Culture and adenoviral infection of sinoatrial node myocytes from adult mice

A new method is described for the culture of fully differentiated pacemaker cells from adult mice. Cultured sinoatrial myocytes retain normal morphological and electrophysiological properties. Robust expression of exogenous proteins is achieved by adenoviral infection. The new method extends the experimental approaches that can be used to study cardiac pacemaking.

Abstract

Pacemaker myocytes in the sinoatrial node of the heart initiate each heartbeat by firing spontaneous action potentials. However, the molecular processes that underlie pacemaking are incompletely understood, in part because of our limited ability to manipulate protein expression within the native cellular context of sinoatrial node myocytes (SAMs). Here we describe a new method for the culture of fully differentiated SAMs from adult mice, and we demonstrate that robust expression of introduced proteins can be achieved within 24–48 h in vitro via adenoviral gene transfer. Comparison of morphological and electrophysiological characteristics of 48 h-cultured versus acutely isolated SAMs revealed only minor changes in vitro. Specifically, we found that cells tended to flatten in culture but retained an overall normal morphology, with no significant changes in cellular dimensions or membrane capacitance. Cultured cells beat spontaneously and, in patch-clamp recordings, the spontaneous action potential firing rate did not differ between cultured and acutely isolated cells, despite modest changes in a subset of action potential waveform parameters. The biophysical properties of two membrane currents that are critical for pacemaker activity in SAMs, the “funny current” (I_f) and voltage-gated Ca²⁺ currents (I_Ca), were also indistinguishable between cultured and acutely isolated cells. This new method for culture and adenoviral infection of fully-differentiated SAMs from the adult mouse heart expands the range of experimental techniques that can be applied to study the molecular physiology of cardiac pacemaking because it will enable studies in which protein expression levels can be modified or genetically encoded reporter molecules expressed within SAMs.

NEW & NOTEWORTHY

A new method is described for the culture of fully differentiated pacemaker cells from adult mice. Cultured sinoatrial myocytes retain normal morphological and electrophysiological properties. Robust expression of exogenous proteins is achieved by adenoviral infection. The new method extends the experimental approaches that can be used to study cardiac pacemaking.

each beat of the heart is triggered by a spontaneous electrical depolarization that originates in the sinoatrial node. Sinoatrial node myocytes (SAMs) are highly specialized pacemaker cells that differ considerably from other cardiac myocytes in terms of morphology, function, and protein expression. SAMs fire spontaneous pacemaker action potentials (APs) that are characterized by the presence of a spontaneous depolarization during diastole and a slow, Ca²⁺-mediated upstroke. The mechanisms responsible for the diastolic depolarization and the generation of spontaneous APs are incompletely understood, but the cardiac funny current (I_f) and voltage-gated Ca²⁺ currents (I_Ca) are among the membrane currents that are critical for pacemaker activity in SAMs (26).

Advances in our understanding of the molecular physiology of cardiac pacemaking are limited in part by the lack of experimental techniques for the manipulation of proteins within the unique cellular context of SAMs. Although many insights into pacemaker mechanisms have been gained through experiments using acutely dissociated SAMs from adult animals, these short-term cell preparations are not amenable to studies requiring alterations in protein expression or the use of genetically encoded reporter molecules. Although other cardiac-derived cells, such as neonatal ventricular myocytes and immortalized cardiac cells lines, can contract spontaneously and be maintained in culture for expression studies, these cell types are not suitable for detailed studies of pacemaker mechanisms because they differ from fully differentiated SAMs in numerous aspects, including the spontaneous AP firing rate, the AP waveform shape, and the properties and expression of I_f and other membrane currents (35, 45).

In the present study, we sought to develop a method to maintain fully differentiated adult SAMs in culture in such a manner as to preserve their characteristic properties while permitting expression of exogenous proteins. Here, we present detailed methods describing the isolation, culture, and adenoviral infection of spontaneously active SAMs from adult mice. We have validated this system by showing that morphological and electrophysiological properties of the cultured cells are similar to those of acutely dissociated SAMs and by demonstrating that robust expression of exogenous proteins can be attained within 24–48 h in vitro by adenoviral-mediated gene transfer. This new culture system expands the range of experimental approaches that can be applied to the study of cardiac pacemaking.

METHODS

Sinoatrial myocyte isolation.

Animal procedures were performed according to protocols approved by the Institutional Animal Care and Use Committee of the University of Colorado Denver, Anschutz Medical Campus.

SAMs were harvested from male C57BL/6J mice of 6–12 wk of age (Jackson Laboratories, Bar Harbor, ME) as we have previously described (16, 18, 38, 14). Animals were anesthetized by inhalation of isoflurane and euthanized by cervical dislocation. Hearts were excised, and the ventricles removed. The sinoatrial node, defined as the region bordered by the crista terminalis, the interatrial septum, and the superior and inferior vena cavae (13, 15, 34, 12, 22, 30), was microdissected at 35°C in a heparinized Tyrode's solution containing (in mM) 140 NaCl, 5.4 KCl, 1.2 KH₂PO₄, 5 HEPES, 5.5 D-Glucose, 1 MgCl₂, 1.8 CaCl₂, and 100 USP/ml heparin, pH adjusted to 7.4 with NaOH. Sinoatrial node tissue was cut into three strips, which were enzymatically digested for 10–15 min at 35°C with 3.75 mg Liberase TM (Roche, Basel, Switzerland) and 4.75 U elastase (Worthington Biochemical, Lakewood, NJ) in a modified Tyrode's solution containing (in mM) 140 NaCl, 5.4 KCl, 1.2 KH₂PO₄, 5 HEPES, 18.5 D-glucose, 0.066 CaCl₂, 50 taurine, and 1 mg/ml BSA, pH adjusted to 6.9 with NaOH. Following digestion, tissue was transferred to a modified Kraft-Brühe (KB) solution containing (in mM) 100 potassium glutamate, 10 potassium aspartate, 25 KCl, 10 KH₂PO₄, 2 MgSO₄, 20 taurine, 5 creatine, 0.5 EGTA, 20 glucose, 5 HEPES, and 0.1% BSA, pH adjusted to 7.2 with KOH at 35°C, and SAMs were dissociated by mechanical trituration with a fire-polished glass pipette (∼2 mm diameter) for 5–12 min. Calcium was gradually reintroduced to the cell suspension in 5 steps over the course of 22 min to a final concentration of 1.8 mM. Cells used for acute studies were stored in KB solution at room temperature for up to 8 h before electrophysiological recordings.

Sinoatrial myocyte plating and culture.

SAMs for culture experiments were collected by centrifugation at ∼3,000 rpm and were resuspended in a plating medium, consisting of Media199 (Sigma-Aldrich, St. Louis, MO; No. M4530) supplemented with 10 mM 2,3-butanedione monoxime (BDM; Sigma-Aldrich), 100 U/ml penicillin (GE Healthcare, Pittsburg, PA), 100 mg/ml streptomycin (Hyclone, Logan, UT), and 5% FBS (Hyclone). Cells were plated at an approximate density of 10–15 cells/mm² onto laminin-coated glass coverslips that were prepared immediately before plating by applying mouse laminin (BD Biosciences, San Jose, CA) at 100 ng/ml in PBS for 1 h at 37°C. FBS was used in the plating media to promote cellular adhesion to the cover glass.

SAMs were allowed to settle and adhere to coverslips for 4–6 h. The medium was then changed to a culture medium consisting of Media199 supplemented with 0.1 mg/ml BSA, 10 mM BDM, 10 μg/ml insulin, 5.5 μg/ml transferrin, 5 ng/ml selenium (ITS; Sigma-Aldrich), 100 U/ml penicillin, and 100 mg/ml streptomycin. SAMs were maintained at 37°C in an atmosphere of 95% air/5% CO₂, and culture medium was gently exchanged every 48 h.

Transient transfection and adenoviral transduction of sinoatrial myocyte cultures.

Transient transfections of cultured SAMs with enhanced green fluorescent protein (eGFP) in the pcDNA3 mammalian expression vector (Life Technologies, Grand Island, NY) were attempted on the day of plating using either Xtreme Gene 9 (Roche) or Lipofectamine 3000 (Life Technologies) at cDNA weight (in μg): reagent volume (μl) ratios ranging from 1:3 to 1:6 according to the manufacturers′ protocols and as we have previously reported for transient transfection of CHO and HEK cells (16, 17). However, no GFP fluorescence was observed in the cells subjected to the transfection reagents, even after 72 h.

Recombinant human adenoviruses type 5 expressing either mCherry or eGFP under the control of the mammalian CMV promoter were obtained from Vector Biolabs (Philadelphia, PA; Cat No. 1767 and No. 1060, respectively). Viruses were applied either singly or together at a MOI (multiplicity of infection) of 100 each to SAMs on the day of plating after the cells had settled. Cells were counted immediately before virus application and were incubated with the virus-containing medium overnight (∼12–14 h) before the medium was exchanged for fresh culture medium.

Criteria for identification of sinoatrial myocytes.

SAMs were isolated from the sinoatrial node, which was defined as the region bordered by the crista terminalis, the interatrial septum, and the superior and inferior vena cavae as in previous studies (16, 18, 38, 14, 25, 33). Individual cells were initially identified visually by the presence of spontaneous contractions and by their characteristic morphology, small size, and lack of striations as we have previously described (14, 16, 18, 38). In electrophysiological recordings, SAMs were additionally defined as cells that fired spontaneous action potentials with characteristic AP waveform parameters including a diastolic depolarization phase and a slow upstroke (14) and by the presence of I_f of >100 pA in response to a 1-s voltage step to −120 mV in voltage clamp experiments (see results). All of the acutely isolated and cultured SAMs that were assayed were also found to express HCN4 channel protein in immunocytochemical experiments (see Fig. 5).

Fig. 5.

Expression of HCN4 channels and properties of the “funny current” (I_f) in cultured and acutely isolated SAMs. A: similar HCN4 immunofluorescence intensity and localization in acutely isolated and cultured SAMs. B: similar I_f current density at −150 mV in acutely isolated (n = 7) and cultured (n = 8) SAMs (P > 0.05, t-test). C: similar voltage-dependence of activation in acutely isolated (n = 8) and cultured (n = 14) SAMs. Insets, representative current families; scale bars 500 pA, 500 ms.

Sinoatrial myocyte electrophysiology.

For electrophysiology, either a 100 μl aliquot of acutely dissociated SAMs in suspension or a fragment of glass coverslip bearing cultured SAMs was transferred to a 200 μl recording chamber on the stage of an inverted microscope. Spontaneous contractions were observed in both acutely isolated and cultured cells immediately upon transfer to the recording chamber, which contained Tyrode's solution at 35°C containing (in mM) 140 NaCl, 5.4 KCl, 1.2 KH₂PO₄, 5 HEPES, 5.55 glucose, 1 MgCl₂, and 1.8 CaCl₂, pH adjusted to 7.4 with NaOH. All electrophysiological experiments were performed at 35 ± 1°C, which was maintained by dual platform and in-line perfusion heaters (Warner Instruments, Hamden, CT), and cells were continuously perfused at 1 to 2 ml/min with Tyrode's solution during the recordings. Although cultured cells began beating immediately upon transfer to the Tyrode's solution in the recording chamber, they were perfused with fresh Tyrode's solution for a minimum of 2 min before recordings to remove any residual BDM from the culture medium. In all recordings, the fast component of pipette capacitance was minimized, and the membrane capacitance was estimated with 10-mV test pulses using the membrane test function in Clampex software (Molecular Devices, Sunnyvale, CA).

Recording and analysis of spontaneous APs.

Spontaneous APs were recorded from cultured and acutely isolated SAMs meeting the above criteria using the amphotericin perforated-patch technique in current-clamp mode without current injection. Borosilicate glass patch pipettes had resistances of 1.5–3 MΩ when filled with an intracellular solution composed of (in mM) 135 KCl, 0.1 CaCl₂, 1 MgCl₂, 5 NaCl, 10 EGTA, 4 Mg-ATP, 10 HEPES, and 200 μg/ml Amphotericin-B (Fisher Scientific, Pittsburgh, PA), with pH adjusted to 7.2 with KOH. Amphotericin-B was prepared fresh daily as a 20 mg/ml stock solution in DMSO, and the amphotericin-containing pipette solution was made fresh hourly by diluting an aliquot of the stock solution into the intracellular solution and vortexing for at least 1 min. The final pipette solution containing amphotericin was stored on ice and protected from light.

Access resistance (R_a) was monitored for each cell during perforation using the membrane test function in ClampEx, and recordings were begun only after obtaining a stable R_a of <10 MΩ (usually within 2–3 min). Data were acquired at 5 kHz and low-pass filtered at 1 kHz using an Axopatch 1D or 200B amplifier, Digidata 1322a or 1440a A/D converter, and ClampEx software (Molecular Devices). Spontaneous AP firing rate was measured in the presence of 1 nM isoproterenol (ISO; EMD Millipore/Calbiochem, Billerica, MA) in the bathing Tyrode's solution, which was used to promote stable AP firing in both acutely isolated and cultured SAMs as previously described (6, 14, 16, 18, 38).

APs were analyzed offline using ClampFit software (Molecular Devices). AP firing rates for each cell are reported as the average instantaneous firing rate measured during 15- to 30-s recording windows. AP waveform parameters were calculated for each cell from average waveforms from 5-s recording windows as we have previously described (14). Maximum diastolic potential (MDP) and V_max were defined as the most negative and positive membrane potentials, respectively. Cycle length was defined as the interval between MDPs of successive APs. The take off potential (TOP) was defined as the membrane potential at which the dV/dt reached 10% of its maximum value. The upstroke and repolarization rates were taken as the maximum and minimum values of the first derivative of the AP waveform (dV/dt_max and dV/dt_min respectively). Action potential duration (APD) was defined as the interval between the TOP and subsequent MDP. The early diastolic depolarization rate (eDDR) was estimated as the slope of a linear fit between 10% and 50% of the diastolic duration, and the early diastolic duration was the corresponding time interval. The nonlinear late diastolic depolarization (LDD) phase was described as the time duration between 1% and 10% of dV/dt_max.

Voltage-clamp recordings and analysis.

Following recording of spontaneous APs in current-clamp mode, each cell was then assayed for the presence of I_f in voltage-clamp mode. To be considered I_f positive, cells exhibited a slowly activating inward current of >100 pA in response to a 1-s voltage step to −120 mV from a holding potential of −50 mV. All acutely isolated and cultured cells in this study that fired spontaneous APs were found to be I_f positive.

For amphotericin perforated-patch voltage-clamp recordings of I_f, SAMs were perfused with Tyrode's solution that contained 1 mM BaCl₂ to block K⁺ currents as we have previously described (12, 13, 15, 34). For measurement of I_f following spontaneous AP recordings, cells were perfused for a minimum of 2 min with Tyrode's solution containing BaCl₂ before I_f recordings were begun. I_f current density and activation kinetics were determined from the amplitudes and double-exponential fits, respectively, of 3-s hyperpolarizing voltage steps to −150 mV from a holding potential of −50 mV. The voltage-dependence of I_f was determined from current families elicited by 3-s hyperpolarizing voltage steps ranging from −60 mV to −160 mV in 10-mV increments, from a holding potential of −50 mV. Interpulse intervals were 5–30 s and were triggered manually, once the holding current returned to its original level between successive sweeps. Conductance (G) was calculated from hyperpolarization-activated inward currents according to Ohm's law: G = I/(V_m − V_r), where I is the time-dependent component of I_f, V_m is the applied membrane voltage (corrected for a +14 mV junction potential error), and V_r is the reversal potential for I_f in SAMs (−30 mV) (17). Conductance for I_f was determined from inward currents because the presence of many other currents in mouse SAMs precludes I_f tail current analysis. Conductances were normalized to the maximum value, plotted as a function of test potential, and fit with a Boltzmann equation to determine the midpoint activation voltage (V_1/2) and slope factor (k) for each cell: f(V) = 1/[1 + e^{(V − V_1/2)/k}].

Whole cell Ca²⁺ currents were recorded as we have previously described with an intracellular (pipette) solution consisting of (in mM) 130 CsCl, 1 MgCl₂, 10 HEPES, 10 EGTA, 4 Mg-ATP, and 0.1 Na-GTP, with pH adjusted to 7.2 with CsOH, and extracellular (bath) solution consisting of 130 TEA-Cl, 2 CaCl₂, 1 MgCl₂, 10 4-aminopyridine, and 10 HEPES, with pH adjusted to 7.4 with CsOH. Total I_Ca was elicited by 200 ms depolarizing voltage steps between −80 and +40 mV in 10-mV increments from a holding potential of −90 mV. Applied voltages were corrected for a −9 mV calculated liquid junction potential. Linear components of leak and capacitive current were cancelled using −P/4 subtraction. Individual and average current-voltage plots for Ca²⁺ currents were fit with a modified Boltzmann equation: I = G_max*(V − V_rev)/{1 + exp [(V − V_1/2)/k_G] }, where I is the peak current at a given voltage (V), V_rev is the reversal potential, G_max is the maximum conductance, V_1/2 is the half-maximal activation potential, and k_G is the slope factor. Ca²⁺ current inactivation rates were determined from double exponential fits to currents elicited by voltage steps to −10 mV.

Immunocytochemistry and cell size measurements.

Acutely isolated or cultured SAMS on laminin-coated glass coverslips were fixed with 4% paraformaldehyde in PBS for 15 min at room temperature. Following three 10-min washes in PBS, cells were permeabilized with 0.02% Triton X-100 in PBS for 30 min at room temperature. Following three more washes in PBS, nonspecific immunoreactive sites were blocked with 10% normal goat serum in PBS for 1 h at room temperature. After three additional PBS washes, cells were incubated overnight at 4°C in a humidified environment with a rabbit polyclonal anti-HCN4 antibody (Alomone Labs Cat No. APC-052) diluted 1:200 in the blocking solution. Cells were again washed three times in PBS and were then incubated in the dark for 1 h with an AlexaFluor 568-conjugated goat anti-rabbit secondary antibody (Invitrogen/LifeTechnologies, Cat No. A11011) at a dilution of 1:1,000 in blocking solution. After three more PBS washes and a brief final rinse with deionized water, coverslips were dried and were mounted onto slides using ProLong Gold anti-fade reagent (Invitrogen). Slides were cured overnight in the dark, and edges of the coverslips were sealed to the slide with nail polish before imaging.

Bright field and epifluorescent images were acquired using an Olympus Q-Color 3 digital camera mounted to an inverted Olympus IX51 microscope with 40× air (fluorescence) or 60× oil (brightfield) objectives. Confocal images were acquired on a 3I MARIANAS inverted spinning disk microscope with a 63× oil objective. SAM size measurements were made using ImageJ software [National Institutes of Health (NIH), Bethesda, MD], with length measured from a line through the center of each cell at its longest point and width measured at the widest point (usually corresponding to the nuclear bulge).

Statistics.

All data are presented as means ± SE. Statistical comparisons were made using unpaired Student's t-tests or one-way ANOVAs as indicated, with P < 0.05 taken as the significance threshold.

RESULTS

Expression of exogenous proteins in cultured SAMs by adenoviral gene transfer.

Spontaneously active SAMs from adult mice could be maintained in culture for up to 1 wk, and cultured cells at all time points exhibited spontaneous contractions immediately upon BDM wash-out. Because the central goal of this study was to establish a system in which exogenous proteins could be introduced into adult SAMs, we pursued several approaches to this end. We first attempted to transiently transfect the cultured cells with a mammalian expression plasmid encoding eGFP using either Lipofectamine 3000 or Xtreme Gene 9 at various ratios of cDNA:reagent. However, no fluorescence was observed even 72 h after transfection (data not shown). In contrast, we obtained high infection efficiency and high expression levels within 24–48 h using adenoviruses to deliver cDNA encoding eGFP and/or mCherry (Fig. 1). Viruses were delivered at a multiplicity of infection (MOI) of 100, which was optimized in trial experiments as the lowest viral titer that produced a high infection efficiency. We found that an MOI of 100 for each adenovirus, administered either separately or together, resulted in nearly 100% infection efficiency for both eGFP and mCherry with no apparent toxicity.

Fig. 1.

Expression of exogenous proteins in sinoatrial node myocytes (SAMs). Bright field (Bright) and epifluorescent images of representative SAMs at either 24 or 48 h after double infection with adenoviruses encoding eGFP (Ad-GFP) and mCherry (Ad-mCherry) at a multiplicity of infection (MOI) of 100 each are shown. Scale bar, 25 μM.

Preserved morphology of cultured sinoatrial myocytes.

We next evaluated the morphology of SAMs in culture. In this and the subsequent functional studies, we chose 48 h as a time point for characterization based on the high level of protein expression achieved within that time frame (Fig. 1). We found that the cultured cells retained an overall morphology that was similar to acutely isolated SAMs (Fig. 2A) and that there were no statistically significant differences in the average length, width, or cross-sectional area of cultured cells compared with acutely isolated cells, although there were trends toward an increase for each of these parameters (n ≥ 8; Fig. 2, B–D). We attribute these trends to a tendency of the cells to flatten as they adhered to the laminin-coated coverslips, and not to an increase in cell volume, because we did not observe a corresponding increase in cellular capacitance in voltage-clamp recordings (n ≥ 20; Fig. 2E).

Fig. 2.

Similar shape and size of acutely isolated and cultured SAMs. A: bright field images of representative SAMs immediately upon isolation (acute) or after 48 h in culture (cultured). B–D: average (±SE) maximum length, width, and cross-sectional area for acutely isolated vs. 48-h cultured SAMs. E: average membrane capacitance from voltage-clamp recordings from acutely isolated or cultured SAMs. NS, not significant: P > 0.05 vs. acute.

AP firing rate and AP waveform parameters of cultured SAMs.

To evaluate the pacemaker function of the cultured cells, we next compared the AP firing rate and AP waveform parameters in cultured SAMs with those in acutely isolated SAMs. Spontaneous APs were recorded in current-clamp mode using the amphotericin perforated-patch technique. As shown in Fig. 3, spontaneous AP firing rates were indistinguishable in acutely isolated and cultured SAMs (acute = 391.5 ± 45.4 APs/min, n = 6; culture = 379.9 ± 32.5 APs/min, n = 14; P > 0.05, t-test). AP waveforms were also similar in acutely isolated and cultured cells (Fig. 4); there were no significant differences in cycle length (CL), maximum diastolic potential (MDP), take-off potential (TOP), repolarization rate, AP duration (total, and at 50% or 90% repolarization), diastolic depolarization rate (DDR), or the total, early, or late phases of the diastolic depolarization (DD). APs in cultured SAMs had small but significant differences in peak voltage (V_max), AP amplitude, and upstroke velocity compared with acutely isolated cells, although these differences did not affect the AP firing rate (Fig. 3).

Fig. 3.

Unchanged action potential (AP) firing rate in cultured SAMs. A: representative current-clamp recordings of spontaneous APs from acutely isolated (top) or 48-h cultured (bottom) SAMs. Dashed lines indicate 0 mV. Scale bars, 20 mV, 200 ms. B: average spontaneous AP firing rate in acute (black; n = 6) and 48-h cultured (gray; n = 14) SAMs.

Fig. 4.

Spontaneous action potential waveform parameters in acutely isolated and cultured SAMs. A: representative APs from amphotericin perforated patch recordings from acutely isolated SAMs (black) or 48-h cultured SAMs (gray) aligned at time of peak. B: comparison of average (±SE) AP waveform parameters in acutely isolated (n = 8) and 48-h cultured SAMs (n = 14). CL, cycle length (ms); MDP, maximum diastolic potential (mV); V_max, maximum voltage (mV); Amplitude, total AP amplitude (mV); TOP, take-off potential (mV); Upstroke V, upstroke velocity (mV/ms); Repol rate, repolarization rate (mV/ms); APD, total action potential duration (ms); APD₅₀, AP duration at 50% repolarization (ms); APD₉₀, AP duration at 90% repolarization (ms); DDR, diastolic depolarization rate (mV/s); DD, total diastolic duration (ms); early DD, early diastolic depolarization duration (ms); late DD, late diastolic depolarization duration (ms). *P < 0.05 vs. acute; all others P > 0.05 (t-tests).

The cardiac funny current (I_f) is a hallmark of SAMs, and I_f contributes to the generation of spontaneous APs. Following current-clamp recordings of spontaneous APs, the amplifier was switched to voltage-clamp mode and each cell was assayed for the presence of I_f. All of the acutely isolated and cultured cells in this study also exhibited I_f of >100 pA in response to a voltage step to −120 mV, consistent with their identity as SAMs (not shown).

No change in I_f or I_Ca in cultured SAMs.

HCN4 channels are the primary molecular correlate of I_f (28), and HCN4 immunoreactivity is used as a marker of the sinoatrial node (4, 19, 12). As shown in Fig. 5A, we observed similar membrane-associated expression of HCN4 in all of the cultured and acutely isolated cells we assayed in confocal immunofluorescence experiments. We next evaluated the biophysical properties of I_f in perforated-patch voltage-clamp experiments. I_f was elicited by 3-s hyperpolarizing voltage steps from −60 to −60 mV from a holding potential of −50 mV. We found no significant differences in the current density, activation kinetics, voltage dependence of activation, or slope factor for I_f in acutely isolated versus cultured SAMs (Fig. 5, B and C; Table 1).

Table 1.

Properties of I_f determined from amphotericin perforated-patch recordings of acutely isolated and 48-h cultured SAMs

				Activation Rate, ms
	Midpoint Activation Voltage, mV	Slope Factor, mV	Current Density, pA/pF	Tau slow	Tau fast
Acutely isolated	−107.9 ± 3.4 (10)	7.99 ± 0.63 (10)	25.4 ± 2.3 (30)	916 ± 181 (8)	117 ± 10 (8)
48-h in vitro	−106.5 ± 1.9 (8)	8.78 ± 0.39 (8)	24.6 ± 6.6 (16)	1342 ± 483 (7)	88 ± 8 (8)

Values are means ± SE. Midpoint activation voltage and slope factor were determined from Boltzmann fits of conductance-voltage plots for each cell. Current density and activation rates were determined from currents elicited by 3-s voltage steps to −150 mV. Number of cells is indicated in parentheses. P > 0.05 for all parameters in acute vs. cultured sinoatrial node myocytes (SAMs) (t-tests). I_f, cardiac funny current.

To further characterize the electrophysiological properties of cultured SAMs, we compared whole-cell calcium currents in cultured SAMs with those in acutely isolated cells. Total I_Ca was measured in response to 200 ms depolarizing test pulses from −80 to +60 mV from a holding potential of −90 mV. There were no significant differences in the peak current density, fast or slow time constants of inactivation, voltage-dependence of activation, or slope factor of I_Ca in cultured compared with acutely isolated SAMs (Fig. 6 and Table 2).

Fig. 6.

Properties of voltage-gated Ca²⁺ currents in acutely isolated and cultured SAMs. A: representative whole cell Ca²⁺ currents recorded from acutely isolated (black) and 48-h cultured (gray) SAMs. Scale bars, 5 pA/pF, 10 ms. B: peak current density for voltage-gated Ca²⁺ currents (I_Ca) in acute (n = 7) and cultured (n = 8) SAMs determined from Boltzmann fits of current-voltage plots. P > 0.05, t-test. C: average fast and slow inactivation rate constants for I_Ca in response to voltage steps to −10 mV in acutely isolated (n = 7)and cultured (n = 8) SAMs. D: voltage-dependence of activation of I_Ca in acute and cultured SAMs.

Table 2.

Properties of I_Ca determined from whole-cell recordings of acutely isolated and 48-h cultured SAMs

				Inactivation Rate, ms
	Maximum Conductance, pS/pF	Peak Voltage, mV	Slope Factor, mV	Tau Slow	Tau Fast
Acutely isolated	0.45 ± 0.04 (7)	−41.0 ± 1.7 (7)	8.8 ± 0.3 (7)	4.01 ± 0.23 (7)	26.38 ± 1.99 (7)
48-h in vitro	0.43 ± 0.07 (8)	−34.9 ± 4.5 (8)	10.7 ± 1.0 (8)	4.08 ± 0.31 (8)	24.36 ± 2.25 (8)

Values are means ± SE. Maximum conductance, peak voltage, and slope factor were determined from Boltzmann fits of conductance-voltage plots for each cell according to Equation 3. Number of cells is indicated in parentheses. P > 0.05 for all parameters in acute vs. cultured SAMs (t-tests). Inactivation rates were determined from double exponential fits of the decaying phases of currents elicited by test pulses to −10 mV. I_Ca, Ca²⁺ current.

DISCUSSION

In this study, we describe for the first time a method for maintaining fully differentiated adult sinoatrial node myocytes in vitro that preserves their characteristic morphology, spontaneous activity, and electrophysiological properties and that is amenable to protein expression studies. Specifically, we showed that exogenous proteins can be expressed at high levels in the cultured cells within 24–48 h via adenoviral gene transfer and that the cultured cells had no significant changes in the AP firing rate, only modest changes in a subset of AP waveform parameters, and no changes in the properties of I_f or I_Ca.

Advantages of the sinoatrial myocyte culture method.

Our new method for culturing adult SAMs and for introducing exogenous proteins into them will greatly expand the range of experimental techniques that can be applied to study the molecular physiology of cardiac pacemaking. For example, the effects of protein overexpression or downregulation (e.g., using RNAi approaches) will enable the dissection of intracellular signaling pathways and the evaluation of the effects of protein interactions. Such gain- and loss-of-function approaches have been mainstays in the study of ventricular myocyte function for many years [e.g., for study of the effects of ion channel interacting proteins (42) and regulatory pathways (24)] but have not been possible to date in sinoatrial myocytes because, until now, there has been no method available that allows protein expression in SAMs. One particularly powerful new experimental approach will be the reintroduction of proteins into cells harvested from knockout animals. This approach will facilitate proof-of-concept rescue experiments and will enable the reintroduction of mutant proteins into an otherwise normal background to study structure-function relationships within the native cellular context.

Another important application of the culture system is that it will enable the use of genetically encoded indicator molecules in sinoatrial node myocytes. Ca²⁺ release from the SR is thought to be critical for pacemaker activity in SAMs (5, 13, 34), and genetically-encoded Ca²⁺ indicators (GECIs) could provide important new insights into this process. GECIs can be targeted to subcellular microdomains and have yielded much important information about localized Ca²⁺ signaling in other types of cardiac myocytes (10, 11, 22, 27, 36). In a similar manner, tethered FRET-based cAMP sensors could be used to characterize spatio-temporal cAMP signaling associated with different GPCRs and phosphodiesterase isoforms, as has been done in other cardiac myocytes (20, 23, 30, 37, 43). cAMP sensors are of particular interest in SAMs, given the fundamental importance of cAMP signaling to sinoatrial myocyte function (15, 16, 41). New genetically encoded sensors are also emerging for many other cellular processes including membrane potential (9); pH, small molecules and lipid metabolites (39), ATP (2, 7), and ROS production (32). Use of such molecules in SAMs could provide novel insights into mechanisms that regulate pacemaking.

The method for culturing SAMs described in the present study fills a gap posed by limitations in existing experimental approaches. For example, although acutely isolated SAMs have yielded a wealth of information about the properties of these highly specialized cells, it is not possible to manipulate protein expression or to conduct longer-term studies in acutely isolated SAMs. Although other cardiomyocytes, such as neonatal ventricular myocytes or HL-1 cells (an immortalized atrial cell line), exhibit spontaneous activity and can be maintained in culture for gene expression studies, these alternative cell types are quite distinct from SAMs. For instance, HL-1 cells exhibit heterogeneous AP waveforms and a large Na⁺ conductance, and contract spontaneously only when they are confluent, indicating that their properties are quite plastic depending on the state of the culture (45). In addition, I_f is expressed in only about one-third of HL-1 cells (45) (when compared with 100% of the cultured SAMs in the present study) and appears to be produced primarily by the HCN1 and HCN2 isoforms (35) (rather than HCN4, which is the predominant isoform in SAMs) (28). Although primary cultures of SAMs from adult rabbits have been reported, the cultured cells exhibited significant morphological dedifferentiation (21, 29, 44) and fired spontaneous APs at a rate at least 50% slower than observed in acutely isolated cells.

Limitations of the new SAM culture system.

One obvious limitation of the mouse SAM culture system is the small size of the mouse SAN and correspondingly small number of SAMs. While adequate cell numbers are available for electrophysiological and imaging studies, the limited amount of tissue precludes extensive biochemical analyses. Although the same problem exists for acutely isolated mouse sinoatrial node tissue, it is exacerbated in culture because the number of cells decreased progressively over time (by ∼50% after 48 h; data not shown). Thus experiments relying on proteins with introduced tags for biochemical purification remain impractical in the mouse SAN. However, we anticipate that our method can be applied with minimal adaptations to prepare SAM cultures from larger species, which may permit such experiments.

In our study, SAMs were cultured in the presence of the reversible myosin ATPase inhibitor, BDM, which prevents contraction by inhibiting cross-bridge cycling (1, 3, 31). However, BDM is well-known to have off-target effects, including nonspecific phosphatase activity, inhibition of cardiac transcription factors (40), and inhibition of sodium channels, calcium channels, and ryanodine receptors (3). Although it would be preferable to avoid the use of BDM, we found it to be an essential component of our SAM culture protocol; we attempted numerous cultures in the absence of BDM, but in each case, the number of spindle-shaped spontaneously active cells was dramatically reduced, and no such cells were observed beyond 24 h in vitro (not shown). These observations are consistent with previous reports in which adult rabbit SAMs cultured in the absence of BDM tended to round-up and dedifferentiate (21, 29, 44).

It is possible that other contractile inhibitors may have a more desirable pharmacological profile than BDM. We examined one such alternative, the myosin II inhibitor, blebbistatin (25 μM), which has been shown to improve viability of ventricular myocyte cultures (7). However, SAM survival and morphology were dramatically compromised in cultures prepared with blebbistatin than with cultures prepared with BDM (not shown). Thus there appear to be differences between the optimal conditions for the culture of different types of cardiac myocytes. Despite its known off-target effects, our data show that BDM has rather minimal consequences on the morphological and electrophysiological properties of SAMs.

Finally, for the purposes of characterizing the cultured cells in this study, we compared them with acutely isolated cells. Although most properties were remarkably unchanged in culture, we observed small but significant differences in a subset (3 of 14) of AP waveform parameters in the cultured SAMs. The mechanism(s) responsible for these changes are not known (although they seem to be independent of both I_f and I_Ca, which were not altered in vitro) and we have not exhaustively characterized all of the other the possible changes that could occur in SAMs in culture. However, for many experimental applications, the most appropriate comparisons would be made between cultured SAMs under different conditions (e.g., in the presence vs. the absence of an introduced protein). Such comparisons would tend to negate the relatively minor changes arising from the culture conditions alone. Furthermore, primary cultures of adult ventricular myocytes also display some changes in vitro, but these systems have nonetheless enabled important advances in our understanding of cardiac cellular physiology. We anticipate that the new method presented herein will allow similar progress in our understanding of sinoatrial node myocytes.

GRANTS

This work was supported by NIH National Heart, Lung, and Blood Institute Grant R01-HL088427 (to C. Proenza). E. J. Sharpe was supported by NIH-5T32-AG000279 from the National Institute on Aging.

DISCLAIMER

The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

Author contributions: J.R.S. and C.P. conception and design of research; J.R.S., E.J.S., and C.P. performed experiments; J.R.S., E.J.S., and C.P. analyzed data; J.R.S., E.J.S., and C.P. interpreted results of experiments; J.R.S. and C.P. prepared figures; J.R.S., E.J.S., and C.P. edited and revised manuscript; J.R.S., E.J.S., and C.P. approved final version of manuscript; J.R.S. and C.P. drafted manuscript.