Ca2+ cycling in cardiomyocytes from a high-performance reptile, the varanid lizard (Varanus exanthematicus).

The varanid lizard possesses one of the largest aerobic capacities among reptiles with maximum rates of oxygen consumption that are twice that of other lizards of comparable sizes at the same temperature. To support this aerobic capacity, the varanid heart possesses morphological adaptations that allow the generation of high heart rates and blood pressures. Specializations in excitation-contraction coupling may also contribute to the varanids superior cardiovascular performance. Therefore, we investigated the electrophysiological properties of the l-type Ca(2+) channel and the Na(+)/Ca(2+) exchanger (NCX) and the contribution of the sarcoplasmic reticulum to the intracellular Ca(2+) transient (Delta[Ca(2+)](i)) in varanid lizard ventricular myocytes. Additionally, we used confocal microscopy to visualize myocytes and make morphological measurements. Lizard ventricular myocytes were found to be spindle-shaped, lack T-tubules, and were approximately 190 microm in length and 5-7 microm in width and depth. Cardiomyocytes had a small cell volume ( approximately 2 pL), leading to a large surface area-to-volume ratio (18.5), typical of ectothermic vertebrates. The voltage sensitivity of the l-type Ca(2+) channel current (I(Ca)), steady-state activation and inactivation curves, and the time taken for recovery from inactivation were also similar to those measured in other reptiles and teleosts. However, transsarcolemmal Ca(2+) influx via reverse mode Na(+)/Ca(2+) exchange current was fourfold higher than most other ectotherms. Moreover, pharmacological inhibition of the sarcoplasmic reticulum led to a 40% reduction in the Delta[Ca(2+)](i) amplitude, and slowed the time course of decay. In aggregate, our results suggest varanids have an enhanced capacity to transport Ca(2+) through the Na(+)/Ca(2+) exchanger, and sarcoplasmic reticulum suggesting specializations in excitation-contraction coupling may provide a means to support high cardiovascular performance.

for most ectothermic vertebrates, resting and maximal cardiovascular performance is severalfold lower than a similarly sized mammal or bird (11, 29). However, certain ectotherms have heart rates, cardiac outputs, and arterial blood pressures that approach those of mammals (29). One such species, the varanid lizard, Varanus exanthematicus, has one of the largest aerobic capacities among reptiles, with maximal rates of oxygen consumption that are twice that of other lizards of comparable sizes at the same temperature (3, 4, 19).

To support this superior aerobic capacity, varanid lizards have a host of cardiac morphological specializations that permit the generation of high heart rates and blood pressures. For example, the varanid ventricular wall is thicker than most ectotherms, and contains a significant compact layer and coronary supply (54). The heart is unique among most noncrocodilian reptiles for possessing a well-developed muscular ridge that facilitates a functionally divided heart during systole, allowing for high “mammalian-like” systemic blood pressures, while pulmonary pressure remains low (8, 53–56). Accordingly, maximal heart rates [120 beats/min (30, 52)] and blood pressures [7–14 kPa (8)] of varanid lizards greatly exceed more sedentary reptiles, such as turtles, snakes [except pythons (51)], and other lizards [∼50 beats/min and 4 kPa (13, 14, 52)].

Specializations in excitation-contraction (E-C) coupling at the level of the isolated cardiac myocyte may also underlie the varanid lizard's superior cardiac performance (16). High resting and maximal heart rates in mammals have been associated with enhanced expression of an intracellular store of Ca2+, the sarcoplasmic reticulum (SR) (22). In most ectothermic vertebrates, the SR is poorly developed and the amount of Ca2+ used for contraction originating from the SR is negligible (10, 16, 23, 37, 45). However, highly active species of fish with high heart rates and blood pressures, such as tuna, are more SR dependent than sedentary species (17, 28, 36, 38, 42). Similarly, varanid lizard cardiac muscle is more sensitive to SR inhibition compared with less-active species of reptiles, such as freshwater turtles (16). Thus, enhanced SR Ca2+ cycling may play a significant role in the development of high heart rates and blood pressures in ectotherms.

In addition to an increased dependence on intracellular stores of Ca2+, active ectothermic species may have an enhanced capacity to cycle Ca2+ across the sarcolemmal membrane (35, 44). In most ectotherms, the majority of Ca2+ necessary for contraction is extracellular in origin, regardless of activity level (1, 10, 16, 24, 37, 45, 49). In fish and turtles, extracellular Ca2+ crosses the sarcolemmal membrane primarily through the l-type Ca2+ channel, although in some species the Na+/Ca2+ exchanger (NCX) can also contribute significantly (15, 16, 25, 27, 46–48). Aside from directly contributing activator Ca2+, the l-type Ca2+ channel and the NCX may also serve to trigger the release of Ca2+ from the SR (26) and contribute to the plateau of the cardiac action potential (AP) (47). Thus, increased sarcolemmal Ca2+ cycling has the potential to positively affect many pathways in E-C coupling and may provide a means to enhance heart function. Indeed, among teleosts the density and kinetics of the l-type Ca2+ channel current (ICa) is greater in bluefin tuna atrial cardiomyocytes compared with the less-active pacific mackerel (35), and high densities of the l-type Ca2+ channel have been recorded in ventricular muscle of another active teleost, the skipjack tuna (44).

We have previously shown intracellular Ca2+ transients (Δ[Ca2+]i) from turtle ventricular myocytes under steady-state conditions at room temperature are insensitive to SR inhibition, and contraction is supported exclusively by sarcolemmal pathways (15). In the present study we investigate the hypothesis that these Ca2+ cycling pathways are enhanced in varanid lizard ventricular cardiomyocytes, thereby contributing to this species' superior cardiovascular performance. To this end, we have used fluorescent microscopy and electrophysiology to quantify the amount of Ca2+ used for contraction that originates from the SR, NCX, and l-type Ca2+ channel. Furthermore, because the morphology of cardiomyocytes have important implications on E-C coupling, we have used confocal microscopy to visualize lizard ventricular myocytes and make morphometric measurements that can be compared with other vertebrate species.

METHODS

Animal Origin and Care

Savannah monitor lizards, V. exanthematicus (body mass range, 67–805 g; heart mass range, 0.29–1.01 g; n = 18) were obtained from Monkfield Nutrition (Hertfordshire, UK). Although this is a large range in body mass, cardiomyocyte size does not scale with body size (21), and unlike mammals, body mass has little effect on heart rate in ectotherms (12). The mean cell capacitance from the smaller animals (body mass, 67 ± 3.3 g; heart mass, 0.29 ± 0.02 g) was 42.2 ± 6.2 pF (n = 16 cells). The mean cell capacitance from the larger animals (body mass, 805 ± 34.2 g; heart mass, 1.01 ± 0.20 g) was 41.2 ± 2.3 pF (n = 55 cells). Lizards were held in 1 × 0.5 × 0.5 m vivariums maintained at 25–30°C by heating lamps allowing for behavioral thermoregulation. Animals had easy access to water and were fed cat food daily. To limit variability of results due to the metabolic status of the animals, all lizards were removed from their enclosure at the same time (9 AM) on the day of experimentation prior to feeding.

Solutions

The isolation solution contained (in mM) 100 NaCl, 10 KCl, 1.2 KH2PO4, 4 MgSO4, 50 taurine, 20 glucose, and 10 HEPES, with pH adjusted to 6.9 via KOH. For enzymatic digestion, 1.5 mg/ml collagenase (type 1A), 1 mg/ml trypsin (type IX), and 1.5 mg/ml fatty acid-free BSA were added to the solution. The composition of the extracellular solution used for recording Δ[Ca2+]i was (in mM) 150 NaCl, 5.4 KCl, 1.5 MgSO4, 0.4 NaH2PO4, 2 CaCl2, 10 glucose, and 10 HEPES. The pH was adjusted to 7.7 with NaOH. For electrophysiological measurements, KCl was replaced with equimolar CsCl to inhibit K+ channels and the pH adjusted to 7.7 via CsOH. For whole cell voltage clamp electrophysiological measurement of NCX current (INCX) the pipette solution contained (in mM) 130 CsCl, 1 MgCl2, 5 Mg ATP, 5 Na2-phosphocreatine, 10 HEPES, 15 tetraethylammonium chloride, and 0.03 Na2 GTP. The EGTA concentration was 0.025 mM. The pH was adjusted to 7.2 with CsOH. The inclusion of tetraethylammonium chloride and Cs+ abolished all K+ currents. ICa was characterized using the perforated patch configuration of the voltage clamp technique. For this, 240 μg/ml amphotericin was added to the pipette solution described above. To be certain that cells were indeed perforated and not in the whole cell configuration, a high concentration of CaCl2 (10 mM) was also included in the pipette solution. All drugs were purchased from Sigma-Aldrich (St. Louis, MO).

Isolated Myocyte Preparation

Ventricular myocytes were obtained by adaptation of isolation protocols previously described for turtles (15). Our protocol was approved by and all procedures were made in accordance with United Kingdom Home Office regulations. Briefly, lizards were decapitated and the heart was excised. A cannula was inserted into the right aortic arch and advanced into the ventricle for perfusion. The heart was perfused in a retrograde manner for 20 min at room temperature, first with isolation solution to clear the heart of blood and to stop the heart contracting and then with proteolytic enzymes (BSA, collagenase, and trypsin) for an additional 40 min. Ventricular tissue was separated from the atria, cut into small pieces with scissors, and placed in the enzyme solution and shaken at 34°C (Grant OLS200 water bath shaker). To check for viable cardiomyocytes, aliquots of tissue suspension were taken every 5 min and placed under a microscope. Once viable cardiomyocytes were observed, the suspension was removed from the shaker, left to settle, and resuspended in fresh isolation solution. Healthy viable cells were obtained after 15–20 min of heating and shaking. Cardiomyocytes were agitated gently, filtered through a nylon mesh, and stored in fresh isolation solution at room temperature for up to 8 h. In some experiments, the initial perfusion and digestion were conducted at 30°C rather than at room temperature. When this protocol was used, the tissue did not require the subsequent shaking step in the water bath, and viable cells were achieved after 20 min of enzyme perfusion followed by gentle agitation of the tissue in fresh isolation solution. While this latter technique (requiring higher temperature) may have given a higher yield of cardiac myocytes, the viability, appearance, and results obtained from these cells were similar to the former technique.

Myocyte Morphometrics

To obtain measurements of myocyte dimensions, cells were imaged using confocal microscopy. The sarcolemmal membrane of the ventricular myocytes was visualized by loading cells for 10 min with the lipophilic fluorescent indicator di-8-ANNEPS (5 μM; Molecular Probes). Cells were then resuspended in fresh isolation solution and imaged using a laser scanning confocal microscope (Leica, Germany) with 488 nm excitation light and detection at > 505 nm. Consecutive plane scans (x-y) were made through the cell to make a three-dimensional model (z stack), from which cell length, width, depth, and volume were calculated using Zeiss LSM 5.0 software (see Table 1).

Table 1.

Morphometric measurements of varanid lizard ventricular cardiomyocytes

Cell Length, μm	Cell Width, μm	Cell Depth, μm	Cell Capacitance, pF	Cell Volume, pl	Ratio, pF/pl
151.2±23.8	5.9±0.9	5.6±0.4	41.2±2.3	2.3±0.1	18.2

Data are means ± SE, n = 5 cardiomyocytes. Cell length, width, and depth was measured by analysis of confocal microscopic images of cardiomyocytes. Cell volume was derived from cell capacitance.

Δ[Ca2+]i Measurements

Isolated ventricular myocytes were incubated in 5 μM Fura-2 AM for 15 min at room temperature and then washed by dilution and left for 20–30 min for deesterification. Cells were then transferred to a perfusion bath mounted on an inverted microscope (Nikon Eclipse) and perfused for 2–3 min in extracellular solution. The cells were alternately excited at 340 and 380 nm with a xenon arc-lamp and monochromator and 510 nm emission detected with a photomultiplier (Optoscan Fluorescence System, Cairn Research, Kent, UK). Amplifier outputs were digitized (Digitdata 1440; Axon Instruments, Sunnyvale, CA) and analyzed (PCLAMP 10; Axon Instruments). Cells were field stimulated at 0.2 Hz, and recordings were made before and after 6 min of perfusion with SR inhibitors ryanodine (10 μM) and thapsigargin (1 μM). Cells were not field stimulated during the drug application. All Δ[Ca2+]i experiments were performed at room temperature (∼21°C).

The Fura dye was calibrated as previously described for trout cardiomyocytes (41). Briefly, Fura-loaded cells were perfused with extracellular solution containing 2 μM rotenone, 5 μM carbonyl cyanide m-chlorophenyl-hydrazone, 5 mM sodium iodoacetate, and the Ca2+ ionophore 4-bromo A23187. For maximum fluorescence ratio signal, the solution also contained a total of 2 mM CaCl2. For minimum fluorescence ratio signal, the solution was nominally Ca2+-free and contained 10 mM EGTA. All ratiometric data were converted to Δ[Ca2+]i using Kd = 336 nM and the equation of (20).

Electrophysiological Measurements of Sarcolemmal Currents

Samples of cardiomyocytes were added to the recording chamber and left to settle and attach to the bottom. Cells were perfused with extracellular solution at a rate of 1–2 ml/min, and all experiments were performed at room temperature (∼21°C). For l-type Ca2+ channel and NCX measurements, the perforated patch and whole cell voltage clamp techniques were used, respectively. Experiments were performed using an Axopatch 200B amplifier (Axon Instruments) with a CV 203BU headstage (Axon Instruments). Patch pipettes were pulled with borosilicate glass (Harvard Apparatus, Holliston, MA) and had a resistance of 2.4 ± 0.02 MΩ (perforated patch configuration, n = 35) and 2.8 ± 0.08 MΩ (whole cell configuration, n = 16) when filled with pipette solution. Once a gigaohm seal had formed, patch pipette resistance (5–6 MΩ) was compensated for. In the whole cell configuration, cell membranes were ruptured by delivering a short electrical pulse (zap) to the membrane. In the perforated patch configuration, resistance (Ra) was monitored using the membrane test function to assess the extent of perforation. Once electrical access to the cell was gained, the cell capacitive currents were compensated for by manually adjusting series Ra and the cell capacitance compensation circuits. Series Ra (perforated patch; Ra = 15.9 ± 2.1 MΩ, n = 35, whole cell; 10.2 ± 1.1, n = 16) and capacitance were measured using the membrane test function of pClamp 9.0 software (Axon Instruments). Ra was monitored throughout experiments to ensure a stable preparation. Signals were analyzed offline using Clampfit 9.0 software (Axon Instruments).

Measurement of ICa.

The voltage clamp waveform protocols for each experiment are provided in the figures. The amplitude of ICa was calculated as the difference between peak inward current and the current at the end of the depolarizing pulse. ICa was normalized to cell area by dividing the amplitude of ICa by the cell capacitance to give pA/pF. To assess the rate of inactivation of ICa, tau fast (τf ), and tau slow (τs), inactivation components were derived by fitting a second-order exponential function to the decaying portion of ICa using the Chebyshev procedure (Clampfit Software; Axon Instruments). Steady-state kinetic parameters were determined by fitting steady-state activation and inactivation data to Boltzmann equations to calculate the half-activating and half-inactivating potential (Vh) and the slope of activation and inactivation (k), as previously described (47). Recovery from inactivation of ICa was assessed by normalizing current amplitude at a constant test pulse (500 ms, −70 to 0 mV) to the constant prepulse value (500 ms, −70 to 0 mV) after various interpulse durations (50–350 ms, −70 mV). The contribution of ICa to total cellular Δ[Ca2+] was calculated from the transferred charges and cell volume. Charge transfer was determined by integrating the inactivating portion of the Ca2+ current for 500-ms square-wave voltage pulses from −70 mV to 0 mV. Cell volume was calculated from the measured cell capacitance (Cm = 41.2 ± 2.3 pF, n = 55) and the surface-to-volume ratio of the cells. The cardiomyocytes were considered to be flat elliptical cylinders with an axis ratio of 1:2 for the elliptical cross section (48). The change in total cellular Ca2+ due to Ca2+ influx through l-type Ca2+ channels was expressed as a function of nonmitochondrial volume (48).

Measurement of the INCX.

The capacity of the NCX to transfer charge is dependent on the transmembrane activities of Ca2+ and Na+ as well as the membrane voltage. We investigated the capacity of the NCX at two voltages; at 0 mV to investigate charge transfer at the plateau of the AP and at 20 mV to investigate charge transfer at the AP overshoot. The voltage-dependence of INCX was measured with a repolarizing voltage ramp (2 s) from 50 mV to −80 mV with 5-s intervals. This experiment was also performed in the absence and presence of KB-R7943 (KBR; 5 μM; Calbiochem) to assess the extent of forward- vs. reverse-mode block of INCX. All NCX protocols were carried out in the presence and absence of 10 mM NiCl2, and INCX was then calculated as the Ni+-sensitive difference current. When measuring INCX, 20 μM nifedipine and a combination of 5 μM ryanodine and 2 μM thapsigargin was added to the perfusate to inhibit ICa and any possible SR Ca2+ release and reuptake, respectively.

Statistical Methods

With the exception of original traces and voltage protocols, data are given as mean values ± SE. The effects of SR inhibition on the amplitude and baseline Δ[Ca2+]i, the decay time constant τ, Ca2+ rise time, and the effects of voltage on (INCX) density were tested with paired t-tests. Effects of KBR inhibition on INCX was tested with a two-way repeated-measures ANOVA. Tests were performed with SigmaStat 3.5 software with a significance level of P < 0.05. Signed Rank Tests were used when the data were not normally distributed. N values in figure legends are for number of cells where the minimum number of animals is five.

RESULTS

Cell Morphology

Dissociation of the lizard heart required longer perfusion times and higher temperatures than those used previously for the turtle and various fish species (15, 27, 35, 46, 48). These differences may be due to the thicker ventricular wall and larger compact layer in varanid lizard ventricle or may relate to the complication of perfusing a three-chambered ventricle with a well-developed muscular ridge (53, 54). Light and confocal microscopy images of isolated lizard ventricular cardiomyocytes are displayed in Fig. 1. Ventricular cardiomyocytes were typically spindle-shaped, being ∼150 μm in length and 6 μm in width and depth (Table 1). When light and confocal images are compared, it is apparent that the sarcomeres of the cardiomyocytes are not associated with T-tubules (Fig. 1). Cardiomyocytes had a small cell volume (∼2 pL), leading to a large surface area-to-volume ratio (18.3; Table 1), typical of ectothermic vertebrates.

Fig. 1.

Morphology of live ventricular cardiomyocytes from the varanid lizard heart. Light (A) and confocal microscopy (B) images of varanid lizard ventricular cardiomyocytes. Scale bar applies to all images. Arrow indicates area where sarcomeres are apparent. Mean morphometric data are provided in Table 1.

To determine the relative importance of the SR to Δ[Ca2+]i in varanid ventricular cardiomyocytes, Δ[Ca2+]i was measured in field-stimulated cardiomyocytes with and without SR inhibition by ryanodine (10 μmol/l) and thapsigargin (1 μmol/l) (Fig. 2). While no effect of the drug application on baseline Δ[Ca2+]i was observed (Fig. 2), SR inhibition significantly decreased the rate of rise of the Ca2+ transient (Fig. 2) and its peak amplitude (Fig. 2), the latter by 40%. The decay constant τ increased with SR inhibition (Fig. 2).

Fig. 2.

Effects of sarcoplasmic reticulum (SR) inhibitors ryanodine (10 μM) and thapsigargin (1 μM) on the intracellular Ca2+ transient (Δ[Ca2+]i) in isolated ventricular cardiomyocytes from the varanid lizard field stimulated at 0.2 Hz and room temperature. A: representative recording. B: baseline intracellular Ca2+ concentration. C: Ca2+ rise slope. D: Tau (τ), the first-order decay constant of cytosolic Ca2+ clearance during relaxation. E: amplitude of the Δ[Ca2+]i. *Statistically significant difference between controls and SR-inhibited treatments (n = 12).

l-type Ca2+ Channel Properties

Characterization of inward currents.

A depolarizing voltage step from −80 mV to 0 mV elicited a fast-inactivating Na+ current (INa) and a more slowly activating and inactivating Ca2+ current (Fig. 3, A and B). As both INa and ICa are activated within the same voltage range, prepulses from −80 mV to −40 mV were applied before each test pulse in all subsequent protocols to fully inactivate INa and experimentally isolate ICa. To be certain that the remaining current originated from l-type Ca2+ channels, we used 50 μM nifedipine, a specific l-type Ca2+ channel blocker, to inhibit ICa (Fig. 3, dotted line).

Fig. 3.

Isolation of l-type Ca2+ current (ICa). A: waveform protocol used to activate Ca2+ channel current (ICa) and Na+ current (INa). B: INa and ICa in a varanid lizard ventricular myocyte (capacitance = 34.9 pF) and the effect of 50 μM nifedipine, a specific blocker of the l-type Ca2+ channel, on ICa. C: ICa current-voltage relations. ICa was normalized to cardiomyocyte capacitance (pF) and expressed as current density (pA/pF). Vm = membrane potential. Values are means ± SE (n = 13).

ICa density, kinetics, and voltage relations.

The current-voltage relationship for lizard ventricular cardiomyocytes is shown in Fig. 3. ICa activated at approximately −40 mV, peaked at 0 mV, and reversed at +60 mV. At peak ICa density (−3.1 ± 0.5 pA/pF), the time constant for τf and τs was 37.4 ± 2.2 ms and 189.2 ± 19.0 ms, respectively (n = 13). Due to this relatively slow inactivation time, charge transfer and, therefore, total Ca2+ influx through l-type Ca2+ channels was particularly high in varanid lizard cardiomyocytes (∼ 64 μmol/l) compared with that observed in other ectotherms and mammals (5, 31, 35, 46, 48) (Table 2).

Table 2.

Characterization of sarcolemmal Ca2+ influx through L-type channels in varanid lizard ventricular cardiomyocytes

Capacitance, pF	ICa Density, pA/pF	Charge Transfer, pC	Charge Density, pC/pF	Δ[Ca2+] Total NonM, μmol/l
41.2±2.3	−3.1±0.5	−11.7±2.2	−0.28±0.04	66.2±9.3

Data are means ± SE, n = 13 cardiomyocytes. Δ[Ca2+] Total NonM, change in total cellular [Ca2+] expressed as a function of nonmitochondrial (NonM) cell volume.

Steady-state activation and inactivation of ICa.

The voltage protocol for measuring steady-state activation and inactivation of ICa and a representative current recording is given in Fig. 4, A and B, respectively. Activation of ICa began positive to −40 mV, and was half maximal (Vh) at −9.5 ± 1.4 mV, while inactivation of ICa, or channel availability, began decreasing positive to −30 mV and was half complete at −24.5 ± 1.0 mV (Fig. 4). The slopes of activation and inactivation (k) were 5.6 ± 0.2 and 3.8 ± 0.1, respectively. At voltages positive to 10 mV, channel inactivation was attenuated, probably due to a reduced driving force and consequently less Ca2+-dependent inactivation (Fig. 4). As a result of overlap between activation and inactivation curves, a window current was evident between −40 and 10 mV. The window current was maximal at approximately −18 mV, where it contributed 4% of maximal conductance (Fig. 4, inset).

Fig. 4.

Steady-state activation and inactivation of ICa in varanid lizard ventricular cardiac cardiomyocytes. A: voltage protocol used to measure steady-state activation and inactivation. B: representative current recording from a varanid ventricular myocyte (capacitance = 56.2 pF) subjected to the voltage protocol given in A. C: mean steady-state inactivation and activation profiles. Values are means ± SE (n = 12). Inactivation (▪) is measured by depolarizing from −70 mV to a test potential for 1 s and then testing the remaining available ICa at 0 mV. Activation (□) is calculated by dividing peak current by apparent driving force [applied membrane potential (Em) − reversal potential (Erev)] according to Ohm's law. Inset: l-type Ca2+ channel window current (product of activation and inactivation at each voltage).

ICa recovery from inactivation.

The voltage protocol for measuring the recovery of ICa from inactivation and a representative current trace is shown in Fig. 5, A and B, respectively. The number of recovered channels increased as the duration between the prepulse and the test pulse was lengthened (Fig. 5). The time constant of recovery from inactivation (τ) was 190.1 ± 21.5 ms, thus at physiologically relevant frequencies of contraction (0.2–1.5 Hz), incomplete restitution of lizard l-type Ca2+ channels is unlikely to occur.

Fig. 5.

Recovery of ICa from inactivation in varanid lizard ventricular cardiomyocytes. A: voltage protocol used to measure recovery from inactivation. B: representative trace from a varanid lizard ventricular myocyte (capacitance = 40.1) subjected to the displayed voltage protocol. C: relationship between current recovery and interpulse duration for ventricular cardiomyocytes. Values are means ± SE, n = 10.

INCX Properties in Lizard Ventricular Myocytes

Characterization of INCX.

A square-wave voltage pulse from −40 mV to 0 mV for 500 ms and then to +20 mV for 500 ms (Fig. 6) gave rise to a maintained outward current, which could be blocked with 10 mM NiCl2, confirming the presence of a Ni+-sensitive Na/Ca2+ exchange current (INCX) (Fig. 6). INCX was integrated to give a measure of charge transfer so that total Ca2+ influx through the NCX could be calculated. Charge transfer was significantly greater at 20 mV (Fig. 6).

Fig. 6.

Stimulus waveform (A) and representative Na+/Ca2+ exchanger current (INCX) (B) from a lizard ventricular myocyte (40 pF) elicited by square depolarizing pulses to 0 and 20 mV (with a prepulse to −40 mV to remove INa). B: calculated total Ca2+ transferred by INCX during a pulse to 0 or 20 mV expressed as μmol/l nonmyofibrillar (see methods). *Significantly more Ca2+ transferred on INCX at 20 mV compared with 0 mV. Values are means ± SE, n = 7.

NCX voltage sensitivity.

The voltage-dependence of INCX was measured using a voltage ramp protocol (Fig. 7, inset). INCX was identified as the Ni+-sensitive current. INCX showed a steep increase with increasing voltage (outward rectification) (Fig. 7), while the inward current increased more slowly during hyperpolariazation. The measured reversal potential of INCX was −10 mV. Using this reversal potential, we calculate an intracellular-free Ca2+ of ∼50 nM. This value is lower than the diastolic value obtained in Fura-2-loaded cells (∼150 nM) and may be due to the fact cells were not undergoing constant stimulation. Certainly a higher intracellular-free Δ[Ca2+] would have shifted the reversal potential of the NCX to more positive voltages.

Fig. 7.

A: current-voltage relationship for INCX in lizard ventricular cardiomyocytes. The current is calculated from the voltage ramp pulse (see inset). •, Ni-sensitive current (10 mM NiCl2); ○, KB-R7943 (KBR)-sensitive current (KBR, 5 μM). *Significant block by KBR (2-way repeated-measures ANOVA, P < 0.05). B: %inhibition of forward and reverse INCX by 5 μM KBR calculated from the KBR-sensitive current as a fraction of INCX at 50 mV on either side of the reversal potential of −10 mV. *Significantly greater inhibition of reverse INCX. All values are means ± SE, n = 8–10 cells.

NCX inhibition by KBR.

We compared the extent of forward- vs. reverse-mode block of INCX with KBR using the voltage ramp protocol (Fig. 7, inset). After recording baseline current (Fig. 7), 5 μM KBR was applied. A steady-state response was achieved within 5 min, and the calculated difference current represented the KBR-sensitive current (Fig. 7). NiCl2 (10 mM) was then added to fully block the NCX and to measure total INCX. From this data the fractional block by KBR was calculated (Fig. 7). KBR significantly decreased reverse mode, but not forward mode, INCX. When measured at a similar driving force KBR inhibited > 50% of reverse-mode INCX (Fig. 7).

DISCUSSION

The varanid lizard heart possesses morphological specializations that allow the generation of high heart rates and blood pressures (8, 53–56). The present study demonstrates that the varanid lizard ventricle is also specialized at the level of the isolated cardiomyocyte. The major findings from this study are that 1) varanid ventricular myocytes are structurally similar to other ectotherms and rely predominantly on sarcolemmal Ca2+ cycling, 2) the varanid NCX has a large capacity to move Ca2+ into and out of the cardiomyocyte, and 3) the varanid SR plays an important role in steady-state E-C coupling. In aggregate, our results suggest that the density and/or kinetics of the SR and NCX in varanid ventricular myocytes are enhanced compared with other ectotherms, which may contribute to this species' superior cardiac performance.

Varanid Cardiac Myocyte Morphology

The structural design of lizard ventricular cardiomyocytes is similar to other ectothermic vertebrates, and reveals a system primed for transsarcolemmal Ca2+ flux. Lizard cardiomyocytes are spindle-shaped and lack T-tubules, which is typical of ectothermic vertebrates (6, 15, 49), and although they are somewhat comparable in length to mammalian ventricular myocytes (lizard, ∼150; rat, ∼141), their width and depth appear much smaller (lizard, ∼6; rat, ∼13) (33). Thus, the surface area-to-volume ratio (18.2) is high compared with mammals (rabbit, 4.6; rat, 6–8) (2, 33), but similar to other ectotherms (trout, 18.2; crucian carp, 19.2; bluefin tuna, 14–17; mackerel, 18–22; turtle, 18.3) (15, 35, 46, 48). This large surface area-to-volume ratio will increase the impact of sarcolemmal Ca2+ transport by reducing the diffusional distance that Ca2+ has to travel to the myofilaments. Moreover, the myofilaments of ectothermic cardiomyocytes lie directly below the sarcolemmal membrane (48). Thus, the ectothermic cardiomyocyte is primed for extracellular Ca2+ cycling, and the morphology of the cardiomyocyte negates the necessity for a T-tubular network.

Varanid Sarcolemmal Ca2+ Cycling

Our results indicate sarcolemmal Ca2+ cycling is the primary source of activator Ca2+ for contraction in varanid lizard ventricular myocytes. This is also the case in the majority of ectothermic vertebrates (1, 15, 16, 25, 26, 35, 46, 48). The peak density at room temperature of varanid ICa (−3.1 ± 0.5 pA/pF) is similar to that found in the turtle, (∼3.2 pA/pF), but appreciably smaller than mammalian ICa (∼10 pA/pF) (32, 39, 46). However, reptilian ICa inactivates more slowly than mammalian ICa (15), allowing for a greater charge transfer over time. Furthermore, the small cell size and large surface-area-to-volume ratio of ectothermic cardiac cardiomyocytes means that the impact of a given Ca2+ influx across the sarcolemma is far greater than in mammalian cardiomyocytes. Thus, when converting ICa to total Ca2+ influx of nonmitochondrial space (66.2 ± 9.3 μmol/l, n = 13), lizard and turtle Ca2+ influx through l-type Ca2+ channels at room temperature is five times that found in adult mammalian ventricular cardiomyocytes (5, 31) and approximately double that of certain fish ventricular cells (35, 46, 48).

In addition to the l-type Ca2+ channel, the NCX appears to play an important role in both the removal and entry of Ca2+ across the sarcolemma. At negative membrane voltages, the varanid ventricular NCX acts as the primary Ca2+ removal pathway, as in other ectotherms and mammalian cardiomyocytes. This is demonstrated by the decay of Δ[Ca2+]i in Fig. 2. When the SR Ca2+ uptake pathway is inhibited, decay is only slowed by ∼25%, suggesting 75% of Ca2+ efflux in varanid ventricular myocytes is supported by the NCX. Compared with turtle ventricular myocytes, which depend exclusively on the NCX to remove Ca2+, the decay constant of varanid Δ[Ca2+]i under SR inhibited conditions (1,093.2 ± 96.8 ms) is faster than the turtle (∼1,350 ms, estimated from Ref. 15), suggesting the varanid NCX has a greater capacity to remove Ca2+. Indeed, varanid NCX density at negative membrane potentials is twofold higher than the turtle (15). Furthermore, at voltages positive to −10 mV, our results suggest the NCX contributes to contraction by entering “reverse mode” (Ca2+ in, Na+ out) during the upsweep of the AP. In fact, total Ca2+ entry via reverse mode NCX in the varanid at 0 mV (179 ± 25 μmol/l) is approximately fourfold greater than values obtained from the turtle or the crucian carp measured under identical experimental conditions (15, 47). In aggregate, these results suggest the density and/or kinetics of the NCX are particularly high in varanid ventricular myocytes, which may contribute to the generation of high heart rates (enhanced rate of Ca2+ removal) and force of contraction (enhanced Ca2+ influx).

Varanid SR Dependence

Ca2+ cycling through intracellular Ca2+ stores appears to be enhanced in the varanid ventricular myocyte with up to 40% of the Ca2+ used for contraction originating from the SR. This is in stark contrast to most ectotherms, including the turtle, where the role of the SR in E-C coupling is small or negligible (10, 15, 16, 23, 37, 45). The decay constant of the varanid Δ[Ca2+]i under control conditions (where the SR is contributing to Ca2+ removal) is ∼30% faster than the SR-independent turtle (15), suggesting the SR may aid in rapid systolic Ca2+ removal and thus facilitate high-contraction frequencies. Consistent with these results, morphological studies show a well-developed SR network in lizards, with a total, free, and junctional SR volume that is more similar to mammals and birds than amphibians or fish (7). Furthermore, isolated ventricular muscle preparations from the varanid lizard exhibit a 15–20% reduction in contractile force when treated with ryanodine (16). The greater effect of SR inhibition on Δ[Ca2+]i in the present study (∼40%) may reflect the use of ryanodine and thapsigargin in combination to inhibit both the Ca2+ release channel and Ca2+ reuptake ATPase. Taken together, these studies suggest both Ca2+ entry and removal in varanid ventricular myocytes can occur through intracellular stores, and this may contribute to high cardiovascular performance in this species.

It should be noted that the contribution of the SR, l-type Ca2+ channel, and NCX during contraction and relaxation of ectothermic cardiomyocytes can change substantially according to many factors, including heart rate, temperature, exercise level, and pH (49). The present study has investigated varanid ventricular myocytes at room temperature stimulated at 0.2 Hz, which lies in the range of frequencies experienced by resting varanid lizards at this temperature (34). However, V. exanthematicus are indigenous throughout most of Africa and the Sahara where temperatures can range from 10–58°C. Furthermore, varanids are highly active predators, and exercise (as with other vertebrates) is associated with high heart rates [up to 120 min in Varanus at 37°C (30)], adrenergic stimulation, and acidosis (9). All of these factors will have profound effects on cardiac function, and the E-C coupling processes that support it (6).

SR dependence in varanid lizard ventricular muscle is greater at lower frequencies of contraction (0.2–0.5 Hz) (16). At present, there is no other information available on the effects of environmental change and physiological status on E-C coupling processes in varanid lizards, although some exists for other species of ectotherms. While acutely cooling cardiac myocytes has a negative impact on l-type Ca2+ channel cycling in ectothermic myocytes, the associated changes in heart rate (and AP duration) tend to offset the negative effects of temperature (42, 43). SR dependence also seems to be temperature sensitive with acute warming increasing SR involvement (37). Alterations in adrenaline levels are known to increase Ca2+ entry via phosphorylation of the l-type Ca2+ channel, and this appears to be temperature sensitive, being greater at cold temperatures (40). Finally, a low pH is known to decrease myofillament Ca2+ sensitivity (18), but the effects of acidosis on other components of the ectothermic E-C coupling cascade have yet to be identified. Obviously, more information is required to make a complete picture of varanid E-C coupling under physiologically relevant conditions. The present study lays the foundation for future work where the roles of the l-type Ca2+ channel, SR, and NCX in varanid cardiac myocytes under different experimental conditions are more fully elucidated.

Perspectives and Significance

To our knowledge, this is the only study that has quantitatively characterized Δ[Ca2+]i from a reptile. While diastolic Ca2+ levels in varanid ventricular myocytes are comparable to other studies on ectotherms and mammals (41, 50), systolic Δ[Ca2+]i amplitude appears to be 10-fold larger than trout atrial myocytes (41) and twice that found in mammalian ventricular myocytes (50). Direct comparison of the present study with mammalian literature is complicated by investigators using different experimental conditions, such as stimulation frequency, type of fluorescent dye, temperature, and concentration of Ca2+. Nevertheless, our results suggest the varanid ventricle may have evolved an E-C coupling system similar to mammalian atrial myocytes, which have a significant SR component while retaining a spindle-like morphology, allowing for greater sarcolemmal Ca2+ transport. This combination provides a robust framework for the transport of Ca2+, and may underlie the varanid heart's ability to generate high frequencies of contraction and contractile force.

GRANTS

This study was funded by The Biotechnology and Biological Sciences Research Council, The Wellcome Trust, and The Anglo-Danish Society.

DISCLOSURES

No conflicts of interest are declared by the author(s).