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Transforming growth factor-1 decreases cardiac muscle L-type Ca²⁺ current and charge movement by acting on the Ca_v1.2 mRNA

¹Department of Biochemistry and ²External Section of Toxicology, Cinvestav, México

Submitted 20 July 2006 ; accepted in final form 11 September 2006

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Transforming growth factors- (TGF-s) are essential to the structural remodeling seen in cardiac disease and development; however, little is known about potential electrophysiological effects. We hypothesized that chronic exposure (6–48 h) of primary cultured neonatal rat cardiomyocytes to the type 1 TGF- (TGF-1, 5 ng/ml) may affect voltage-dependent Ca²⁺ channels. Thus we investigated T- (I_CaT) and L-type (I_CaL) Ca²⁺ currents, as well as dihydropyridine-sensitive charge movement using the whole cell patch-clamp technique and quantified Ca_V1.2 mRNA levels by real-time PCR assay. In ventricular myocytes, TGF-1 did not exert significant electrophysiological effects. However, in atrial myocytes, TGF-1 reduced both I_CaL and charge movement (55% at 24–48 h) without significantly altering I_CaT, cell membrane capacitance, or channel kinetics (voltage dependence of activation and inactivation, as well as the activation and inactivation rates). Reductions of I_CaL and charge movement were explained by concomitant effects on the maximal values of L-channels conductance (G_max) and charge movement (Q_max). Thus TGF-1 selectively reduces the number of functional L-channels on the surface of the plasma membrane in atrial but not ventricular myocytes. The TGF-1-induced I_CaL reduction was unaffected by supplementing intracellular recording solutions with okadaic acid (2 µM) or cAMP (100 µM), two compounds that promote L-channel phosphorylation. This suggests that the decreased number of functional L-channels cannot be explained by a possible regulation in the L-channels phosphorylation state. Instead, we found that TGF-1 decreases the expression levels of atrial Ca_V1.2 mRNA (70%). Thus TGF-1 downregulates atrial L-channel expression and may be therefore contributing to the in vivo cardiac electrical remodeling.

calcium channel; atrial fibrillation; muscle disease

Compelling evidence has accumulated during the last 20 years, which demonstrates that the TGF- signaling is essential to the structural remodeling seen during cardiac development and disease. For example, Thompson et al. (42) showed, in an animal model of myocardial infarction, that cardiomyocytes increase the expression levels of TGF-1 in response to myocardial stress. More recently, an increase on expression levels of TGF-1 has been also detected in patients with idiopathic hypertrophic cardiomyopathy (25), dilated cardiomyopathy (37), and lone atrial fibrillation (AF) (AF without ischemic heart disease, Ref. 39) as well as in animal models of norepinephrine-induced hypertrophy (4), progressive coronary artery occlusion (47), pressure overload (46), and congestive heart failure (16). Perhaps more important than a simple correlation is the striking observation that TGF-1 per se reproduces most of the hallmarks seen in structural remodeling (for recent reviews see Refs. 8, 14, 31, 38). Thus, not surprisingly, significant efforts are being currently made to discover potential therapeutic roles for TGF- signaling in cardiac disease (2, 7, 24, 26, 32, 34, 35). In contrast, little is known with regard to potential TGF--related electrophysiological effects, despite that electrical remodeling is an important substrate for cardiac disease.

Thus we have hypothesized that a chronic exposure of cardiomyocytes to TGF-1 might reproduce critical events associated to electrical remodelling. Specifically, we investigated whether TGF-1 provokes significant alterations in the function and expression of voltage-dependent Ca²⁺ channels. Overall, our results indicate TGF-1 provokes a selective inhibition on L-channel expression from atrial but not ventricular myocytes. We propose these results may contribute to explaining in vivo electrical remodeling of the heart.

	MATERIALS AND METHODS

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Reagents. Unless specified, the reagents were purchased from Sigma (St. Louis, MO). TGF-1 was obtained from either Sigma or PrepoTech (Rocky Hill, NY). Likewise, okadaic acid was from either Sigma or Calbiochem, EMD Biosciences (San Diego, CA). Media, serum, enzymes, reagents, and materials for tissue culture were purchased from Invitrogen (Carlsbad, CA).

Primary cultures of cardiomyocytes. Primary cultures of cardiomyocytes were obtained from male neonatal (4 days old) Wistar rats, which were decapitated to isolate the hearts; all animal manipulations were conducted in accordance with the Mexican Official Norm NOM-062-ZOO-1999 and the Guide for the Care and Use of Laboratory Animals as adopted by the National Institutes of Health. Subsequently, ventricular and atrial tissues were dissected and enzymatic digested separately. Only the midlower half of the ventricles was used (inferior ventricle) to avoid myofibroblasts coming from the valves. The cardiac tissue was further dispersed mechanically and centrifuged, and the resulting cells were preplated 2 h at 37°C. This step reduces fibroblast content on the grounds that fibroblasts attach faster to a plastic surface, as opposed to cardiomyocytes. Subsequently, the semipurified cardiomyocytes were plated on glass coverslips and kept at 37°C in culture medium consisting of Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 2% of either horse serum or fetal bovine serum, 100 U/ml penicillin, 100 µg/ml streptomycin, and 4 mM L-glutamine. Identical results were obtained if using horse serum or fetal bovine serum. The myocytes were kept in culture from 0 to 3 days (i.e., from day 0 to day 3). At day 1 or day 2, some myocytes were transferred from the control culture conditions to a culture medium supplemented with 5 ng/ml TGF-1 and then cultured for additional 6–48 h (TGF-¹-treated myocytes). At this concentration, TGF-1 elicits nearly maximal effects on cardiomyocytes gene expression (27).

Voltage-clamp experiments. Myocytes were transferred from the culture medium to an external solution containing (in mM): 140 choline-Cl, 20 TEA-Cl, 2 CaCl₂, 10 glucose, 10 HEPES, and 0.03 tetrodotoxin (pH = 7.4 with TEA-OH). Approximately 15–60 min thereafter, the cells were subjected to voltage-clamp experiments, using the whole cell patch-clamp technique, as described previously (29). The patch electrodes were elaborated from borosilicate glass capillaries and exhibited electrical resistances of 2–4 m. The standard pipette filling solution consisted of (in mM) 135 Cs-Asp, 10 MgCl₂, 10 EGTA, 1 CaCl₂, 10 HEPES, 5 glucose, 5 ATP-Mg, and 0.05 GTP-Na₂ (pH = 7.4 with CsOH). We estimate 10 mM EGTA plus 1 mM of total Ca²⁺ should buffer the final concentration of free Ca²⁺, close to 15 nM. In some experiments (see Fig. 6), the pipette solution was supplemented with either 2 µM okadaic acid or 200 µM cAMP. Cell capacitance (C_m) was determined by integration of linear capacity transients resulting from control pulses (from –80 mV to –100 mV) and was used to normalize currents (pA/pF) from different myocytes. The holding potential (HP) was –80 mV, unless otherwise specified. The current signals were filtered at 1 kHz, using a four-pole Bessel filter, and recorded at 50 kHz (10- to 30-ms pulses) and 5 kHz (100- to 200-ms pulses). Dissociation of I_CaT and I_CaL was performed either changing the HP from –80 mV to –50 mV [current-voltage (I-V) curves] or applying a two-pulse protocol that consists of a long (either 100 or 200 ms) pulse from –80 mV to –30 mV, which preferentially activates and inactivates T-type Ca²⁺ current (I_CaT); followed by a second pulse to +20 mV, which primarily elicits L-type Ca²⁺ current (I_CaL), since I_CaT remains inactivated. Current densities corresponding to I_CaT and I_CaL were plotted independently as a function of test potential (I-V curves) and fitted according to the following equation:

(1)

where G_max is the maximal channel conductance, V_m is the test potential, V_rev is the extrapolated reversal potential, V_G(1/2) is the potential for half-maximal activation of G_max, and k_G is a slope factor. The Ca²⁺ channels voltage dependence of inactivation was accessed by application of 1-s prepulses of different voltage amplitudes, which were followed by 200-ms pulses to either –30 mV or +20 mV. Subsequently, T- (–30 mV) and L-current (+20 mV) densities were plotted as a function of the membrane potential during the prepulse and fitted according to:

(2)

I_max represents the maximal current density, V_i(1/2) is the potential for half-maximal inactivation of I_max, and k_i is a slope factor.

View larger version (21K): [in this window] [in a new window]   Fig. 1. Transforming growth factor (TGF)-1 regulates L-type (ICaL) but not T-type (ICaT) Ca2+ current in atrial myocytes. A, B: representative ICaT (A) and ICaL (B) recorded from atrial myocytes cultured 3 days under control conditions (day 3; control, left), or 2 days under control conditions, plus one additional day in the presence of TGF-1 (i.e., from day 2 to day 3; TGF-1, right). A double test-pulse protocol was used, setting test potential (Vm) at –50 mV during a 30-ms interpulse. C: average values of cell membrane capacitance (Cm) (a), ICaT (b), and ICaL (c) obtained from experiments as in A and B. The numbers of investigated myocytes are shown in a. Exposure of myocytes to TGF-1 started at day 2. Solid triangles, control; shaded triangles, TGF-1. D, E: ICaT (D) and ICaL (E) I-V curves obtained from a total of 14 control (day 2 plus day 3, solid triangles) and seven TGF-1-treated (1 day, from day 2 to day 3, shaded triangles) atrial myocytes. Average kinetic parameters describing the corresponding voltage dependences of activation are shown in Table 1 (atria).

View larger version (30K): [in this window] [in a new window]   Fig. 2. Voltage dependence of activation and inactivation of Ca2+ channels is unaffected. A: representative traces of ICaL (top) and ICaT (bottom) recorded from a control atrial myocyte (day 3), following 1-s inactivating prepulses to the indicated potentials (Vm, left). B, C: normalized inactivation (squares) and activation (triangles) curves, obtained from current traces as in A and the current-voltage (I-V) curves shown in Fig 1. Both current density (squares) and channel conductance (triangles) were normalized to their corresponding maximal values (i.e., Imax and Gmax) that were calculated from each particular myocyte. The average maximal values are given in Table 1 (Gmax) and Table 2 (Imax). Solid symbols, day 2-day 3 control atrial myocytes; shaded symbols, 1 day-treated atrial myocytes (from day 2 to day 3).

View larger version (20K): [in this window] [in a new window]   Fig. 3. TGF-1 does not regulate Ca2+ currents in ventricular myocytes. A, B: representative ICaT (A) and ICaL (B) obtained from ventricular myocytes cultured either 3 days under control conditions (left, control), or 2 days under control conditions plus one additional day in the presence of TGF-1 (right, TGF-1). The current traces were acquired as described in Fig. 1A. C: average values of cell Cm (a), ICaT (b), and ICaL (c) that were obtained from control (solid circles) and treated (shaded circles) ventricular myocytes, which were kept different times in culture. The treatment with TGF-1 started at day 2. D, E: ICaT (D) and ICaL (E) I-V curves obtained from a total of 15 control (day 2 and day 3, solid circles) and 14 TGF-1-treated (1 day, from day 2 and day 3, shaded circles) ventricular myocytes. The corresponding kinetic parameters describing voltage dependences of activation are shown in Table 1 (ventricle).

View larger version (29K): [in this window] [in a new window]   Fig. 4. TGF-1 reduces charge movement in atrial but not ventricular myocytes. A, B: analysis of nonlinear capacitive currents recorded from atrial (A, squares) and ventricular (B, circles) myocytes. a: representative nonlinear capacitive currents that were recorded from control (top) and TGF-1-treated (bottom) myocytes. The currents were elicited from a holding potential (HP) of –50 mV, to the indicated test potentials (left). b, c: average voltage dependence of charge moving outward (QON) on depolarization (b), and corresponding normalized curves (c). Results were obtained from day 2 and day 3 control myocytes (control, solid symbols), and myocytes that were treated from day 1 to either day 2 or day 3 (TGF-1, shaded symbols). The absolute values of QON from each particular myocyte were divided by Cm (nC/µF, b) and then fitted according to a Boltzmann equation (Eq. 3). Subsequently, QON was normalized by the corresponding maximum intramembrane charge movement (or Qmax) estimated from the fitting procedure (normalized curves, c). Average Boltzmann parameters as well as the total number of experiments are given in Table 3. The dashed lines in c represent normalized L-channel activation curves, obtained under similar cationic concentrations, as the charge movement. The activation curves represent the average from seven atrial (Ac, dashed line) and five ventricular (Bc, dashed line) control myocytes. The average values of VG(1/2) and kG for atrial and ventricular myocytes were, respectively (in mV): 11.9 ± 4.0 and 9.7 ± 1.1, and 13.0 ± 3.1 and 9.4 ± 0.3.

View larger version (17K): [in this window] [in a new window]   Fig. 5. Effect on Qmax is parallel to ICaL reduction. The figure shows average values of Qmax (estimated as in Fig. 4) that were obtained from atrial myocytes, as a function of the days in culture. Solid circles, control; shaded circles, TGF-1 (applied since day 1). Inset: time course of ICaL inhibition by TGF-1 (shaded triangles) in atrial myocytes treated since day 1. Average values of ICaL were obtained from a minimum of six (TGF-1; day 3) and a maximum of 33 (control; day 2) experiments.

View larger version (17K): [in this window] [in a new window]   Fig. 6. TGF-1 decreases atrial L-channel expression but not permanent phosphorylation state. A: average ICaL that was estimated at +20 mV (by a two-pulse protocol), using the following internal solutions: standard (left), standard supplemented with 2 µM okadaic acid (OA, middle), and standard supplemented with 100 µM cAMP (cAMP, right). Solid bars, control myocytes (day 1-day 3); shaded bars, treated myocytes (1 day-2 day, from day 1). ICaL was normalized by the corresponding absolute values obtained from control myocytes (–5.4 ± 0.5 pA/pF, standard; –4.5 ± 0.6 pA/pF, OA; and –14.9 ± 2.7 pA/pF, cAMP). B: L-channel I-V curves that were obtained from the same cells as in A (cAMP), using 30-ms pulses delivered from a HP of –50 mV. Solid squares, control myocytes; shaded squares, treated myocytes. C: quantitative levels of mRNA encoding to the principal subunit of L-channels (CaV1.2), which were estimated from control (solid bars) and TGF-1-treated (shaded bar) myocytes, according to the threshold cycle method. Data represent the mean value from six independent determinations. mRNA values were corrected by parallel determinations of 18S rRNA (t), and the relative changes estimated according to: 2[exp – (t – tref)], where tref is t from day 3, control myocytes.

(3)

where Q_max, V_Q(1/2), and k_Q have their usual meanings with regard to charge movement.

Real-time quantitative PCR detection. PCR reactions were performed using an ABI PRISM 7000 sequence detector system (Perkin-Elmer Applied Biosystems, Foster, CA) and analyzed using the comparative threshold cycle (C_t) method, as previously described (28). Briefly, total mRNA was isolated from control and treated myocytes using the TRIzol reagent according to manufacturer’s instructions (Invitrogen Life Technologies Carlsbad, CA). Subsequently, quantitative real-time PCR assay of the cDNA transcripts was prepared from 2 µg of total RNA using a SuperScript First Strand Synthesis kit. The genes encoding Ca_V1.2 and 18S ribosomal RNA (rRNA, endogenous control) were simultaneously amplified in a single PCR reaction to allow for normalization of the mRNA data. The PCR reaction mixture consisted of probes (0.25 µM), specific primers (0.9 µM), cDNA (4 µl), and 1X TaqMan Universal PCR Master Mix. The following primers and prove FAM were used to analyze the Ca_V1.2 mRNA: 5'-CGAAGCTCAACTCAACTGTTTCTAC-3'(forward); 5'-GCATTGGCATTCATGTTGGCAT-3'(reverse); 5'-CCATAGTTGGAACCTCC-3'(probe FAM).

Data analysis and statistics. All data are expressed as the means ± SE. Significant differences were determined at the P < 0.05 level, unless specified. Student’s unpaired t-tests (2-tailed) were used to compare a single mean value between two independent cell groups (two-sample comparisons). The acquisition and analysis of experimental results were performed using commercially available software suites (Microsoft Excel 2002, Redmond, WA, and pClamp V9.0, Axon Instruments, Union City, CA).

	RESULTS

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TGF-1 downregulates I_CaL in atrial but not ventricular myocytes. We initially aimed to compare the control values of atrial C_m, I_CaT, and I_CaL during long-term culture conditions. Toward this end, we applied a two-pulse protocol that allows us to simplify dissociation of I_CaT and I_CaL (Figs. 1, A and B). Under control conditions (Fig. 1C, closed symbols), I_CaL drastically increased between days 0–1 and then remained relatively constant. In addition, between days 0 and 3, C_m increased approximately twofold, and I_CaT was relatively stable. In general, cardiomyocytes were spherically shaped at day 0, progressively spread out within days 1–3, and then became flattened by day 4. In fact, by day 4 of culture, it was practically impossible to perform patch-clamp experiments, probably because of these morphological alterations. Thus we only focused on myocytes cultured up to 3 days.

We first decided to initiate the treatment with TGF-1 at day 2. As a result, TGF-1 markedly reduced I_CaL, following applications of 1 day, up to a 45% of their respective control value (i.e., control-day 3); in the absence of significant changes in either C_m or I_CaT (Fig. 1C). The underlying molecular mechanisms apparently involve slow cellular processes, as opposed to a fast and direct modulation in the L-channels activity. This is because the effect exhibited a relatively slow onset. In fact, the effect was not statistically significant at 0.5 days (Fig. 1C). Moreover, the effect seems to be very persistent, because it was observed following 15–60 min of TGF-1 withdrawal (the experiments were performed in a growth factor-free recording solution, see the composition of external solution in MATERIALS AND METHODS).

Ca²⁺-channel current-voltage (I-V) curves were also compared between control and TGF-1-treated atrial myocytes (Fig. 1, D and E). Interestingly, whereas TGF-1 significantly reduced I_CaL (55%) at several membrane potentials (Fig. 1E), the I-V curves corresponding to T-channels were practically unchanged (Fig. 1D). The I-V curves shown in Fig. 1, D and E, were fitted according to a Boltzmann equation (Eq. 1), and the resulting kinetic parameters are shown in Table 1. Results from this fitting procedure indicate that the growth factor selectively reduces (50%) the G_max in the absence of significant alterations on the other Boltzmann parameters describing L-channel activation curves (Table 1; atria).

We also investigated the voltage dependence of the inactivation of T- and L-channels (Fig. 2; normalized current, squares). The treatment of atrial myocytes with TGF-1 produced a marked reduction (50%) in the maximal value of I_CaL, without significantly altering the maximal value of I_CaT, or the other Boltzmann parameters describing inactivation curves [i.e., V_i(1/2) and k_i; Table 2]. These results strongly suggest that the inhibitory effect of TGF-1 on I_CaL cannot be explained by a possible increase in the fraction of inactivated channels.

Figure 3, A–C, shows values of C_m, I_CaT, and I_CaL, estimated from ventricular myocytes that were kept in culture from 0 to 3 days. By day 2, some myocytes were transferred to TGF-1 and cultured for an additional 12 or 24 h. In contrast to atrial myocytes, the growth factor did not affect I_CaL, and we did not detect either significant differences on C_m or I_CaT. On the other hand, the Ca²⁺ currents shown in Fig. 3, A–C were measured at –30 (I_CaT) and +20 mV (I_CaL), and it was conceivable that TGF-1 might be regulating Ca²⁺ channels at other voltages. Thus we compared current-to-voltage relationships, or I-V curves, between control and TGF-1-treated ventricular myocytes. We found, however, that the growth factor did not affect I-V curves corresponding to either I_CaT (Fig. 3D) or I_CaL (Fig. 3E). This suggests that the growth factor was unable to regulate the conductance of T- and L-channels at different membrane potentials. Consequently, the Boltzmann parameters describing the corresponding voltage dependences of activation were practically identical between control and TGF-1-treated myocytes (Table 1; ventricle). Altogether, results shown in Figs. 1–3 strongly suggest that the growth factor preferentially regulates L-channels in atrial but not ventricular myocytes.

TGF-1 downregulates dihydropyridine (DHP)-sensitive charge movement in atrial but not ventricular myocytes. It is possible that TGF-1 regulates the number of L-channels on the plasma membrane, since the corresponding kinetic parameters of I_CaL were unaffected. Thus we decided to investigate whether the growth factor affects the intramembrane charge movement associated to L-channels. To isolate charge movement that could potentially be associated to L-channels, we used a HP of –50 mV and blocked Ca²⁺ currents by adding CdCl₂ (3 mM) and LaCl₃ (0.1 mM) to the external solution. We next investigated whether this charge movement could be also regulated by TGF-1. Figure 4, A,a and B,a, shows representative traces of nonlinear capacitive currents elicited at different test potentials. In both control and TGF-1-treated myocytes, the capacitive currents tend to saturate at positive potentials and exhibit an activation threshold of approximately –30 mV. Interestingly, however, in the TGF-1-treated atrial myocyte (Fig. 4A,a, bottom), capacitive currents tend to be smaller. On average, the Q_ON estimated from TGF-1-treated atrial myocytes was significantly smaller at different potentials compared with the respective control condition (Fig. 4A,b). According to this, the maximum amount of charge movement (Q_max) obtained from the treated atrial myocytes was 60% reduced (Table 3). These results suggest that in atrial myocytes, a chronic exposure to TGF-1 significantly decreases the number of voltage sensors associated to L-channels. Moreover, since the growth factor did not affect the voltage dependence of charge movement (Fig. 4A,c and Table 3), further support is added to the notion that it is the number of channels that is being regulated as opposed to the corresponding functional properties.

As mentioned before, we directly added to the external solution 3 mM Cd⁺² plus 0.1 mM La³⁺ to block Ca²⁺ currents. However, both divalent and trivalent inorganic cations are known to shift toward more positive potentials the activation of ion channels (18). Therefore, we investigated the effect on I_CaL of adding similar cationic concentrations as those used for isolating charge movement (Fig. 4, Ac and Bc, dashed lines). Specifically, we added 3.15 mM CaCl₂ to the standard external solution to yield a total Ca²⁺ concentration of 5.15 mM. Under these conditions, the membrane potential required to activate 50% of G_max [or V_G(1/2)] was 11.9 ± 4.0 mV for atrial myocytes and 13.0 ± 3.1 mV for ventricular myocytes. These values are 14 mV more positive than using 2 mM Ca²⁺ (see Table 1) and 9 mV more positive than the potential for half-maximal activation of charge movement [or V_Q(1/2); see Table 3]. Thus, under similar cationic concentrations, the charge movement-to-voltage relationships (Q-V curves) are located at more negative potentials compared with the activation curves of L-channels (G-V curves; Fig. 4, Ac and Bc). This particular distribution of charge and conductance, across the voltage axis, represents an important requirement for charge movement to actually reflect the corresponding activity of the voltage sensor.

To corroborate that the charge movement shown in Fig. 4 is DHP sensitive, we used 4 µM of the DHP antagonist nifedipine applied to the Cd²⁺/La³⁺ external solution. As a result, the average Q_ON was significantly reduced to only a 6% of the standard condition (data not shown). We also aimed to determine whether control pulses used to subtract linear components might be eliciting nonlinear responses. For this, we applied hyperpolarizing test pulses within the membrane potentials that are used by our standard p/-3 subtraction protocol (i.e., from –80 mV to –120 mV). Subsequently, we applied control pulses of opposite polarity (i.e., from –80 to –65 mV) to subtract the corresponding elicited responses. These results indicated that, at these voltages, both capacitive and resistive asymmetric components were practically absent (data not shown). Thus 4 µM nifedipine suppresses 94% of nonlinear charge movement in neonatal cardiomyocytes at a HP of –50 mV. It is possible therefore that this charge movement originates mostly (if not entirely) from the voltage-sensing activity of L-channels.

Figure 5 shows onset of inhibitory effects by TGF-1 on both Q_max and I_CaL. The effects are parallel, suggesting the presence of a common molecular mechanism. In addition, the inset in Fig. 5 extends an important observation made from Fig. 1, in the sense that reduction of I_CaL requires more than 12 h (Fig. 1 C,c) to be statistically significant. According to this, in Fig. 5, I_CaL was not significantly altered following 6 h of treatment. It is important to remark that in contrast to Fig. 1, the treatment in Fig. 5 was now initiated at day 1 (instead of day 2). This allowed us to resolve that both effects (i.e., changes on I_CaL and Q_max) persist for a minimum of 2 days (i.e., from day 1 to day 3), corroborating that they remain active during long-term conditions and may thus be significant under steady-state situations in vivo.

Molecular mechanisms point to alterations on L-channel expression but not phosphorylation. A reduced I_CaL is a hallmark of atrial electrical remodeling in AF (44, 49). Recent results suggest that it is a decreased L-channel phosphorylation that keeps low I_CaL. This is supported by the observations that in atrial myocytes derived from certain models of AF, both okadaic acid (a phosphatase inhibitor) and cAMP overstimulate I_CaL compared with the corresponding control myocytes. Consequently, both compounds quickly eliminate (in seconds or minutes) significant differences showing a reduced atrial I_CaL in AF (5, 9). Motivated by these previous observations, we aimed to determine whether inhibition of atrial I_CaL by TGF-1 could be reverted by similar manipulations on L-channels phosphorylation state. However, adding 2 µM of okadaic acid to the internal solution did not affect the corresponding 50% reduction on I_CaL (Fig. 6A, middle). As a matter of fact, this compound did not affect either the absolute values of I_CaL in both control and TGF-1-treated myocytes (see the legend of Fig. 6).

Atrial myocytes were also dialyzed with an internal solution containing 100 µM cAMP to activate directly the cAMP-dependent protein kinase (PKA). In contrast to okadaic acid, cAMP provoked a marked increase in the absolute values of I_CaL, in both control and TGF-1-treated myocytes (compare Fig. 6B with Fig. 1E). Nevertheless, identical to okadaic acid, cAMP did not affect the TGF-1-induced 50% reduction on I_CaL (Fig. 6, A and B). This is explained by the fact that percentages of I_CaL stimulation by cAMP were comparable between control (180%) and TGF-1-treated (160%) myocytes. Moreover, in both experimental conditions, cAMP also shifted the L-channel activation curves to a similar extent (approximately –7 mV). Specifically, the average values of V_G(1/2) obtained in the presence of cAMP were the following: –9 ± 0.9 mV and –6.0 ± 1.8 mV for control and TGF-1-treated myocytes, respectively (compare these numbers with the corresponding values shown in Table 1). Taken together, results shown in Fig. 6, A and B, strongly suggest that permanent alterations in the L-channels phosphorylation state are not contributing to explaining I_CaL reduction by TGF-1.

A reduced L-channel expression has been also proposed to explain atrial I_CaL reduction by AF (6, 13, 22, 43). Thus we next decided to investigate whether TGF-1 might be regulating atrial L-channel expression. Specifically, we investigated the expression levels of mRNA encoding to Ca_V1.2, the principal subunit of L-channels. Interestingly, as can be seen in Fig. 6C, in the TGF-1-treated myocytes (from day 2 to day 3, gray bar) the Ca_V1.2 mRNA levels were significantly reduced (70%) compared with the corresponding control conditions (Fig. 6C, black bars). These results strongly suggest that a chronic exposure of atrial myocytes to TGF-1 drastically decreases L-channel expression.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Our study shows that a chronic exposure of atrial myocytes to TGF-1 markedly reduces both I_CaL and DHP-sensitive charge movement, in the absence of significant alterations on cell C_m, I_CaT, or channel kinetics. A similar exposure of ventricular myocytes to TGF-1 was unable to regulate I_CaL, I_CaT, C_m, or DHP-sensitive charge movement. Reduction of atrial I_CaL by TGF-1 was not affected by the presence of intracellular okadaic acid and cAMP, two compounds that promote L-channel phosphorylation. Instead, inhibition of atrial I_CaL and charge movement was associated to a significant reduction on steady-state levels of Ca_V1.2 mRNA. It is possible, therefore, that TGF-1 might be acting by reducing Ca_V1.2 gene expression, although we cannot discard a potential effect on mRNA stability or degradation.

Comparison to previous studies. To our knowledge, only a previous study has attempted to identify DHP-sensitive charge movement in neonatal rat cardiomyocytes (from the ventricle, Ref. 12). However, using a HP of –60 mV, Field et al. (12) found a negative charge movement induced by nifedipine, which did not allow characterizing further DHP-sensitive components. In addition, in a cell held at 0 mV (1 min; in the absence of nifedipine), a brief hyperpolarization stimulus produced asymmetric charge (relative to a control step from 0 mV to +20 mV), complicating estimations of the absolute amount of nonlinear components. In our experiments, however, we did not find evidence for a negative charge induced by nifedipine nor identified asymmetric charge within the voltages used to estimate nonlinear components (–65 mV to –120 mV). In a related study, Josephson and Sperelakis (19) reported that in embryonic chick ventricular myocytes, the nonlinear charge movement is barely detectable (<5% of the total) at hyperpolarized potentials (–100 mV to –120 mV). Our present results are in line with this interpretation.

In a previous interesting study, Lalevée et al. (23) showed that aldosterone upregulates in situ T- and L-channel expression in ventricular but not atrial myocytes (obtained from neonatal rats). In contrast, we showed here that TGF-1 downregulates I_CaL in atrial but not ventricular myocytes. Thus differential responses of atrial and ventricular myocytes, to a particular extracellular stimulus, are probably manifestations of a more general functional heterogeneity.

I_CaT is practically absent in adult ventricular myocytes. In fact, it can only be resolved at the earliest stages of development. On the other hand, cardiac remodeling involves, in addition to hypertrophy, regression to what is known as the "fetal gene expression program". Apparently, this process includes reexpression of ventricular I_CaT (reviewed in Ref. 48). Most interestingly, TGF-1 reproduces certain aspects of regression to the fetal gene expression program (36). Thus, TGF-1 would be expected to augment T-channel expression in ventricular myocytes. Conversely, we did not find significant differences on ventricular I_CaT in response to TGF-1. The possibility still remains, however, that in adult ventricular myocytes, TGF-1 provokes I_CaT reexpression, which may be part of its ability to promote regression to the fetal gene expression program.

Molecular mechanisms. A hallmark of electrical remodeling in AF is a significant reduction of I_CaL (44, 49). However, the precise molecular mechanisms that keep low I_CaL remain controversial. The following mechanisms represent perhaps the most commonly accepted: 1) reduced level of channel expression (6, 13, 22, 43), and 2) reduced level of channel phosphorylation (5, 9). Conceivable, different times of the presence and development of AF, as well as the intrinsic differences in experimental models, might contribute to explaining this controversy.

In addition to alterations on gene expression, the TGF- signaling promotes activation of protein phosphatase 1 and 2A (i.e., PP1 and PP2A; reviewed in Ref. 10). On the other hand, an increased expression and activity of PP2A is thought to be responsible for I_CaL reduction in human AF (9). Thus, conceivable I_CaL reduction by TGF-1 could be explained by a possible increase in phosphatase activity and a subsequent reduction on L-channel phosphorylation. However, the absence of significant alterations on L-channel kinetics strongly argues against this possibility. We nevertheless decided to investigate a potential role for L-channel phosphorylation, motivated by the current controversy, regarding to the molecular mechanisms involved on I_CaL reduction by AF.

Okadaic acid is a commonly used inhibitor of PP1 and PP2A (11). These phosphatases promote in turn L-channel dephosphorylation and reduce therefore I_CaL (20). Moreover, okadaic acid eliminates rapidly (in seconds or minutes) significant differences on I_CaL (50–70% reduction), which are associated to AF (5, 9). In our experiments however, okadaic acid was unable to revert the inhibitory effect on I_CaL by TGF-1 (Fig. 6A). Thus reduced I_CaL by TGF-1 cannot be explained by a potential increase on the fraction of PP1/PP2A-dephosphorylated L-channels. In addition to okadaic acid, cAMP also eliminates significant differences on atrial I_CaL provoked by AF (5). We found, however, that cAMP did not affect either the TGF-1-induced 50% reduction in I_CaL (Fig. 6, A and B). This suggests that L-channels from control and TGF-1-treated myocytes respond similar to PKA stimulation and exhibit also a common basal state of phosphorylation.

As mentioned before, another mechanism proposes a decreased channel expression (6, 13, 22, 43) to explain I_CaL reduction by AF. The following observations indicate that a similar mechanism explains the present I_CaL reduction by TGF-1. First of all, TGF-1 reduced 50% the maximal channels conductance or G_max, which is directly proportional to the number of channels of the plasma membrane. Second, TGF-1 significantly reduced (60%) the maximal values of DHP-sensitive charge movement (Q_max) to a similar extent as the corresponding effects on I_CaL and G_max. Thus, inhibition of I_CaL and G_max are both explained by a significant reduction on the density of L-channel voltage sensors. Finally, the growth factor significantly reduced (70%) the steady-state levels of Ca_V1.2 mRNA, which encodes to the principal L-channels subunit. This suggests that the growth factor decreases transcriptional rate of the Ca_V1.2 gene, promotes degradation of the corresponding mRNA, or provokes both effects. The reduced mRNA levels, in turn, should be expected to decrease the synthesis of L-channels. It is important to remark that we investigated only long-term effects (from 6 to 48 h). Thus it is still possible that TGF-1 might also decrease I_CaL in seconds or minutes, by promoting an increased PP2A activity and a subsequent L-channel dephosphorylation. Obviously, further experiments are needed to eventually prove these hypotheses.

Possible physiological relevance. TGF- signaling is also important for heart development, in addition to disease. In fact, several members of the TGF- superfamily have been involved in cardiac differentiation (8, 31). The corresponding effects involve induction of cardiac-specific genes, most likely including Ca_V1.2. Thus TGF--stimulated stem cells (or TGF--preprogrammed stem cells) are often used to improve the therapeutic benefit of stem cells transplantation in diseased hearts (2, 26). Within this context, our present study can be interpreted to suggest that TGF-1 may not represent the most effective candidate to successfully promote cardiac regeneration from TGF--preprogrammed stem cells. This is because our results suggest that this particular isoform most likely restrains cardiac L-channel expression instead of promoting induction. Further support to this interpretation comes from the fact that TGF-1 provokes regression to the fetal gene expression program, instead of promoting development to the adult phenotype (36). Moreover, in a recent study, TGF-2, but not TGF-1 or –3, was shown to markedly increase cardiac differentiation (41). There are numerous of noncompensated functions between TGF- isoforms, which suggests that a precise combination of different TGF-s may be required to achieve proper cardiac differentiation. Thus it will be interesting to investigate in the near future potentially distinct effects on I_CaL by these isoforms.

It was recently shown that transgenic mice overexpressing TGF-1 exhibit an increased predisposition to develop AF (45). In that study, however, TGF-1 overexpression did not provoke significant alterations on action potential duration (APD; APs were triggered at 7 Hz). Thus predisposition to AF was associated to an increased fibrosis instead of potential electrophysiological effects. Unfortunately, Verheule et al. (45) did not investigate I_CaL or a potential loss in the capability of APD to adapt to rate (which would have suggested an in vivo regulation of I_CaL). Thus whether TGF-1 actually provokes in vivo electrical remodeling still remains to be elucidated.

As mentioned before, it is intriguing that TGF-1 affects only I_CaL from atrial but not ventricular myocytes. Interestingly, whereas transgenic mice overexpressing TGF-1 develop an increased vulnerability to AF (45), mutations in the gene encoding TGF-3 are linked to a human ventricular cardiomyopathy (3). Moreover, TGF-1 exerts opposite effects on atrial (inhibition) and ventricular (stimulation) hypertrophy (30). These previous observations, combined with our present results, could be interpreted to suggest that different TGF- isoforms might preferentially regulate either, atrial or ventricular failure.

In conclusion, the beneficial effects of certain neurohumoral blockers are partially explained by their ability to inhibit electrical remodeling (21, 33, 40). Nevertheless, whereas certain drugs can be successfully used to treat AF, they can often result in potentially lethal alterations in ventricular electrophysiology. Thus identification of the factors that selectively control ion channel expression in atrial versus ventricular myocytes could be advantageous to find a more effective treatment of both AF and heart failure.

	GRANTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
This work was supported by a Conacyt grant to GA (39512).

	ACKNOWLEDGMENTS

 
We thank Drs. Robert T. Dirksen and Gabriel Cota for continuous support. We also thank Dr. Georges Christé for advice and helpful comments, and Mario Rodriguez for excellent technical assistance.

	FOOTNOTES

 

G. Avila, Dept. of Biochemistry, Cinvestav, Mexico DF 007000, Mexico (e-mail: gavila{at}cinvestav.mx)

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.

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