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Pro-oxidant effect of transforming growth factor-β1 mediates contractile dysfunction in rat ventricular myocytes

Shumin Li, Xun Li, Hong Zheng, Bin Xie, Keshore R. Bidasee, George J. Rozanski
DOI: http://dx.doi.org/10.1093/cvr/cvm022 107-117 First published online: 1 January 2007

Abstract

Aims Transforming growth factor-β1 (TGF-β1) is a multifunctional cytokine that contributes to pathogenic cardiac remodelling via mechanisms that involve oxidative stress. However, the direct impact of TGF-β1 on contractile function of ventricular myocytes is incompletely understood.

Methods and results Reactive oxygen species (ROS) production and intracellular glutathione (GSH) were measured by fluorescence microscopy in isolated rat ventricular myocytes pretreated with TGF-β1 (0.1–10 ng/mL). In separate studies, video edge detection measurements were made to evaluate myocyte contractile function, and confocal microscopy was used to monitor evoked Ca2+ transients. TGF-β1 (1 ng/mL) for 3–4 h significantly increased ROS production by 90% (P < 0.05) and decreased GSH by 34% (P < 0.05) compared with control. These changes paralleled a significant decrease in the rate of myocyte shortening and relaxation by 33% and 43%, respectively (0.5 Hz; P < 0.05), whereas fractional shortening was not altered. Ca2+ transients in TGF-β1-treated myocytes were characterized by a delayed peak and slowing in the rate of decay but no change in peak Ca2+ amplitude. Increased ROS production and GSH depletion by TGF-β1 were prevented by an NAD(P)H oxidase inhibitor or a free radical scavenger, both of which significantly mitigated TGF-β1-induced myocyte contractile dysfunction. Moreover, pretreating myocytes with exogenous GSH or the GSH precursor N-acetylcysteine also prevented myocyte contractile impairment and abnormal Ca2+ transients elicited by TGF-β1.

Conclusion Our data suggest that TGF-β1-induced cardiomyocyte contractile dysfunction is associated with enhanced ROS production and oxidative alterations in Ca2+ handling proteins regulated by endogenous GSH.

Keywords
  • TGF
  • Redox
  • Glutathione
  • NAD(P)H oxidase
  • Oxidative stress

1. Introduction

Cellular dysfunction mediated by reactive oxygen species (ROS) contributes to pathophysiological processes in cardiovascular diseases where a critical role for oxidative stress has been demonstrated. It is known that cardiac tissues are rich sources of ROS, including superoxide anion (O2•−), hydrogen peroxide (H2O2), and hydroxyl radical (OH).1 These ROS are derived from multiple sources, including cellular oxidase complexes and the mitochondrial electron transport system.2 The mechanisms underlying the generation of ROS are complex and include indirect production stimulated by hormones, growth factors, and cytokines.3

TGF-β1 is a multifunctional cytokine that participates in a number of biological events such as proliferation, differentiation, apoptosis, and cell death.4 These events have been shown to be related to increased ROS formation and cellular redox imbalance, as indicated by the depletion of GSH, an essential regulator that maintains normal cellular redox status.57 The mechanisms underlying the generation of ROS induced by TGF-β1 appear to differ between various cell types, but recent evidence suggests the involvement of NAD(P)H oxidase complexes, a major source of myocardial ROS production.8,9

In heart, TGF-β1 synthesis is increased during the development and progression of cardiac hypertrophy, fibrosis, and myopathy, in which oxidative stress has been clearly documented.10,11 Recent studies show that inhibition of endogenous TGF-β1 prevents myocardial fibrosis and remodelling with corresponding alleviation of cardiac dysfunction in pressure-overloaded rats12 and post-infarction mice,13 implying that TGF-β1 plays an important role in the pathogenesis of heart failure. Although these reports provide evidence that TGF-β1 is associated with cardiac dysfunction, the underlying mechanisms are unclear.

The present study examined mechanisms of redox alteration and cellular dysfunction induced by TGF-β1 in isolated rat ventricular myocytes. Our data suggest that TGF-β1 increases myocyte ROS production and depletes intracellular GSH, which contribute to altered contractile function and Ca2+ transients. Furthermore, increased ROS production by TGF-β1 is correlated with increased activity of NAD(P)H oxidase and expression of membrane subunits of the NAD(P)H oxidase complex. Hence, the effects of TGF-β1 on myocyte function are abolished by NAD(P)H oxidase inhibitors and by exogenous GSH and N-acetylcysteine (NAC), a precursor of GSH. These results indicate that TGF-β1 contributes to myocyte contractile dysfunction in part by a pro-oxidant effect that impacts redox-mediated antioxidant mechanisms.

2. Methods

2.1 Cell isolation

This investigation conformed with the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996). Rats were given an overdose of pentobarbital sodium (100 mg/kg, i.p.) and single ventricular myocytes were dissociated from excised, perfused hearts by a collagenase digestion protocol described previously.14,15 The atria were discarded after the digestion procedure and dissociated myocytes from all ventricular regions were suspended in Dulbecco's modified Eagle's medium, seeded on laminin-coated glass coverslips, and incubated at 37°C until used, usually within 6 h of isolation. Myocytes used for measuring mechanical properties were not attached to coverslips. Unless stated otherwise, all reagents used in our experiments were purchased from Sigma-Aldrich (St Louis, MO, USA).

2.2 Measurement of ROS and intracellular GSH

Cellular levels of O2•− and OH were monitored by the fluorescent probe TEMPO-9-AC (Molecular Probes, Carlsbad, CA, USA) as previously described.16 Briefly, cells were loaded at room temperature with 30 µmol/L TEMPO-9-AC in a solution containing (in mmol/L): 138 NaCl, 4.0 KCl, 1.2 MgCl2, 1.0 CaCl2, 18 glucose, and 5.0 HEPES, pH 7.4. Cells were then transferred to a recording chamber mounted on the stage of inverted microscope and fluorescence intensity (excitation, 358 nm; emission, 424 nm) was collected with a photometer assembly (Photon Technology International, Lawrenceville, NJ, USA) through a 20 × 20 µm optical sampling window positioned over the center of each myocyte. ROS levels in treated myocytes were expressed as a percent of untreated controls using fluorescence intensities measured as counts/s per unit cell volume (µm3) as estimated previously.14

Intracellular GSH was measured using the fluorescent probe monochlorobimane (mBCl, Molecular Probes).14 For these studies, myocytes were suspended in the same solution as above containing 0.4 mmol/L mBCl plus 2 mmol/L probenecid to inhibit cellular export of the GSH–bimane adduct. Cells were loaded with mBCl at room temperature for 50–60 min and transferred to a recording chamber on an inverted microscope. Fluorescence intensity (excitation, 360 nm; emission, 460 nm) was collected as for TEMPO-9-AC experiments and [GSH] calculated as described previously14 from calibration curves generated with graded concentrations of GSH plus rat liver glutathione-S-transferase to generate the fluorescent GSH–bimane adduct. Measured fluorescence was converted to [GSH] in amol/µm3 using the calibration curve, the known dimensions of the sampling window and an estimate of cell thickness derived previously.14

2.3 Enzyme assays

Basal activities of two major enzymes that regulate GSH, glutathione reductase and γ-glutamylcysteine synthetase, were measured by standard methods.17,18 For glutathione reductase, myocyte suspensions were homogenized in 0.1 mol/L Tris buffer and centrifuged at 4°C (6000 g). A 200 µL aliquot of supernatant was added to a cuvette containing 0.2 mol/L KH2PO4 buffer plus (in mmol/L): 2 EDTA, 20 oxidized glutathione (GSSG), and 2 NADPH. The change in absorbance at 340 nm was monitored for 5 min at 30°C by spectrophotometer (ThermoSpectronic, Waltham, MA, USA). A milliunit (mU) of glutathione reductase activity was defined as the amount of enzyme catalyzing the reduction of 1 nmol NADPH per minute. To measure γ-glutamylcysteine synthetase activity, cells were prepared as above and a 50 µL aliquot of supernatant was added to a reaction mixture of 0.1 mol/L Tris buffer containing (in mmol/L): 150 KCl, 5 Na2-ATP, 2 phosphoenolpyruvate, 10 l-glutamate, 10 l-aminobutyrate, 20 MgCl2, 2 Na2-EDTA, and 0.2 NADH, plus 17 µg pyruvate kinase and 17 µg lactate dehydrogenase. The change in absorbance at 340 nm was monitored for 5 min at 37°C and activity was expressed in mU, defined as the amount of enzyme converting 1 nmol of NADH to NAD per minute. Protein concentration in all samples was measured using Coomassie blue and enzyme activities were expressed per milligram of protein.

NAD(P)H oxidase activity was assessed19 in suspensions of myocytes incubated with 250 µmol/L NADPH or NADH. The utilization of NADPH or NADH was monitored by the decrease in absorbance at 340 nm over 10 min. To verify specific oxidase activity, the rate of NADPH consumption inhibited by 30 µmol/L diphenyleneiodonium (DPI) was measured by adding the inhibitor 30 min before assay. All results were expressed in nmol/L of substrate/min/106 cells. Cell number was determined by counting with a haemocytometer and viability was assessed on the basis of exclusion of 0.4% Trypan blue.

2.4 Western blot analysis

The abundance of NAD(P)H oxidase subunits and selected Ca2+ handling proteins was analysed by western blotting. Specifically, cells were sonicated in buffer (10 mmol/L PBS, 1% N P-40, 0.5% sodium deoxycholate, and 0.1% SDS) containing 1 mmol/L PMSF and a protease inhibitor cocktail and centrifuged at 750 g for 10 min at 4°C. Forty micrograms of sample was separated by 10% SDS–polyacrylamide gel electrophoresis and the proteins transferred to a polyvinylidene difluoride membrane (Millipore, Billerica, MA, USA). The membranes were probed with primary antibodies against the following NAD(P)H oxidase subunits: p22phox, p47phox, p67phox (Santa Cruz Biotechnology, Santa Cruz, CA, USA), or gp91phox (BD Biosciences, San Diego, CA, USA). Sarcoplasmic reticulum Ca2+-ATPase and phospholamban were similarly probed with primary antibody (Affinity Bioreagents, Golden, CO, USA), while GAPDH (Santa Cruz) was used as the loading control for all analyses. The blots were detected with HRP-labelled rabbit anti-mouse IgG or goat anti-rabbit IgG polyclonal antibodies and quantified by densitometry by means of chemiluminescence (ECL, Amersham Bioscience).

2.5 Myocyte mechanical properties and Ca2+ transients

Mechanical properties of ventricular myocytes were measured using a video-based edge detection system (IonOptix, Milton, MA, USA).20,21 Briefly, cells were placed in a perfusion chamber mounted on the stage of an inverted microscopy and superfused at room temperature with the same external solution used for fluorescence studies. Cells were field-stimulated (10 V, 10 ms) through a pair of platinum wires placed on opposite sides of the chamber. Resting cell length, shortened cell length, and rates of shortening (−dL/dt) and re-lengthening (+dL/dt) were measured by IonWizard software (IonOptix). Ca2+ transients were measured with the fluorescent Ca2+ indicator Fluo-3-acetoxymethyl ester (Fluo-3 AM, Molecular Probes).20 Isolated myocytes attached to glass coverslips were loaded with 5 µmol/L Fluo-3 AM in culture medium for 30 min at 37°C. Cells were then washed with external solution to remove extracellular probe and placed in a chamber on the stage of a confocal microscope (LSM 410, Zeiss) equipped with an argon–krypton laser. During field-stimulation of myocytes (10 V, 10 ms) at room temperature, fluorescence measurements were made in line-scan mode (6 ms intervals), along the long axis of the cell (excitation, 488 nm; emission, 515 nm). Fluorescence signals (ΔF) were calculated as maximum fluorescence (Fmax) minus basal cell fluorescence (F0), normalized to F0: ΔF = (FmaxF0)/F0 expressed as arbitrary units/s. The time course of Ca2+ transients was assessed by measuring the time to peak and the duration at 50% of ΔF in the decay phase (T50 of decay). Since Ca2+ transients are triggered by the influx of Ca2+ through the sarcolemma, a subset of myocytes was studied using the whole-cell configuration of the patch–clamp technique as described previously,21 to assess changes in the properties of L-type Ca2+ channels.

2.6 Statistical analysis

Results are expressed as a mean ± SEM. Comparisons of two groups were made using a Student's t-test, whereas more than two groups were compared by ANOVA. When a significant difference among groups was indicated by the initial analysis, individual paired comparisons were made using a Student–Newman–Keuls t-test. Differences were considered significant at P < 0.05.

3. Results

3.1 Oxidative stress and contractile dysfunction elicited by TGF-β1

The pro-oxidant effects of TGF-β1 on ventricular myocytes were examined in initial experiments using TEMPO-9-AC to monitor cellular ROS production.16 Figure 1A shows the time-dependent change in TEMPO-9-AC fluorescence intensity elicited by 1 ng/mL TGF-β1. Normalized fluorescence reached a maximum level by 3–4 h of TGF-β1 incubation and remained significantly elevated for up to 6 h. In parallel experiments using mBCl, intracellular levels of GSH were also monitored as an index of oxidative stress. Figure 1B shows that the same concentration of TGF-β1 elicited GSH depletion that lagged slightly behind the increase in ROS production. By comparison, [GSH] in untreated myocytes did not change significantly over the incubation periods studied: mean [GSH] at 0, 1–2, 3–4, and 5–6 h was 3.9 ± 0.1, 3.9 ± 0.1, 3.8 ± 0.2, and 3.8 ± 0.2 amol/µm3, respectively (P > 0.05). Concentration–response data in Figure 1C illustrate that TGF-β1 (3–4 h) at less than 1 ng/mL produced correspondingly lower levels of TEMPO-9-AC fluorescence (filled circles) but that higher concentrations did not produce greater fluorescence. Moreover, the TGF-β1 concentration–response curve for [GSH] mirrored the curve for ROS production. Given the time- and concentration-dependent responses to TGF-β1 shown in Figure 1AC, we chose an experimental protocol of 1 ng/mL TGF-β1 for 3–4 h for all subsequent experiments. Finally, Figure 1D illustrates that increased ROS production and GSH depletion elicited by TGF-β1 were blocked by 1 µg/mL actinomycin D or 100 µmol/L cycloheximide, indicating that the pro-oxidant effect of TGF-β1 was mediated by synthesis of new protein.

Figure 1

Alterations in myocyte levels of ROS and GSH by TGF-β1. (A and B) Myocytes were incubated with 1 ng/mL TGF-β1 for up to 6 h and the cellular levels of ROS and GSH were measured by fluorescence microscopy using the probes TEMPO-9-AC and mBCl, respectively. (C) ROS and GSH levels were measured as a function of TGF-β1 concentration (3–4 h treatment). (D) Pretreatment of myocytes with actinomycin D (1 µg/mL, ActD) or cycloheximide (100 µmol/L, CHX) prevented changes in GSH content (grey bar) and ROS levels elicited by TGF-β1. Each bar represents the mean from 30 to 40 myocytes isolated from five rat hearts. *P < 0.05 vs. control.

To determine if TGF-β1 depleted GSH by directly inhibiting regulatory pathways, myocyte suspensions were assayed for two major enzymes that control endogenous GSH levels. First, we found that the activity of γ-glutamylcysteine synthetase, which controls de novo GSH synthesis, was not affected by TGF-β1 (TGF-β1-treated, 111.2 ± 21.3 mU/mg protein, n = 5; control, 119.0 ± 13.4 mU/mg protein, n = 5; P > 0.05). Moreover, the enzyme glutathione reductase which generates GSH from its oxidized form, GSSG, was also not significantly different between TGF-β1-treated (52.6 ± 10.3 mU/mg protein, n = 5) and control (42.5 ± 6.7 mU/mg protein, n = 5; P > 0.05) myocytes. These data suggest therefore that TGF-β1 at the test concentration used in these experiments did not directly inhibit GSH-related regulatory pathways.

The functional effects of TGF-β1 on ventricular myoyctes were explored by measuring the rate and extent of myocyte shortening and re-lengthening by video edge detection during field stimulation. As illustrated by the superimposed traces in Figure 2A, TGF-β1 slowed the rate of cell shortening and re-lengthening with little change in extent of shortening compared with untreated myocytes. In parallel confocal studies measuring Ca2+ transients (Figure 2B), TGF-β1-treated myocytes showed a prolonged time to peak Ca2+ and T50 of decay compared with control. The rising phase of the Ca2+ transient in treated myocytes was clearly biphasic in 25 of 30 cells studied and was characterized by a slowing in the development of the peak or a secondary peak as shown in the inset of Figure 2B. Nevertheless, TGF-β1 did not affect the maximum rate of rise of the Ca2+ transient (TGF-β1-treated, 69.5 ± 2.3 f.a.u./s, n = 12; untreated 75.6 ± 4.4 f.a.u./s, n = 13; P > 0.05) or its peak amplitude. Table 1 summarizes the alterations in myocyte mechanical properties (0.5, 1, and 2 Hz) and Ca2+ transients (0.5 Hz) elicited by TGF-β1. The rate of shortening (−dL/dt) and re-lengthening (+dL/dt) in untreated myocytes was marked by a frequency-dependence that was much smaller or absent in TGF-β1-treated myocytes. Hence, abnormal myocyte mechanical properties elicited by TGF-β1 were unmasked at low stimulus frequency.

Figure 2

Myocyte contractile dysfunction elicited by TGF-β1. (A) Representative traces of mechanical responses in a control and TGF-β1-treated (1 ng/mL, 3–4 h) myocyte measured by edge-detection (0.5 Hz). (B) Comparison of the intracellular Ca2+ transient in a TGF-β1-treated and control myocyte (0.5 Hz) measured by confocal microscopy. Inset figure shows a second example from a TGF-β1-treated myocyte illustrating a secondary peak of the Ca2+ transient.

View this table:
Table 1

Effect of in vitro TGF-β1 treatment on myocyte mechanical properties and Ca2+ transients

Frequency (Hz)−dL/dt (µm/s)+dL/dt (µm/s)FS
Mechanical properties
Untreated0.5211.0 ± 27.6166.7 ± 20.00.87 ± 0.023
TGF-β10.5141.7 ± 20.1*95.0 ± 19.0*0.90 ± 0.030
Untreated1.0165.1 ± 21.4128.2 ± 15.60.89 ± 0.01
TGF-β11.0133.5 ± 12.196.7 ± 10.10.91 ± 0.03
Untreated2.0153.3 ± 13.4126.6 ± 16.00.90 ± 0.01
TGF-β12.0130.3 ± 23.297.1 ± 12.50.92 ± 0.03
Frequency (Hz)Time to peak (ms)Peak Ca2+ (f.a.u.)T50 decay (ms)
Ca2+ transients
Untreated0.537.7 ± 2.32.1 ± 0.16352.2 ± 17.3
TGF-β10.545.8 ± 2.1*1.9 ± 0.12410.6 ± 24.0*
  • Values are means ± SEM. Mean data for mechanical studies based on 30–35 myocytes. Resting length for untreated and TGF- β1-treated myocytes was 125.5 ± 5.6 and 124.8 ± 7.3 µm, respectively (P > 0.05). Mean data for Ca2+ transients based on 25–30 myocytes. In all experiments, myocytes were treated with 1 ng/mL TGF-β1 for 3–4 h. −dL/dt, maximum rate of myocyte shortening; +dL/dt, maximum rate of re-lengthening; FS, fractional shortening, calculated as resting minus shortened cell length normalized to resting length; f.a.u., fluorescence intensity in arbitrary units measured as (FmaxF0)/F0 (where F is fluorescence). *P < 0.05 compared with untreated myocytes.

3.2 Effects of ROS scavengers and the role of NAD(P)H oxidase

The functional impact of increased ROS production by TGF-β1 was also explored using structurally distinct ROS scavengers and enzyme inhibitors. In initial fluorescence microscopy studies, ROS levels were measured in myocytes pretreated for 1 h with radical scavengers or an NAD(P)H oxidase inhibitor before adding TGF-β1 for 3–4 h. Figure 3A shows that the radical scavenger Mn(III)terakis(1-methyl-4-pyridyl)porphyrin-pentachloride (MnTPyP, 50 µmol/L) and the NAD(P)H oxidase inhibitor DPI (30 µmol/L) blocked the increase in ROS production induced by TGF-β1. Similarly, GSH depletion by TGF-β1 was also prevented by MnTPyP and DPI (Figure 3B). It should also be noted that the radical scavenger Tiron (20 mmol/L) gave quantitatively similar results as MnTPyP and DPI, whereas the xanthine/xanthine oxidase inhibitor oxypurinol (100 µmol/L) or the NOS inhibitor L-NAME (100 µmol/L) failed to prevent the pro-oxidant and GSH-depleting effect of TGF-β1 (data not shown). In parallel studies examining mechanical properties of myocytes, MnTPyP and DPI markedly blunted the slowing of the rate of cell shortening and re-lengthening in myocytes treated with TGF-β1 (Figure 3C and D).

Figure 3

Effects of ROS inhibitors on myocytes treated with TGF-β1. MnTPyP (50 µmol/l) and DPI (30 µmol/l) prevented the increase in ROS levels (A) and GSH depletion (B) induced by 1 ng/mL TGF-β1. (C and D) The rates of myocyte shortening (−dL/dt) and re-lengthening (+dL/dt) were diminished by TGF-β1 alone and prevented by MnTPyP or DPI. Each bar represents the mean from 30 to 35 myocytes from five rat hearts. *P < 0.05 vs. control.

The inhibition of pro-oxidant and mechanical effects of TGF-β1 by DPI suggests that NAD(P)H oxidase was a major source of ROS generation. To address this mechanism further, the rate of NADPH utilization by suspensions of isolated myocytes was used as a measure of NAD(P)H oxidase activity. Figure 4A shows representative experiments illustrating that myocytes pretreated with TGF-β1 (filled circles) utilized exogenous NADPH at a much faster rate than control myocytes. Figure 4B compares the rates of NADPH consumption in cell suspensions under various conditions clearly showing the profound increase in utilization by TGF-β1-treated myocytes and the blockade of this effect by pretreatment with 100 µmol/L cycloheximide or 30 µmol/L DPI. When NADH was used in place of NADPH, its rate of utilization was not significantly different between TGF-β1-treated (0.46 ± 0.1 nmol/L min/106 cells) and untreated (0.43 ± 0.1 nmol/L min/106 cells) myocytes (P > 0.05).

Figure 4

Increased NADPH utilization by myocytes treated with 1 ng/mL TGF-β1. (A) Rate of NADPH consumption was greater in TGF-β1-treated myocytes than in untreated cells. (B) Increased NADPH utilization was abolished by DPI (30 µmol/L) or cycloheximide (CHX, 100 µmol/L). Each bar represents the mean from 30 to 35 myocytes from five rat hearts. *P < 0.05 vs. control.

To determine whether TGF-β1 affected protein abundance of NAD(P)H oxidase subunits, western blots were analysed from myocyte suspensions treated with TGF-β1 or left untreated for 3–4 h. Figure 5A shows representative immunoblots from four experiments illustrating that the expression of two membrane subunits of the NAD(P)H oxidase complex, gp91phox and p22phox, were increased by TGF-β1 compared with controls. By comparison, the expression of two cytosolic subunits, p47phox and p67phox, was not different between groups of myocytes. Mean data from these analyses, which are summarized in Figure 5B and C, show that the protein abundance of the membrane subunits (gp91phox and p22phox) after TGF-β1 treatment was increased from control by 2.1- and 1.8-fold, respectively (P < 0.05), whereas the cytosolic subunits (p47phox and p67phox) were not different from control.

Figure 5

Protein abundance of NAD(P)H oxidase subunits in TGF-β1-treated myocytes. (A) Representative western blots of gp91phox, p22phox, p47phox, and p67phox in suspensions of control and TGF-β1-treated myocytes. (B) Mean densitometry measurements of gp91phox (left panel) and p22phox expressed relative to GAPDH. *P < 0.05 vs. control. (C) Mean densitometry measurements of p47phox (left panel) and p67phox expressed relative to GAPDH. Each bar represents the mean from myocyte suspensions isolated from three to seven rat hearts.

3.3 Antioxidant role of GSH and myocyte contractile function

The role of GSH in regulating cell function was explored in greater detail by measuring TEMPO-9-AC fluorescence in myocytes pretreated for 1 h with 10 mmol/L GSH or NAC (a precursor of GSH) before exposure to TGF-β1. Figure 6A shows that both GSH and NAC prevented the increase in ROS production elicited by TGF-β1 compared with control levels. By contrast, the reducing agent dithiothreitol did not prevent TGF-β1-mediated increased ROS production. When the functional effects of exogenous GSH and NAC on myocyte contractile properties were examined, the decreases in the rates of cell shortening and re-lengthening induced by TGF-β1 were also prevented (Figure 6B and C). Finally, to further examine the impact of GSH on cell contractile function, intracellular Ca2+ transients were measured. Figure 7 shows that GSH and NAC prevented TGF-β1-induced changes in time to peak Ca2+ (A) and T50 decay (B). It should also be noted that in parallel experiments in which GSH or NAC was added after TGF-β1 treatment (3–4 h), the altered contractile and Ca2+ transient kinetics were normalized (data not shown). These findings suggest that oxidative modifications of Ca2+ handling proteins by TGF-β1 are reversed by mechanisms that directly or indirectly involve intracellular GSH.

Figure 6

Effects of reducing agents on myocytes treated with TGF-β1. (A) GSH or NAC (10 mmol/L) prevented the increase in ROS production elicited by TGF-β1, whereas dithiotheitol (DTT) had no effect. (B and C) Changes in the rates of myocyte shortening (−dL/dt) and re-lengthening (+dL/dt) induced by TGF-β1 were prevented by pretreatment with GSH or NAC. Each bar represents the mean from 25 to 30 myocytes isolated from five rat hearts. *P < 0.05 vs. control.

Figure 7

Effects of reducing agents on Ca2+ transients in myocytes treated with TGF-β1. GSH and NAC abolished the increases in time to peak (A) and T50 decay (B) elicited by TGF-β1. Each bar represents the mean from 25 to 30 myocytes isolated from five hearts. *P < 0.05 vs. control.

3.4 Protein targets of TGF-β1-induced ROS

The mechanisms of TGF-β1-induced mechanical dysfunction were explored in greater detail by experiments that focused on changes in the properties of Ca2+ handling proteins, particularly voltage-gated Ca2+ channels, ryanodine receptors (RyR2), and sarco-endoplasmic reticulum Ca2+-ATPase (SERCA2a). First, to assess changes in L-type Ca2+ channels, we conducted whole-cell patch–clamp experiments to measure the peak amplitude and kinetics of the L-type Ca2+ current (ICa). In these studies, TGF-β1 did not alter the current–voltage relationship of ICa or its inactivation kinetics. Specifically, peak ICa density measured at 0 mV was −27.4 ± 2.0 and −26.0 ± 2.4 pA/pF in TGF-β1- treated (n = 13) and untreated (n = 11) myocytes, respectively (P > 0.05). The kinetics of inactivation of ICa at 0 mV was also not affected by TGF-β1 (data not shown). Secondly, spontaneous Ca2+ sparks were analysed in quiescent myocytes loaded with Fluo-3 AM to assess possible changes in the function of RyR2 to release Ca2+ from the sarcoplasmic reticulum (SR). However, neither the frequency nor the peak amplitude of Ca2+ sparks was altered by TGF-β1: mean spark frequency in TGF-β1-treated (n = 18) and control (n = 18) myocytes was 6.5 ± 0.8 and 5.9 ± 0.9 sparks/50 µm/s (P > 0.05), whereas mean peak Ca2+ amplitude was 1.59 ± 0.10 (n = 32) and 1.64 ± 0.17 (n = 17) f.a.u., respectively (P > 0.05). Thirdly, the total amount of releasable Ca2+ inside the SR was estimated in Fluo-3 AM-loaded myocytes that were field-stimulated with a train of four beats (0.1 Hz) and then challenged 30 s after the last stimulus with 10 mM caffeine.21 The mean peak amplitude of caffeine-induced Ca2+ release, a measure of releasable SR Ca2+, was 2.18 ± 0.17 and 2.17 ± 0.27 f.a.u. in TGF-β1-treated (n = 7) and untreated (n = 5) myocytes, respectively (P > 0.05).

Western blot analyses were also done in treated and untreated myocytes to examine the phosphorylation status of phospholamban, an important regulator of SERCA2a function to transport cytosolic Ca2+ back into the SR. Figure 8A shows that the amount of phosphorylated phospholamban in TGF-β1-treated myocytes was significantly less than in control cells. In parallel analyses, the level of dephosphorylated phospholamban was significantly increased in treated myocytes compared with untreated cells (Figure 8B), whereas SERCA2a protein abundance was not affected by TGF-β1 (Figure 8C).

Figure 8

Phosphorylation status of phospholamban in TGF-β1-treated myocytes. Protein abundance of phosphorylated (P-PLB, A) and dephosphorylayed (D-PLB, B) phospholamban was examined by western blot in myocyte suspensions from four hearts treated with TGF-β1. *P < 0.05 vs. control. (C) Protein abundance of SERCA2a measured in cell suspensions from three hearts. Mean densitometry values are expressed relative to GAPDH.

4. Discussion

4.1 Cardiac dysfunction and oxidative stress elicited by TGF-β1

TGF-β, a locally generated cytokine, has three isoforms (TGF-β1, -β2, and -β3) of which TGF-β1 is the major isoform in heart.10 Several extrinsic factors such as mechanical stress, ischaemia, angiotensin II, and TNF-α are known to activate TGF-β1 synthesis in myocytes,10,22 but its role in cardiac pathology is not fully understood. On the one hand, increased TGF-β1 expression is implicated in the process of structural remodelling that underlies ventricular dysfunction in chronic disease states.23 Thus, inhibition of TGF-β1 by a soluble TGF-β-type II receptor 3 days post-myocardial infarction in mice improves contractile function and decreases mortality by inhibiting myocardial fibrosis.13 Blockade of TGF-β1 by anti-TGF-β neutralizing antibodies also inhibits myocardial fibrosis and prevents diastolic dysfunction in pressure-overloaded rats.12 However, endogenous TGF-β1 may also exert cardioprotective effects. For example, pretreating mice with the soluble TGF-β-type II receptor to block TGF-β1 signalling exacerbates ventricular dysfunction and increases mortality 24 h after myocardial infarction compared with untreated, infarcted mice.24 When initiated several days after infarction, TGF-β1 blockade inhibits myocyte hypertrophy and interstitial fibrosis compared with untreated mice.24 At the myocyte level, our study shows that exogenous TGF-β1 causes contractile dysfunction manifest as a prolongation in the rates of cell shortening and re-lengthening (Figure 2 and Table 1), which is consistent with previous in vivo studies.12 Nevertheless, the cellular mechanisms mediating impaired myocyte function by TGF-β1 are still unclear.

It is well known that oxidative stress is a central mechanism in the pathogenesis of heart failure from different etiologies,1,2,11 but the factors initiating oxidative stress are not fully known. Recent experiments suggest that NAD(P)H oxidases are a major source of O2•− generation in myocytes as compared with xanthine oxidase, arachidonic acid metabolism, or mitochondrial oxidases.9 Cardiac NAD(P)H oxidases are complexes that consist of at least two constitutive membrane subunits (gp91phox and p22phox) and four cytosolic subunits (p47phox, p67phox, p40phox, and Rac1).9 Recent studies indicate that activation of this oxidase may occur by phosphorylation and translocation of cytosolic subunits25,26 or by an increase in the synthesis of one or more subunits.27,28 Once activated, NAD(P)H oxidase produces O2•− which is converted to H2O2,27,28 and these ROS may impair cellular function via oxidation of membranes, proteins, or DNA. Our present studies testing the functional impact of ROS scavengers and NAD(P)H oxidase inhibitors (Figure 3) suggest a major role for this oxidase in the cellular effects of TGF-β1 that is mediated by an increase in subunit expression (Figure 5). Moreover, our finding that both cycloheximide and actinomycin D abolished the pro-oxidant effect of TGF-β1 (Figures 1D and 4B) suggests that increased expression of gp91phox and p22phox was due to up-regulation of transcriptional and/or translational pathways for these subunits. However, it cannot be excluded that other mechanisms may participate in TGF-β1-induced increase in ROS generation.

Oxidative stress is not only caused by generating excess ROS but also by attenuating the capacity of antioxidants. Although our experiments found that TGF-β1 treatment was correlated with an increase in ROS levels, the effect of this cytokine on the antioxidant potential of myocytes is still unknown. GSH is a major thiol-based antioxidant that directly scavenges free radicals and serves as co-factor for the breakdown of H2O2 by glutathione peroxidase.29,30 GSH levels are controlled by γ-glutamylcysteine synthetase and glutathione reductase,27 and recent studies in non-cardiac cells suggest that TGF-β1 can deplete cellular GSH by directly inhibiting these enzymes.6 Our studies found that while [GSH] was dose-dependently decreased by TGF-β1 (Figure 1), the activities of γ-glutamylcysteine synthetase and glutathione reductase were not different from control. Hence, these results suggest that TGF-β1 depletes cellular GSH secondary to an increase in ROS production.31 This is supported by data in Figure 1A and B which show that the TGF-β1-mediated decline in GSH begins slightly after the marked increase in ROS production.

Our data also suggest that therapies that maintain cellular GSH levels are protective against the contractile dysfunction elicited by TGF-β1. When intracellular GSH levels drop in pathophysiological states, one approach to maintain them is to supplement cells with exogenous GSH, as in our experiments (Figures 6 and 7). However, mammalian cells do not take up intact GSH in significant amounts.32 Instead, it is proposed that exogenous GSH increases cellular uptake of cysteine, the rate-limiting amino acid in GSH synthesis, by its initial extracellular enzymatic degradation followed by amino acid uptake and re-synthesis of GSH in the cytoplasm.32 Increased cysteine availability for GSH synthesis is also the proposed mechanism to explain the cellular effects of NAC,33 and we have shown that NAC increases intracellular [GSH] in depleted myocytes.20 Nevertheless, while our present data suggest an antioxidant role for GSH, it is not known whether this essential tripeptide directly scavenges ROS or increases ROS degradation via glutathione peroxidase.30 Also we cannot completely rule out the possibility that exogenous reducing agents blocked the TGF-β1 effect by directly inactivating TGF-β1 molecules, rather than by an antioxidant mechanism.34 However, our finding that the reducing agent dithiothreitol did not block the increase in ROS production by TGF-β1 (Figure 6A) argues against this hypothesis. Nevertheless, further studies are required to determine if exogenous reducing agents block other molecular steps in TGF-β1 signalling, such as ligand binding capacity.34

4.2 Redox-sensitivity of Ca2+ handling proteins

Oxidative stress is known to elicit cellular Ca2+ dysregulation and contractile impairment of the myocardium.35 Normal contractile activity at the myocyte level is controlled by the interplay of several Ca2+ handling proteins, including L-type Ca2+ channels, RyR2, SERCA2a, Na+/Ca2+ exchanger (NCX), and contractile myofilaments. Experimental studies suggest that most of these proteins are susceptible to oxidation by ROS, which may explain cardiac dysfunction associated with oxidative stress.36,37 The evoked Ca2+ transients measured in TGF-β1-treated myocytes in the present study were characterized by a delayed peak and slow decay with no significant change in peak Ca2+ amplitude compared with control (Figure 2B and Table 1). These changes largely paralleled alterations in the kinetics of evoked mechanical responses. Our voltage–clamp experiments suggest that Ca2+ entering through voltage-gated channels, which triggers the Ca2+ transient, was not affected by TGF-β1, but the increased time to peak Ca2+ implies that Ca2+ mobilization from intracellular stores was altered. In terms of SR Ca2+ release, the tetrameric RyR2 channel has several cysteine residues whose thiol groups are accessible to oxidizing or nitrosylating molecules that impact SR function,38,39 and it has been shown that high ROS levels decrease the rate of Ca2+ release by RyR2.39 However, significant oxidative alteration of RyR2 function was not evident in our studies since the maximal rate of rise and peak of Ca2+ transients were similar in TGF-β1-treated and control myocytes (Figure 2B), and there were no differences in frequency or amplitude of spontaneous Ca2+ sparks. If the functions of Ca2+ and RyR2 channels were not altered by TGF-β1, then it is likely that the depressed rate of mechanical shortening (Figures 3C and 6B) was due in part to a decrease in the Ca2+ sensitivity of contractile myofilaments or slower crossbridge cycling. Nevertheless, a distinguishing feature of TGF-β1-treated myocytes was a pronounced delay in the Ca2+ peak or a secondary peak (Figure 2B) which may also have contributed to the slowing in the rate of shortening. The cellular basis for this characteristic requires further study but it may reflect depressed function of a small population of RyR2 channels or slow release of Ca2+ from non-SR stores.40

The prolonged decay of Ca2+ transients with TGF-β1 is consistent with slowed re-lengthening observed in mechanical studies and is likely mediated by changes in SERCA2a function and/or NCX. First, ROS may directly decrease SERCA2a activity and SR Ca2+ uptake by blocking ATP binding or through membrane peroxidation.41,42 Decreased SERCA2a activity may also involve the small, regulatory phosphoprotein phospholamban.43 Indeed, we found that TGF-β1 decreased levels of phosphorylated phospholamban (Figure 8A), which would be expected to impede its dissociation from SERCA2a and potentiate its inhibitory influence. However, while oxidative alterations in phospholamban may have decreased the rate of Ca2+ uptake by SERCA2a, it did not significantly affect overall Ca2+ uptake, since the releasable pool of SR Ca2+, as assessed by caffeine challenge, was not altered by TGF-β1. Secondly, delayed Ca2+ transients can be expected from a decrease in NCX activity, but experimental studies generally show that transporter activity in myocytes is increased by oxidants, which would be expected to accelerate the decay of Ca2+ transients. Hence, our data indicate that increased ROS generation by TGF-β1 contributes to SERCA2a dysfunction via phospholamban, thereby slowing the rate of Ca2+ uptake and myocyte relaxation in a redox-sensitive manner. We cannot however rule out the possibility that ROS depressed the rate of Ca2+ uptake in our myocytes by direct inhibition of SERCA2a.41,42

In summary, redox imbalance induced by TGF-β1 in cardiac myocytes is mediated by increased ROS generation and depletion of intracellular GSH. The generation of ROS is derived mainly from NAD(P)H oxidase which underlies contractile dysfunction and altered Ca2+ transients elicited by TGF-β1. Moreover, the reversal of functional deficits elicited by TGF-β1 by exogenous GSH or NAC suggests that endogenous GSH plays a key antioxidant role to protect Ca2+ handling proteins from oxidative damage.

Acknowledgements

This work was supported by grants from the National Heart, Lung, and Blood Institute (HL 66446, GJR; HL 085061, KRB) and the American Diabetes Association (1-06-RA-11, KRB).

Conflict of interest: none declared.

References

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