordings From Intact Cardiac Tissue
Intracellular APs were measured from all three cardiac chambers (right atrium, left atrium, and ventricle) of a spontaneously beating whole heart preparation. The heart was accessed through removal of a 2 cm × 2 cm piece of the plastron using a bone saw after death by decapitation, which for anoxia-acclimated turtles occurred underwater in the plastic containers. The chambers of the excised heart were then medially opened, spread, and gently fixed with insect pins to the Sylgard-coated bottom of a 10-ml, water-jacketed tissue chamber filled with physiological saline containing (in mmol/l) 125 NaCl, 2.5 KCl, 2 CaCl2, 1 MgSO4, 1 NaH2PO4, 10 HEPES, and 5 glucose, as well as a physiologically relevant tonic (1 nmol/l) epinephrine concentration (glucose and epinephrine were added immediately before use). The desired temperature of the saline (see below) was maintained with a circulating water bath, and saline was bubbled continuously with either O2 (for normoxia-acclimated hearts) or N2 (for anoxia-acclimated hearts; saline Po2 was ∼2 kPa). Saline pH was adjusted (Teopal P600, Teo-Pal, Espoo, Finland) to 7.75 with NaOH at 21°C and allowed to change with temperature. Thus pH was ∼7.95 at 5°C. Saline was refreshed every 30 min throughout the 4- to 6-h recording period to avoid potential buildup of anaerobic waste products with anoxia-acclimated hearts and epinephrine degradation. Saline for anoxia-acclimated hearts was prebubbled with N2.
Hearts were allowed to stabilize to the experimental conditions for 30–45 min before recordings were made. For normoxia-acclimated hearts, APs and spontaneous fH were recorded first at the acclimation temperature of the animal (i.e., 21 or 5°C), and then the saline temperature acutely changed either from 5 to 21°C or from 21 to 5°C; hearts were allowed 45–60 min to stabilize before APs and spontaneous fH were rerecorded for that preparation. For anoxia-acclimated hearts, APs and spontaneous fH were recorded only at the acclimation temperature of the animal but were also recorded after a switch to acidotic saline (pH 7.25 at 21°C and pH 7.55 at 5°C, still continuously bubbled with N2), which mimicked in vivo blood plasma pH of turtles exposed to anoxia for 6 h (warm acclimated) or 14 days (cold acclimated) (63, 72). A 25- to 30-min stabilization period was allowed after the switch to the acidotic saline before APs and spontaneous fH were rerecorded for that preparation.
Cardiac APs were recorded with high-resistance, sharp microelectrodes (6–30 MΩ when filled with either 3.0 or 0.3 mol/l KCl), fabricated from borosilicate glass with an internal filament (World Precision Instruments, Sarasota, FL) and using an L/M-3P-A vertical puller (List Medical, Darmstadt, Germany). KCl (0.3 mol/l) was utilized in some experiments to ensure that observed changes in resting membrane potential (RMP) with temperature were not an artifact of the 3.0 mol/l KCl. Microelectrode signals were amplified by a high-impedance amplifier (KS-700, World Precision Instruments), digitized at a sampling rate of 2 kHz (Digidata 1200, Axon Instruments, Union City, CA), and recorded to computer using Axotape 2.2 acquisition software. Spontaneous fH was calculated from the peak-to-peak intervals of right atria contraction force, which was obtained via attachment of one edge of the right atria to a force transducer (FT03, Grass Instruments, West Warwick, RI) by a small metal hook and braided silk thread. The contraction force signal was amplified (7D, Grass Instruments), routed through the digitizer, and stored to computer at a sampling rate of 200 Hz. APs and contraction force recordings were analyzed off-line using Clampfit 9.2 (Axon Instruments, Foster City, CA).
Whole Cell Voltage Clamp From Isolated Myocytes
Whole cell voltage-clamp experiments were performed on individual ventricular myocytes to assess the effect of cold temperature acclimation and prolonged anoxia acclimation on ion current density of INa, ICa, IK1, and IKr.
Single ventricular myocytes were enzymatically isolated by adapting an established isolation protocol for teleost fish (51, 65). The turtle heart was accessed as describe above, excised, and cannulated through the left aortic arch into the ventricle. The heart was then perfused retrograde at room temperature (21 ± 1°C) from a height of 50 cm, first with a nominally Ca2+-free, low-Na+ isolation saline solution (containing, in mmol/l, 100 NaCl, 10 KCl, 4 MgSO4, 1 NaH2PO4, 1.2 KH2PO4, 50 taurine, 20 glucose, and 10 HEPES, with pH adjusted to 6.9 at 21°C with KOH) for 10 min and then for 20 min with fresh isolation solution supplemented with the proteolytic enzymes collagenase (1.5 mg/ml; type IA) and trypsin (1 mg/ml; type IX) and with 1.5 mg/ml of fatty acid-free BSA. All chemicals were purchased from Sigma-Aldrich (St. Louis, MO). The isolation solutions were continuously bubbled with O2 for normoxia-acclimated hearts or N2 for anoxia-acclimated hearts, and the enzyme-supplemented isolation solution was recycled with the use of a peristaltic pump and retained after perfusion. The ventricle was then dissected from the atria and the sinus venosus, minced with scissors in fresh isolation solution, and transferred to the retained enzyme-supplemented isolation solution. Ventricular tissue was gently stirred with a small magnetic bar at room temperature with periodic trituration through the opening of a Pasteur pipette for 10–20 min or until individual viable myocytes were observed by light microscopy. The solution was then left to settle, and myocytes were resuspended in fresh isolation solution and stored at 6°C. Cells were usually recorded from within 4–6 h, as done previously for recordings from anoxia-acclimated crucian carp (Carassius carassius) cardiomyocytes (42, 67). However, in some instances, cells were stored for up to 12 h because of the extreme difficulty in obtaining reliable measurements from viable cells.
Electrophysiological measurements and analysis of sarcolemmal current densities were achieved with the use of established methods and solutions for teleost fish that were adapted for the turtle (14, 15, 42, 51, 55, 65, 66). Specific details for each current measured are given below; however, in all instances, an aliquot of dissociated myocytes was placed into a recording chamber mounted on the stage of an inverted microscope and left to adhere to the bottom of the chamber. Cells were then superfused at a rate of 1–2 ml/min with an extracellular saline solution. Temperature of the extracellular solution was regulated either by water bath circuits that chilled or heated the inflow tube carrying the extracellular solution to the recording chamber or a Peltier device. Thermocouples positioned no less than 5 mm from the cell under investigation were used to continuously monitor temperature. For cells from normoxia-acclimated animals, current density was recorded in the same cell first at the acclimation temperature of the animal (i.e., 21 or 5°C) and then after an acute exposure to the common experimental temperature of 11°C (temperature change was accomplished within 3–5 min). This was done to distinguish cold-acclimation effects from direct temperature effects. For cells from anoxia-acclimated animals, current density was recorded only at the acclimation temperature of the animal, and the extracellular solution was continuously bubbled with N2 (Po2 of the extracellular solution in the recording chamber was ∼5 kPa).
Patch pipettes were pulled from borosilicate glass without an internal filament and had a resistance of 2–4 MΩ when filled with pipette solution. Offset potentials were zeroed just before formation of the gigaohm seal, and pipette capacitance was compensated after formation of the gigaohm seal. The patch was ruptured by delivering a short voltage pulse (zap) to the cell, and capacitive transients were eliminated by iterative adjustments of series resistance and cell capacitance circuits. Mean series resistance was 5.3 MΩ (SD 4.9) (n = 165), and mean cell capacitance was 50.2 ± 1.0 pF (n = 165; mean ± SE).
INa recordings were made (at the University of Joensuu) with an EPC-9 amplifier in conjunction with Pulse v8.65 software (HEKA, Lambrecht, Germany) and a temperature-controlled 500-μl recording chamber (RCP-10T; Dagan, Minneapolis, MN). Patch pipettes were pulled from borosilicate glass (Garner F-78045, Claremont, CA) using a two-stage vertical puller [either a L/M-3P-A (List Medical) or PP-83 (Narishige, Tokyo, Japan)]. Cells were first superfused with normal K+-based extracellular solution containing (in mmol/l) 125 NaCl, 2.5 KCl, 2 CaCl2, 1 MgSO4, 1 NaH2PO4, 10 HEPES, 5 glucose, and 0.01 nifedipine (to block L-type Ca2+ channels; Ref. 12), where gigaohm seal and whole cell patch-clamp recordings of the myocytes were established. Internal perfusion of the myocytes with pipette solution (containing, in mmol/l, 5 NaCl, 130 CsCl, 1 MgCl2, 5 EGTA, 5 MgATP, and 5 HEPES; pH adjusted to 7.4 at 21°C with CsOH) continued for at least 3 min to allow buffering of intracellular Ca2+ with EGTA. The extracellular solution was then switched to a low-Na+ solution (containing, in mmol/l, 19 NaCl, 108.5 CsCl, 1 MgSO4, 1 NaH2PO4, 2 CaCl2, 10 HEPES, 5 glucose, and 0.01 nifedipine; pH adjusted to 7.75 at 21°C with CsOH) without inducing contracture in the patched myocyte, as previously accomplished in teleost fish myocytes (15, 16). INa was elicited in the low-Na+ extracellular solution from the holding potential of −80 mV by 10-ms (21°C) or 30-ms (5°C) depolarizing square pulses to voltages between −100 and 60 mV in 10-mV steps that were preceded by a 20-ms prepulse to −120 mV to remove inactivation. Sampling rate was 20 kHz, and the signal was filtered on-line with a 10-kHz Bessel filter. Voltage off-set caused by series resistance was compensated (70%; 10 μs). Leak current was estimated from current at end of depolarizing pulses at −100, −90, −80, and −70 mV and subtracted off-line. The amplitude of INa was calculated as the peak inward current during the depolarizing pulses.
ICa recordings were made (at the University of British Columbia) with an Axopatch 200B amplifier, a CV 203BU headstage, and ClampEx v9.2 software (Axon Instruments). An RC-26GLP recording chamber (234 μl; Warner Instruments, Hamden, CT) was used, and temperature was regulated with a PHC-2 heater/cooler jacket in conjunction with a SC-20 dual in-line solution heater/cooler and a CL-100 bipolar temperature controller (Warner Instruments). Pipettes were pulled from borosilicate glass (GC150T-7.5; Harvard Apparatus, St. Laurent, QC, Canada) with a Sutter P-97 puller (Sutter Instruments, Novato, CA). Myocytes were superfused with a Cs+-based extracellular solution to eliminate contaminating K+ currents. Extracellular solution contained (in mmol/l) 125 NaCl, 2.5 CsCl, 2 CaCl2, 1 MgSO4, 1 NaH2PO4, 10 HEPES, and 5 glucose; pH was adjusted to 7.75 at 21°C with CsOH. The pipette solution contained (in mmol/l) 130 CsCl, 1 MgCl2, 5 Na2-phosphocreatine, 4 MgATP, 0.03 Na2GTP, 5 EGTA, 15 tetraethylammonium chloride (to block K+ currents; Ref. 23), and 10 HEPES; pH was adjusted to 7.4 at 21°C with CsOH. ICa was elicited from the holding potential of −70 mV by 500-ms depolarizing square pulses to voltages between −70 and +70 mV in 10-mV steps. A preceding 50- or 100-ms prepulse to −40 mV was used to inactivate voltage-gated Na+ channels and eliminate fast Na+ currents, as the turtle ventricular myocytes are relatively insensitive to the specific Na+ blocker TTX (12). Sampling rate was 10 kHz, and signals were low-pass filtered at 2 kHz on-line with the Axopatch amplifier. Signals were analyzed off-line with Clampfit 9.2 software (Axon Instruments). The amplitude of ICa was calculated as the difference between peak inward current and the current at the end of the depolarizing pulse.
Additionally, because ICa can run down over time when measured in whole cell configuration and the experimental protocol utilized here required repeated measurements of ICa, the magnitude of ICa rundown was assessed in a separate group of myocytes. This was accomplished by repeatedly measuring ICa at 3-min intervals over a period of 15 min (Fig. 1).
IK1 and IKr.
IK1 and IKr recordings were made (at the University of Joensuu) with an Axopatch 1D amplifier, a CV-4 1/100 headstage, and ClampEx v8.2 software (Axon Instruments). A RC-26 recording chamber (150 μl; Warner Instruments) was used, and temperature was regulated with water bath circuits. Patch pipettes were pulled from borosilicate glass (Garner F-78045) with the use of a two-stage vertical puller [either an L/M-3P-A (List Medical) or PP-83 (Narishige)]. Cells were superfused with normal K+-based extracellular solution (described above). Pipette solution contained (in mmol/l) 140 KCl, 1 MgCl2, 5 EGTA, 4 MgATP, and 10 HEPES; pH was adjusted to 7.4 at 21°C with KOH. IK1 was measured relative to zero membrane current at the end of 1,000-ms square voltage pulses that were elicited from a holding potential of −80 mV to voltages between −120 and 20 mV in 20-mV steps. No action was taken to abolish the faster INa, either by pharmacological blockade or by a prepulse to −40 mV, during IK1 recordings because Na+ channel activation and inactivation were completed ∼500 ms (at 5°C) and ∼800 ms (at 21°C) before the time at which IK1 was measured. IKr was measured as an outward tail current at −40 mV after 4,000-ms depolarizing square pulses between −80 and 80 mV in 20-mV steps elicited from the holding potential of −40 mV. Signals were sampled at 2 kHz, low-pass filtered on-line at 2 kHz, and analyzed off-line with Clampfit 9.2 software (Axon Instruments).
Data and Statistical Analysis
All results are expressed as means ± SE. The number of observations (n) for cardiac AP data was number of turtles (i.e., 4 or 5 for each of the 4 acclimation groups), with AP characteristics from one to six cells per tissue per animal averaged for each individual. AP shape and duration were quantified by measuring RMP and peak potential and calculating duration to 0 mV (APD0), 50% (APD50), 90% (APD90), and 100% (APD100) repolarization. AP upstroke rate was calculated by dividing the difference in RMP and peak potentials by upstroke duration. Number of myocytes constitutes n for whole cell voltage-clamp experiments. Cells were obtained from 2–10 animals for each exposure condition. Densities of INa, ICa, IK1, and IKr (expressed as pA/pF) were calculated by dividing measured currents by the cell capacitance. Slope conductance (pS/pF) of inward-rectifier K+ channel was calculated for the linear region of the IK1 current-voltage plot by dividing the difference in IK1 between −120 and −100 mV by the change in voltage (i.e., 20 mV). Statistically significant differences in AP characteristics and ion current densities between either 21- and 5°C-acclimated turtles or normoxia- and anoxia-exposed turtles of the same acclimation temperature were determined with a two-way ANOVA or a t-test where appropriate. Two-way repeated measures ANOVA tests, or paired t-tests where appropriate, were used to compare AP characteristics or ion current densities after an acute temperature change, and introduction of acidic saline (i.e., AP characteristics). In all instances, P < 0.05 was used as the level of significance. Where appropriate, multiple comparisons were performed with Student-Newman-Keuls tests.
Effect of Temperature on Spontaneous Normoxic fH
The initial spontaneous fH of 21°C- and 5°C-acclimated heart preparations were 35.3 ± 1.3 beats/min (n = 5) and 5.5 ± 0.7 beats/min (n = 5), respectively. This sixfold difference in spontaneous fH with acclimation to 5°C corresponds to a Q10 value of 3.2. Acute exposure of 21°C-acclimated hearts to 5°C decreased spontaneous fH to a rate (3.0 ± 0.7 beats/min; n = 3) statistically similar to the 5°C-acclimated fH. Conversely, acute exposure of 5°C-acclimated hearts to 21°C significantly increased spontaneous fH to 27.3 ± 0.2 beats/min (n = 3), but this rate was significantly lower than the 21°C-acclimated fH. These results suggest that direct temperature effects rather than temperature acclimation were predominant in setting the spontaneous fH under normoxic conditions.
Effect of Temperature on Cardiac Action Potentials in Normoxia
Like spontaneous fH, the shape and duration of the cardiac AP were considerably modified by temperature. Primarily, after acclimation to 5°C, the RMP was significantly less polarized (by 18–26 mV) in all chambers of the heart (Figs. 2, 3, A and C, 4, A and C, and 5, A and C). Also, acclimation to 5°C significantly decreased AP upstroke rate by 4.7- to 6.8-fold in all cardiac chambers, which corresponded to Q10 values of 3.3 for the right atria, 2.8 for the left atria, and 2.6 for the ventricle (Figs. 2, 3, A and C, 4, A and C, and 5, A and C; Table 1). Furthermore, APD was prolonged at least fourfold in all cardiac chambers compared with 21°C-acclimated hearts (Figs. 2, 3, A and C, 4, A and C, 5, A and C). Specifically, APD100 of 5°C-acclimated hearts was increased by 4.4-fold in the right atria (614.0 ± 39.0 to 2,680.5 ± 113.5 ms; Fig. 2, A and C), 4.9-fold in the left atria (590.0 ± 50.5 to 2,894.0 ± 198.5 ms; Fig. 3, A and C), and 4.2-fold in the ventricle (850.0 ± 38.5 to 3,559.0 ± 529.5 ms; Figs. 2 and 5, A and C). Q10 values for the prolongation of APD at 5°C (calculated from reciprocal values of APD90) were 2.4, 2.6, and 2.6 for the right atria, left atria, and ventricle, respectively. These Q10 values point to direct temperature effects playing a predominant role in contributing to the prolongation of APD with cold acclimation, a conclusion that was confirmed by the results for the acute temperature change.
Acute exposure of 21°C-acclimated heart to 5°C, as well as acute exposure of 5°C-acclimated heart to 21°C (Fig. 6), mimicked many of the AP differences observed between the temperature-acclimated hearts. For instance, the AP upstroke rate was reduced with an acute 5°C exposure for all cardiac chambers (Fig. 6; Table 1) such that the AP upstroke rates for the left atria and ventricle were identical to those measured in cardiac tissue acclimated to 5°C. Also, for all cardiac chambers, all of the indexes of APD (APD0, APD50, APD90, and APD100) were not significantly different from those of the 5°C-acclimated heart (Fig. 6, A, C, and E). Similarly, the ventricular AP shape of 5°C-acclimated heart acutely exposed to 21°C was not statistically significant different from that of the 21°C-acclimated heart (Fig. 6F). Thus the changes in AP shape and duration associated with cold acclimation were predominantly determined by a direct temperature effect rather than the chronic effect of temperature acclimation.
Nevertheless, not all changes in AP characteristics with cold acclimation could be attributed solely to a temperature effect, as some changes were not replicated by an acute temperature change. Notably, acute exposure of 21°C-acclimated hearts to 5°C did not result in a statistically significant depolarization of RMP like that associated with 5°C acclimation (Table 1). Even so, acute exposure of all cardiac chambers from 5°C-acclimated hearts to 21°C did result in a significant decrease in RMP to a membrane potential statistically similar to the 21°C-acclimated RMP (Table 1). These disparate findings indicate that the mechanism underlying the increase in RMP with cold acclimation can be more quickly reversed than initiated. Furthermore, APD90 and APD100 of right and left atria were significantly longer in 5°C-acclimated hearts acutely exposed to 21°C than APD90 and APD100 of 21°C-acclimated atria (Fig. 6, B and D).
Effect of Temperature on Ventricular Sarcolemmal Ion Channel Current Densities
Consistent with the changes in AP characteristics with cold acclimation, current densities of ventricular sarcolemmal ion channels involved in generating APs, as elicited from square voltage-clamp pulse protocols, were drastically reduced in 5°C-acclimated ventricular myocytes compared with 21°C-acclimated cells (Fig. 7). However, the manner through which current density is depressed with cold acclimation differed with each channel type (Figs. 8–10).
Consistent with the reduced AP upstroke rate at 5°C, in 5°C-acclimated ventricular myocytes, INa density was significantly reduced (Fig. 7A), and the kinetics of sodium channel activation and inactivation were slower (Fig. 9) than shown for 21°C-acclimated ventricular myocytes. Peak INa density was 7.3-times less in 5°C-acclimated ventricular myocytes (−1.2 ± 0.1 pA/pF) than in 21°C-acclimated ventricular myocytes (−8.7 ± 0.9 pA/pF). The Q10 values were 3.4 for the decrease in INa, 2.5 for sodium channel activation, and 1.9 for INa inactivation time (the latter values were calculated from reciprocal values at −20 mV) (Figs. 8A and 9). When measured at a common temperature of 11°C, peak INa density did not differ between 5°C-acclimated (−3.3 ± 0.4 pA/pF; Q10 was 5.4 for the acute temperature change) and 21°C-acclimated (−4.1 ± 1.2 pA/pF; Q10 was 2.1 for the acute temperature change) ventricular myocytes (Fig. 8A). Thus, although the density of functional sodium channels on ventricular myocytes has a clear temperature dependency, it was unchanged with cold acclimation.
The reduction in ICa with acclimation to 5°C (Fig. 7B) was more profound than INa. Peak ICa density of 5°C-acclimated myocytes (−0.43 ± 0.03 pA/pF) was ∼1/13th the peak ICa density of 21°C-acclimated myocytes (−5.7 ± 0.5 pA/pF). The Q10 for this depression in ICa with acclimation to 5°C was 5.0. When measured at a common temperature of 11°C, peak ICa density of 5°C-acclimated myocytes (−1.3 ± 0.09 pA/pF; Q10 was 6.3 for the acute temperature change) remained significantly lower than 21°C-acclimated myocytes (−2.2 ± 0.3 pA/pF; Q10 was 2.5 for the acute temperature change) (Fig. 8B). Therefore, in addition to the negative effect of cold temperature and unlike INa, a component of the decreased ICa of 5°C-acclimated myocytes involved the downregulation of functional L-type Ca2+ channels.
The inward IK1 density and the inward slope conductance of inward-rectifier K+ channels were significantly reduced with acclimation to 5°C (Fig. 7C), findings consistent with the depolarized RMP and prolonged APD of 5°C-acclimated ventricular tissue. At −120 mV, inward IK1 density of 5°C-acclimated myocytes (−4.6 ± 0.3 pA/pF) was 26% less than the inward IK1 density of 21°C-acclimated myocytes (−6.2 ± 0.7 pA/pF). Inward slope conductance of inward-rectifier K+ channels between −120 and −100 mV was reduced by almost 50% from 214.7 ± 16.8 pS/pF at 21°C to 125.3 ± 9.6 pS/pF at 5°C. When measured at a common temperature of 11°C, 5°C-acclimated ventricular myocytes displayed greater inward IK1 density at −120 mV (−5.5 ± 0.3 pA/pF compared with −3.3 ± 0.4 pA/pF; Fig. 10) and greater inward rectifier K+ channel slope conductance (191 ± 8.6 pS/pF compared with 89.8 ± 11.0 pS/pF; Fig. 8C) than 21°C-acclimated ventricular myocytes. The fact that the Q10 for the reductions in slope conductance was 1.4 when comparing acclimated hearts but around 2 for an acute temperature change lends support to the notion that the density of functional inward-rectifier K+ channels was upregulated with cold acclimation (Fig. 8C). In contrast to inward IK1 density, no significant difference in outward IK1 density (i.e., at −80 and −60 mV) existed between 5°C-acclimated ventricular myocytes and 21°C-acclimated ventricular myocytes (Fig. 7C). This finding indicates that temperature did not affect IK1 equally at all voltages. However, like inward IK1 and inward slope conductance, outward IK1 density of 5°C-acclimated myocytes was significantly greater than 21°C-acclimated myocytes at −80 and −60 mV when measured at the common temperature of 11°C (Fig. 10). This finding further supports the notion that the density of functional inward-rectifier K+ channels was upregulated with acclimation to 5°C to partially compensate for the negative effect of cold temperature on IK1.
IKr was minor and no different for 21- and 5°C-acclimated ventricular myocytes (Fig. 7D), and this situation was unchanged after acute exposure to 11°C (data not shown).
Effect of Prolonged Anoxia on Spontaneous fH
Spontaneous fH of 5°C anoxic turtles (3.0 ± 0.7 beats/min; n = 5) was significantly 45% lower than the spontaneous fH (5.5 ± 0.7 beats/min; n = 5) of 5°C normoxic turtles under normoxic conditions. At 21°C, spontaneous fH of anoxic preparations was 27.4 ± 4.6 beats/min (n = 5); although this was 32% lower than the 21°C spontaneous fH (35.3 ± 1.3 beats/min; n = 5) under normoxic conditions, there was no statistically significant difference. These findings are consistent with our other work (56) demonstrating that prolonged anoxia at both 21 and 5°C resets intrinsic fH to a rate 25–53% lower than during normoxia.
Effect of Prolonged Anoxia on Cardiac Action Potentials
In contrast to the large effects of temperature on AP shape and duration, prolonged anoxia exposure caused few and only small changes in cardiac APs. Furthermore, the effect of anoxia on cardiac APs was cardiac chamber specific unlike the chamber-independent effects of temperature on AP shape and duration. At 21°C, APs for the right and left atria were not significantly modified after 6 h of anoxic exposure (Figs. 3, A and B, 4, A and B). Likewise, AP shape or duration of all cardiac chambers remained unchanged after 14 days of anoxia at 5°C (Figs. 2, 3, C and D, 4, C and D, and 5, C and D). In contrast, ventricular APD50, APD90, and APD100 were increased significantly by 39%, 49%, and 47%, respectively, after anoxia exposure at 21°C (Figs. 2 and 5, A and B). Therefore, the prolongation of the ventricular APD after prolonged anoxia at 21°C is of the same magnitude as the reduction in spontaneous fH.
Effect of Prolonged Anoxia on Ventricular Sarcolemmal Ion Channel Current Densities
Of the four ventricular sarcolemmal membrane currents examined, prolonged anoxia at 21°C only significantly altered INa. Specifically, peak INa density doubled from −8.7 ± 0.9 to −16.1 ± 1.7 pA/pF with 6 h of anoxia at 21°C (Fig. 11A), although without affecting activation and inactivation kinetics of INa (Fig. 9). No significant changes in ICa, IK1, or IKr density occurred as a result of 6 h of anoxia exposure at 21°C (Fig. 11, B–D).
Similar to findings at 21°C, peak ICa density was unaffected by 14 days of anoxia at 5°C (Fig. 12B). However, peak INa of anoxic 5°C-acclimated ventricular myocytes (−1.1 ± 0.2 pA/pF) was not significantly different than that for normoxic 5°C-acclimated myocytes (−1.2 ± 0.1 pA/pF; Fig. 12A), unlike at 21°C. When recorded with higher (125 mM) extracellular sodium concentration to enhance INa current density, there was still no difference between anoxic (−17.6 ± 2.6 pA/pF) and normoxic (−18.0 ± 2.6 pA/pF) 5°C-acclimated myocytes. This result excludes the possibility that the measurement of INa under reduced sarcolemmal Na+ gradient resulted in minimal INa current density at 5°C and thus an inability to detect small changes in INa occurring with prolonged anoxia exposure. At 5°C, and similar to the results at 21°C, prolonged anoxia exposure did not significantly alter the activation and inactivation kinetics of the sodium current (Fig. 9). Thus, although the effect of prolonged anoxia exposure on INa density was temperature dependent, the effect of anoxia on sodium channel activation and inactivation kinetics was temperature independent.
The effects of prolonged anoxia on IK1 were temperature dependent. Prolonged anoxia at 5°C significantly reduced inward IK1 density and inward slope conductance of inward-rectifier K+ channels (Fig. 12C), whereas neither effect occurred with anoxia at 21°C (Fig. 11C). IK1 density of 5°C anoxia-acclimated myocytes was significantly 33% and 18% lower than IK1 density of 5°C normoxia-acclimated myocytes at −120 and −100 mV, respectively. Inward slope conductance of inward-rectifier K+ channels was significantly reduced by 45% from 125.3 ± 9.6 to 68.3 ± 6.6 pS/pF. In contrast, no statistical differences in outward IK1 density occurred with prolonged anoxia exposure at 5°C. IKr remained minor after prolonged anoxia at 5°C (Fig. 12D).
The Combined Effect of Acidosis and Anoxia on Spontaneous fH and Cardiac Action Potentials
Spontaneous fH of anoxia-acclimated spontaneously beating whole heart preparations was unaffected by acute acidosis exposure. At 21°C, spontaneous fH of anoxia-acclimated heart preparations before acidosis was 21.6 ± 5.2 beats/min (n = 5) and 19.3 ± 3.5 beats/min (n = 5) with acidosis. Similarly, at 5°C, spontaneous fH was unchanged before (3.8 ± 0.7 beats/min; n = 5) and with acidosis (4.3 ± 0.3 beats/min; n = 5).
In contrast, acute acidosis exposure altered the AP shape and duration depending on the acclimation temperature and cardiac chamber. At 21°C, right atrial APD0, APD50, APD90, and APD100 were increased significantly by 18–20% with acidosis (Fig. 3B), but left atria and ventricular AP shape and duration were unaffected (Figs. 4B and 5B). At 5°C, APD50 increased by 18% in the left atria with acidosis (Fig. 4D), but right atria and ventricular AP shape and duration were unaffected (Figs. 3D and 5D).
The present study examined whether modification of electrophysiological properties of the turtle heart facilitates the downregulation of cardiac activity that accompanies cold acclimation and prolonged anoxia exposure. To this end, we compared cardiac APs of spontaneously contracting turtle heart preparations and whole cell current densities of key sarcolemmal ion channels of ventricular myocytes isolated from turtles that had been acclimated to either 21 or 5°C and exposed to either normoxia or prolonged anoxia. Our findings revealed that the substantial modifications of cardiac APs and reduction of ion current densities that likely contribute significantly to decreased cardiac activity were more evident with cold temperature, both direct and acclimation effects, than with prolonged anoxia exposure. In view of this finding, we suggest that the reduction in electrophysiological activity of the turtle heart associated with cold exposure is a critical component in the preparation of cardiac muscle so that turtles can successfully overwinter under anoxia conditions.
Critique of Methods
AP recordings from intact cardiac tissue in vitro.
A difficulty with in vitro work is that both the isolation procedure and the test conditions may not exactly mimic the in vivo situation, especially when trying as we did to maintain the anoxic condition of the tissue. Clearly, a caveat with our AP results for anoxia-acclimated turtles is that the recording chamber saline Po2 could not be completely depleted of oxygen due to practical limitations of the experimental setup. However, we were greatly encouraged by spontaneous fH for the normoxic 21- and 5°C-acclimated heart preparations being close to previously reported in vivo and in vitro intrinsic rates (8, 20, 72). Similarly, the reduced spontaneous fH of anoxic 21- and 5°C-acclimated heart preparations is qualitatively and quantitatively comparable with previous in vivo and in vitro work (20, 55). Thus, if there had been a problem with anoxia-acclimated hearts reverting to their normoxic state during isolation and measurement, we would not have detected the decreased intrinsic fH or seen the similarity to previous studies. Furthermore, the clear, chamber-specific effect of anoxia acclimation on APs at 21°C despite similar measurement conditions argues against the possibility that the lack of effects of anoxia acclimation on cardiac APs at 5°C were a measurement artifact. Therefore, we are confident that our novel recordings of turtle APs for all three cardiac chambers and under all four acclimation conditions were from viable cardiac tissues and representative for the prior exposure history of the animal. Nevertheless, our methods could have underestimated some effects of anoxia.
Whole cell patch clamp in isolated ventricular myocytes.
For turtles, we have provided the first measurements of ventricular sarcolemmal INa, IK1, and IKr at any acclimation temperature or condition (i.e., normoxia or anoxia exposed). Our novel measurements of ventricular ICa of warm-acclimated, anoxia-exposed turtles and cold-acclimated, normoxia- and anoxia-exposed turtles add to a very recent report of ICa for 20- to 21°C-acclimated yellow-bellied turtles (Trachemys scripta scripta) (12).
Like the anoxia-acclimated turtle AP data, a caveat with our recordings of INa, ICa, IK1, and IKr from anoxia-acclimated turtles is that cardiomyocytes could not be maintained anoxic throughout all steps of the isolation and recording protocols. In particular, cells were not continuously bubbled with N2 during storage at 6°C and were exposed to air when left to settle in the recording chamber, a step that lasted only 1–2 min at 21°C and up to 5 min at 5°C, and recording chamber saline Po2 could not be completely depleted of oxygen. However, we are confident that our ion-channel current recordings from anoxia-acclimated turtles faithfully represent the prior anoxia exposure history of the animal. First, this is because differences in ion channel current densities were indeed found for cardiomyocytes before and after anoxia acclimation, the currents affected by anoxia acclimation were temperature dependent (i.e., INa at 21°C and IK1 at 5°C), and the directional changes in current amplitude with anoxia exposure were channel dependent (Figs. 11 and 12). Furthermore, the finding that peak ICa remained unchanged with anoxia exposure appears legitimate, at least for 5°C-acclimated turtles, since twitch force and time-to-peak force of ventricular tissue from 5°C-acclimated turtles does not differ between normoxia- and anoxia-acclimated animals (41). Finally, given that at the organismal level the changes in cardiovascular status with anoxia take ∼1 h and ∼24 h at 21 and 5°C, respectively (18–21, 57, 59), it seems highly unlikely that these changes would all be reversed during the relatively short (i.e., 9 min) recording periods when perfusate oxygen levels were not zero.
Beyond the issues noted above for the isolation and test conditions, a limitation of the whole cell patch-clamp technique is the disruption of the native intracellular milieu by the pipette solution. This disruption affects intracellular ion balance and buffering capacity, interferes with normal cellular signaling by intracellular pH, second messengers, and covalent modification (47), and leads to deterioration (i.e., rundown) of currents over time (34). At the outset of this study, turtle cardiac myocytes had never been investigated with the whole cell patch-clamp technique. Thus our intracellular and extracellular solutions were modified from established whole cell patch-clamp methodologies for teleost fish (15, 16, 42, 51, 55, 65, 66) but were relevant to the freshwater turtle in terms of ionic composition and pH (see Refs. 17, 27, 28 for detailed description of turtle blood ionic composition and pH). Specifically, pipette pH was set to 7.4 at 21°C and was 7.6 at 5°C (49, 71) (Stecyk JAW, Bock C, Overgaard J, Wang T, Farrell AP, Pörtner HO, unpublished observations). Furthermore, to minimize changes in the amplitude of ICa, a current particularly sensitive to rundown, throughout the duration of the experimental protocol 5 mmol/l EGTA was included in the pipette solution (23). In the present study, ICa was −5.7 ± 0.5 pA/pF for 21°C-acclimated normoxic turtles, and in all experimental groups, this current remained unchanged for at least 9 min with repeated measurements of ICa (Fig. 1). The magnitude of ICa and the repeatability of its measurement are comparable to recent reports for cardiomyocytes of yellow-bellied turtles using the perforated-patch technique (12), a technique that overcomes some of the concerns with whole cell patch clamp. Therefore, we are confident of our measured currents and that our reported alterations in current densities with acute temperature change, which were all completed within 9 min, are not due to variation in current density over time.
Nevertheless, the possibility exists that our reported current densities differ from the in vivo condition. Clearly, the square voltage-clamp pulses used to characterize INa, ICa, IK1, and IKr do not emulate the change in membrane potential that occurs with an AP. Furthermore, we did not mimic all the changes in intracellular and extracellular environment that accompany prolonged anoxia exposure (for detailed descriptions, see Refs. 17, 27–29, 49, 71) (Stecyk et al., unpublished observations). For instance, adrenergic stimulation of cardiomyocytes likely exists at a tonic level under normoxia and perhaps at an increased level under anoxia because circulating catecholamines are greatly elevated (31, 32, 70). Epinephrine increases the open probability of L-type Ca2+ channels (45) but was not utilized during measurements of ICa. Because this study was a first step in comparing electrophysiological properties of 21- and 5°C-acclimated turtles exposed to normoxia and anoxia, future studies, some of which have already commenced, utilizing physiologically relevant AP pulse-protocols and faithfully mimicking the extracellular changes in pH and epinephrine that accompany prolonged anoxia are logical next steps for research. Finally, it should be noted that the 20-ms prepulse to −120 mV from the holding potential of −80 mV utilized to remove steady-state inactivation of voltage-gated Na+ channels for measurement of INa may not have been sufficient to fully recover channels from inactivation at 5°C (15). This could partially contribute to the small INa reported for cold-acclimated turtles.
Modification of Turtle Heart Electrophysiology by Temperature and Anoxia
The coordinated pumping action of the turtle heart first involves production of APs in pacemaker cells, which set the spontaneous rhythm of cardiac contraction through a synchronized propagation of excitation throughout the atria and ventricle. The generation of the AP and its expression throughout the heart require the integrated activities of a number of sarcolemmal ionic currents. Thus the potential exists for numerous types of cardiac control mechanisms that are intrinsic to the sarcolemma and that could modify cardiac performance in response to a change in whole body blood flow demand, extracellular conditions, and ambient temperature.
The present study focused on the effects of cold temperature acclimation and prolonged anoxia exposure on turtle cardiac APs, and our goal was to elucidate the contribution of sarcolemmal ion channels to the known reduction in cardiac activity. Four key sarcolemmal ion channel currents were studied in ventricular myocytes. INa, a fast inward Na+ current via voltage-gated Na+ channels, is the first current to be activated in atrial and ventricular cells, determines the amplitude and slope of the AP upstroke (10), is linked to excitation-contraction coupling of cardiac myocytes via the sarcolemmal Na+/Ca2+ exchange (3), and allows for subsequent activation of other ion channels involved in AP generation (33). ICa, the inward Ca+ current via voltage-gated L-type Ca2+ channels, is responsible for the plateau phase of the AP (2) and is the predominant source of free intracellular Ca2+ needed to bind to the myofilaments and trigger contraction in the turtle heart (11, 12). IK1, an outward K+ current via inward-rectifier K+ channels, is primarily responsible for maintaining a stable RMP and terminal repolarization of the AP but not thought to be present during AP plateau (35, 48). IKr, a repolarizing K+ current, gradually develops during the plateau phase of the AP, conducts outward current at more positive voltages than the IK1, and thus is important in balancing ICa and contributing to the plateau of the cardiac AP (46).
Modification of Turtle Heart Electrophysiology With Cold Temperature
In vivo cardiac activity of cold-acclimated freshwater turtles is substantially lower than that of warm-acclimated turtles, with a 5- to 15-fold decrease in fH after acclimation to 5°C from 21–22°C driving similar reductions in Qsys and POsys (19, 57, 59). Similarly, spontaneous fH is six to seven times lower in in vitro heart preparations from 5°C-acclimated turtles than in 21°C-acclimated turtles (56). The present findings not only confirm these findings for spontaneous fH but also indicate that, at least for ventricular tissue, cold temperature can induce changes in the density of sarcolemmal ionic currents in response to direct temperature effects and acclimation. These changes are presumed to serve to alter cardiac AP characteristics and contribute to the depression of cardiac activity in vivo.
To summarize these temperature effects for normoxic turtles, 5°C-acclimated turtle heart exhibited depolarization of RMP by 18–26 mV, 4.7- to 6.8-fold decreases in AP upstroke rate and a prolongation of APD by 4.2- to 4.9-fold in all cardiac tissues examined (Figs. 2, 3, A and C, 4, A and C, 5, A and C). In the ventricle, the increased RMP and APD are consistent with the 50% reduction of IK1 conductance (Fig. 7C), whereas the decreased AP upstroke rate is consistent with the sevenfold reduction in peak density of INa (Fig. 7A). The 13-fold reduction in ventricular ICa of 5°C-acclimated turtles (Fig. 7B) is consistent with reductions in twitch force and time to peak force of 5°C-acclimated turtle ventricular tissue compared with that shown in ventricular tissue from 21°C-acclimated turtles (41).
By making additional measurements of ventricular sarcolemmal current densities after acutely switching the temperature to a common temperature of 11°C, we found that direct temperature effects could be clearly dissected from those due to cold temperature acclimation. Specifically, we found that INa decreased (also reflected in the decreased AP upstroke rate of 5°C-acclimated turtles) predominantly as a result of a direct temperature effect (Figs. 5, A and C, and 8A), whereas density of functional L-type Ca2+ channels was downregulated as part of the cold acclimation (Fig. 8B). These findings contrast in one respect with the effect of cold acclimation in anoxia-tolerant crucian carp (Carassius carassius), where functional Na+ channels were reduced (15), but share a common response to cold acclimation of active downregulation of functional L-type Ca2+ channels (67). It remains to be clarified whether the cold acclimation-induced change in ventricular ICa for turtles is due to temperature-dependent changes in channel phosphorylation, transcription, translation, rate of protein degradation, or trafficking of channels to the sarcolemmal membrane. Furthermore, we found that density of functional inward rectifier K+ channels was upregulated with cold acclimation to partially compensate the negative direct effect of cold temperature on IK1, despite an overall decrease in IK1 conductance (Fig. 8C). This active regulation of inward-rectifier K+ channel density with cold temperature acclimation is consistent with the indication from AP recordings that the increase in RMP to less negative values with cold acclimation is not solely a result of direct temperature effects (Table 1). Again, the possible mechanisms underlying the cold acclimation-induced change in ventricular IK1 and subsequently RMP, such as changes in channel phosphorylation, transcription, translation, rate of protein degradation, or trafficking of channels to the sarcolemmal membrane, remain to be clarified. However, it is tempting to speculate that the known suppression of turtle cholinergic cardiac tone with cold acclimation (20) plays an important role. In rainbow trout atrial myocytes, acetylcholine activates inwardly rectifying K+ currents and stabilizes RMP at more negative voltages (38). Therefore, if a similar phenomenon exists for turtle cardiomyocytes, it is conceivable that the absence of turtle cardiac cholinergic tone with cold acclimation allows for the decreased IK1, the less negative RMP, and the potential for control of IK1 by other mechanisms.
The reduced ventricular sarcolemmal ion currents and the prolongation of cardiac APs of 5°C-acclimated turtles compared with 21°C-acclimated turtles are consistent with the concept of inverse thermal compensation as a strategy to cope with cold temperature. Because of the inhibitory effect of cold temperature on rates of physiological processes, ectothermic vertebrates exhibit a variety of strategies to cope with the cold. Some ectotherms show physiological compensation that allows for the continuation of an active lifestyle at cold temperature. At the cardiac level, established compensatory changes include increased relative ventricular mass (14), increased myofibrillar ATPase activity and decreased refractoriness of the heart (1), proliferation of the sarcoplasmic reticulum (5), modulation of Ca2+ cycling (50–53), increased INa (15), and alterations in K+ conductances that shorten APD (42, 66). However, for a freshwater turtle that becomes inactive during a prolonged period of winter anoxia, physiological processes must be primed to conserve fuel and make compensatory changes maladaptive (22). Accordingly, these organisms reduce activity, routine metabolic rate, and subsequently cardiac activity with cold exposure in anticipation of winter anoxia conditions (18, 26). Similarly, cold-acclimated crucian carp reduced fH (37, 64), reduced the rate of cardiac contraction (61), decreased the maximal conductance of INa by 4.4-fold (15), and decreased the peak ICa density by 6.1-fold (67). Therefore, similar to the findings for crucian carp, the reduced peak density of 5°C-acclimated turtle ventricular INa and ICa as well as inward slope conductance of inward-rectifier K+ channels suggest inverse thermal compensation at the electrophysiological level, which would provide for energetic savings through reducing the cost of ion pumping, one of the largest energy-consuming processes of cells (22). In principle, decreased INa and ICa would reduce demands on the Na+-K+-ATPase, which extrudes a proportion of the Na+ that enters the myocyte during AP upstroke and also via the sarcolemmal Na+/Ca2+ exchange. Furthermore, the current required to trigger an AP is likely to be less with a smaller IK1 and less negative RMP, making myocytes more readily excitability at cold temperature.
Nevertheless, still needed to confirm these suggestions are investigations of direct and acclimatory temperature effects on INa and ICa with pulse protocols representative of physiological APs and in-depth examination of Na+ and Ca2+ channel activation/inactivation kinetics. This is because of the possibility that the prolonged AP and slower activation/inactivation kinetics at cold temperature could allow channels to be open longer, resulting in an increase in total charge transferred despite the decreased peak densities. The possibility is certainly in evidence for atrial myocytes of rainbow trout (Oncorhynchus mykiss), a cold-active species that exhibits positive thermal compensation; the charge carried by ICa, although temperature dependent with square pulses, becomes temperature independent when stimulated with physiologically relevant AP pulse protocols (51). Of course, the reductions in peak INa and ICa densities with cold acclimation in the turtle are much larger than the proportional prolongation of APD, which suggests that these reductions should indeed reduce total charge transferred and contribute to energy conservation. Furthermore, tonic adrenergic stimulation, which likely occurs in vivo, influences ICa (45), and certainly plays a role in cold rainbow trout (54), could be important during cold acclimation in turtles. Future studies investigating the role of epinephrine on ICa would thus be very insightful. Similarly, future investigation on the effect on temperature on Na+/Ca2+ exchanger current is needed to clarify the apparent juxtaposition of decreased ICa and prolonged ventricular APD at 5°C and to determine whether reversed Na+/Ca2+ exchanger current maintains the long AP in the cold.
A surprising discovery in the present study is that IKr was not a predominant current in turtle ventricular myocytes under any experimental condition (Figs. 7D, 11D, and 12D). In contrast, IKr is the predominant repolarizing current in mammalian (46) and fish cardiomyocytes. In fact, cold acclimation markedly increases IKr in rainbow trout (66), and in the burbot (Loto lota), a cold stenothermic fish, IKr is much larger than IK1 (55). This interspecific difference in IKr does not appear to be related to the different cold survival strategies among these species because IKr is present in the crucian carp and is upregulated with cold acclimation (M. Vornanen, personal communication). Whether this phenomenon is unique to turtles remains to be determined.
Finally, the longer APD90 and APD100 of atrial tissue, but not ventricular tissue, of 5°C-acclimated turtles when acutely warmed (Fig. 6, B and D) suggest that the density of K+ currents and/or the effect of cold acclimation on K+ currents differs among cardiac tissue types. A similar phenomenon was reported for rainbow trout where substantial differences in IK1 exist between atrial and ventricular myocytes (66). Again, future studies are needed to clarify this possibility.
Given these important direct and acclimation temperature effects, we can return to the quantitative issue of how these effects might account for the depression of cardiac activity in cold-acclimated turtles. Even the longest tissue APD100, i.e., 3,559.0 ms measured in the ventricle, would allow a fH of ∼17 beats/min if no refractory period existed. Therefore, APD is unlikely to represent a major restriction on the spontaneous fH of ∼5 beats/min. Because cholinergic cardiac inhibition is not involved in the depression of cardiac activity with cold acclimation (20), the decrease in contraction frequency must be due to changes in cardiac refractoriness and other mechanisms such as intercellular electrical coupling and/or pacemaker mechanisms. In ventricular tissue of the anoxia-tolerant crucian carp, the ventricular refractory period increases by sixfold with cold acclimation (61). If refractory period of ventricular tissue increased in cold-acclimated turtles by an amount similar to that observed with crucian carp, an intrinsic fH of ∼4–5 beats/min at 5°C (Ref. 56; present study) could be fully accounted for. Indeed, depolarization of RMP at 5°C (Figs. 2 and 5, A and C) should theoretically result in slower recovery of voltage-gated Na+ channels from inactivation and an increased refractory period. However, unless properties of turtle voltage-gated Na+ channels are different from those of fish and mammals, the depolarized RMP would inactivate practically all voltage-gated Na+ channels. Future studies are needed to resolve these various possibilities.
Modification of Turtle Cardiac Electrophysiology With Prolonged Anoxia Exposure
In contrast to the large effects of temperature on AP shape and duration, prolonged anoxia exposure resulted in few and only small changes in cardiac APs. Nevertheless, the 47% increase in ventricular APD100 with prolonged anoxia exposure at 21°C (Fig. 5, A and B) closely matches the reduction of ventricular contraction rate of ∼30%. Thus, for warm-acclimated turtles, a prolongation of APD is proportional to the reduction in spontaneous cardiac contraction frequency with anoxia. At 5°C, 14 days of anoxia exposure did not affect AP characteristics of any cardiac chamber (Figs. 2, 3, C and D, 4, C and D, and 5, C and D). Therefore, the resetting of intrinsic fH with prolonged anoxia at cold acclimation temperatures (Ref. 56; present study) results primarily from either a marked prolongation of the pacemaker potential, increased contraction and relaxation times, increased muscle refractoriness, or some combination. However, time-to-peak twitch force and time-to-relaxation were similar at 5°C for normoxia- and anoxia-acclimated turtle ventricular strips (41). Thus intrinsic mechanisms related to increased refractoriness of the heart, modification of intercellular electrical coupling, and modification of pacemaker mechanisms and/or other extrinsic modifiers of sarcolemmal ion currents must account for the reduction in intrinsic fH with prolonged, cold anoxia exposure. The present findings exclude the prospect of acidosis as one of these possibilities because exposure of anoxia-acclimated heart preparations to an acidosis equivalent to the in vivo situation (i.e., pH of 7.55 at 21°C or pH of 7.25 at 5°C) did not produce any marked, chamber-independent alterations of APs (Fig. 3, B and D, 4, B and D, and 5, B and D).
Some changes in ventricular sarcolemmal currents were induced by prolonged anoxia exposure without affecting APs. For instance, peak INa density doubled with 6 h of anoxia exposure at 21°C but not at 5°C (Fig. 11A). This finding suggests that an upregulation of functional voltage-gated Na+ channels with anoxia in warm-acclimated, but not cold-acclimated turtles is needed to maintain myocyte excitability and perhaps compensate the depressive effect of an increased extracellular K+ concentration (17, 27, 28) on INa. However, the temperature-dependent effect of anoxia exposure on INa may be due to the temperature-dependent effect of prolonged anoxia on IK1 (Figs. 11C and 12C). The ability of INa to depolarize the membrane is dependent on repolarizing currents such as IK1 that overlap INa at the voltage range of AP onset and thus decrease the net depolarizing current (13). Thus the reduced IK1 density and conductance with a 14-day anoxic exposure at 5°C would mean that less Na+ current is needed to trigger an AP. Consequently, an increased INa with anoxia at this temperature would not be necessary. In contrast, the lack of change in IK1 density with prolonged anoxia at 21°C seems to necessitate an increased INa to maintain myocyte excitability.
The reason why changes in current densities did not affect AP shape for anoxia-acclimated turtle hearts is unclear. Too low a statistical power to distinguish minor differences in AP shape is a possibility. For instance, the final repolarization of 5°C anoxia-acclimated ventricular APs appears to be slightly prolonged (but not statistically significant) compared with that of APs from 5°C normoxia-acclimated ventricles (Fig. 5, C and D) and consistent with reduction in IK1 with prolonged anoxia at 5°C. Another difference is that saline solutions utilized with whole heart preparations contained 1 nM epinephrine, whereas no epinephrine was present for the recording of current densities. Finally, the discrepancy could be due to the prolonged time between myocyte isolation and current recordings for anoxia-acclimated myocytes, which occurred due to the extreme difficulty in obtaining reliable recordings. Even so, our density findings are consistent with previous studies. For example, the constancy of peak ICa density with a 14-day anoxic exposure (Fig. 12B) is consistent with the lack of change in twitch force, time-to-peak force, and relaxation time of ventricular tissue from 5°C anoxia-acclimated turtles compared with 5°C normoxia-acclimated animals (41).
Perspective: Prevalence of Channel Arrest in the Turtle Heart
Prolonged anoxia tolerance requires a matching of ATP demand to the reduced ATP supply available from anaerobic metabolism. Channel arrest is a proposed mechanism through which ATP supply and ATP demand could be matched during anoxia through the reduction in the density and/or activity of ion channels to decrease the energetic cost of ion pumping (22). Such a strategy has been demonstrated in freshwater turtle brain and liver, where Na+, K+, and Ca2+ channel activity are all downregulated with anoxia (4, 6, 7, 44). Given the profound reduction in turtle cardiac activity with prolonged anoxia and the high demand for ATP to support cardiac contraction, the turtle heart would appear to be an ideal, but previously uninvestigated tissue to study channel arrest.
For warm-acclimated turtles, present findings indicate that channel arrest during prolonged anoxia exposure is not a ubiquitous means of energy conservation. For the ventricle, there were no significant changes in ventricular ICa, IK1, and IKr densities (Fig. 11, B–D). Similarly, inward slope conductance of inward rectifier K+ channels was unaffected by prolonged anoxia at 21°C. Furthermore, the doubling of INa with anoxia exposure at 21°C (Fig. 11A) directly opposes the concept of cardiac channel arrest.
However, our findings for ventricular INa and IK1 of 5°C-acclimated anoxia exposed turtles are consistent with the channel arrest hypothesis. First, the upregulation of INa that occurred with 6 h of anoxia exposure at 21°C did not occur with a 14-day anoxic exposure at 5°C. Theoretically, because a proportion of the Na+ that enters the myocyte during AP upstroke is actively extruded from the cell, the lack of upregulation of INa with anoxia at 5°C could lead to reduced demands on the Na+-K+-ATPase and conserve ATP during anoxia. Second, inward IK1 density was reduced by 33% at 120 mV and 18% at −100 mV, and inward slope conductance of inward-rectifier K+ channels diminished by almost half with prolonged anoxia at 5°C (Fig. 12C). In principle, the downregulation of IK1 density and conductance with prolonged anoxia could also serve to limit K+ leakage and lower ATP demand. Inward-rectifier K+ channels allow for continuous K+ efflux from resting cardiac myocytes and also contribute to phase 3 repolarization of the cardiac AP (46). Therefore, during both diastole and systole, IK1 creates a K+ leakage pathway across the sarcolemma and consequently places demands on the Na+-K+-ATPase. Reducing this K+ leak current would thus conserve ATP.
As a point of comparison, there is no evidence for cardiac channel arrest in crucian carp during prolonged, cold anoxia exposure. IK1 conductance, inward-rectifier K+ channel activity, and the number of Ca2+ channels remain unchanged in the crucian carp heart with prolonged anoxia at 4°C (42, 67). However, crucian carp do not exhibit the massive reduction in cardiac activity that is associated with prolonged anoxia in the turtle. In fact, the crucian carp can maintain cardiac activity near normoxic levels for at least 5 days in the complete absence of oxygen (58) and the normoxic cardiac ATP demand is likely within the cardiac glycolytic capability (9). Furthermore, unlike the turtle, no channel arrest occurs in the brain of crucian carp during anoxia exposure (30, 68). This difference in the utilization of channel arrest to conserve energy during anoxia between the turtle and crucian carp is most likely related to the differing anoxia-survival strategies exhibited by these two organisms (9, 36). Briefly, in contrast to the turtle, which enters a comatose-like state during anoxia, crucian carp do not become comatose (39) and continue to swim, albeit at a reduced level compared with that shown at normoxia (40).
In summary, we compared cardiac APs from spontaneously contracting whole heart preparations and INa, ICa, IK1, and IKr of ventricular myocytes obtained from either 21- or 5°C-acclimated, normoxia-, or anoxia-exposed turtles. Our results revealed that both direct and acclimatory cold temperature effects modify turtle cardiac electrophysiology, which serves to decrease cardiac activity with cold acclimation and also preconditions the heart for winter anoxia conditions. Specifically, exposure to cold results in an extensive prolongation of cardiac APDs. Furthermore, decreased peak densities of INa and ICa and decreased conductance of IK1 will conserve ATP by reducing the cost of ion pumping. In contrast, prolonged anoxia exposure at 5°C had only few changes on cardiac APs and ventricular whole cell ion current densities. This finding contrasts the effect of prolonged anoxia at 21°C, where an increase in ventricular APD is proportional to the decrease in spontaneous fH and indicates that the resetting of intrinsic fH to a reduced level that occurs with prolonged anoxia exposure in the turtle involves mechanisms other than an increase in cardiac cycle length. Nevertheless, present findings do suggest the occurrence of some form of channel arrest in turtle cardiac tissue during prolonged anoxia exposure at cold temperature.
This research was supported by Natural Sciences and Engineering Research Council of Canada research grants to A. P. Farrell and J. A. W. Stecyk, a University of British Columbia graduate fellowship to J. A. W. Stecyk, and Research Council of Academy of Finland funding to M. Vornanen.
We give special thanks to Dr. Holly Shiels for advice and innumerable recommendations and to Jaakko Haverinen for instruction on how to record APs.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
- Copyright © 2007 the American Physiological Society