Abstract

Recent studies have suggested that puerarin may attenuate cardiac hypertrophy in mice; however, the underlying mechanism remains unclear. To investigate the role of puerarin in transforming growth factor beta 1 (TGF-β₁)-induced cardiac hypertrophy and the underlying mechanisms. Primary neonatal rat cardiomyocytes (NRCMs) were isolated form the heart of neonatal Wistar rats (1- to 2-day-old) and treated with different doses of puerarin (0,0.1,1or 5 g/L) in the presence of TGF-β₁ (3 μg/L) to investigate the effect of puerarin on TGF-β₁-induced hypertrophic changes and Smad protein alterations. RNA and protein biosynthesis in NRCMs were evaluated by synpropidium iodide staining and [³H]-leucine incorporation, respectively. Gene expression alterations were determined using quantitative real-time PCR and Western blot analysis. Compared with Smad2 knockdown, puerarin treatment (1 g/L) exhibited similar but stronger effects in abrogating TGF-β₁-induced RNA and protein biosynthesis as well as fetal gene upregulation in NRCMs. In addition, puerarin treatment (1 g/L) could remarkably reverse TGF-β₁-induced upregulation of Smad protein expression in NRCMs, suggesting that deactivation of the Smad signaling is possibly involved in the antihypertrophic role of puerarin. Our data suggest that puerarin may protect cardiomyoctyes against TGF-β₁-induced hypertrophic responses and Smad signaling activation, providing basic dosage information and possible targets for puerarin treatment in animal models.

1. Introduction

Cardiac hypertrophy refers to increases in the heart size and mass resulting from adaptive responses to increased hemodynamic stress. It is characterized by enlargement of cardiomyocytes, enhancement of protein synthesis, and reactivation of fetal genes like atrial natriuretic factor (ANF) and β-myosin heavy chain (β-MHC) in cardiomyocytes. Pathological cardiac hypertrophy is an independent risk factor for cardiovascular morbidity and mortality, leading to cardiac dysfunction, heart failure, and even sudden death ^{1, 2, 3}. Currently, cardiac hypertrophy therapy is focused on blocking the hypertrophic signaling pathways ⁴, including sarcomeric Z-disc and its associated proteins that drive mechanical stress-induced signal transduction, the insulin-like growth factor-1 and transforming growth factor beta (TGF-β) signaling that are responsible for the eutrophic myocyte growth, and the G-protein-coupled receptors signaling that is linked to the progression to heart failure ^{5, 6}. Exploring therapeutic agents that target the molecules involved in these pathways would generate effective remedies for patients with cardiac hypertrophy.

Puerarin(7,4′-dihydroxyisoflavone-8β-glucopyranoside), a major active ingredient in the Chinese medicine Pueraria Radix isolated from the kudzu root (Pueraria lobota), exhibits therapeutic effects on various disorders, including hypertension ⁷, myocardial ischemia ⁸, diabetes mellitus ⁹, and cerebral ischemia ¹⁰, due to its antioxidant ¹¹, antiplatelet ¹², antiinflammatory ¹³, antiarrhythmic ¹⁴, and antidiabetic ¹⁵ properties ¹⁶. Recent studies have suggested that puerarin can effectively inhibit cardiac function impairments, cardiac fibrosis, heart size increase, and fetal gene upregulation in angiotensin II- or pressure overload-induced hypertrophy in mice ^{17, 18, 19, 20, 21}. The antihypertrophic effect of puerarin has been linked to the blockade of phosphoinositide 3-kinase (PI3K), c-Jun N-terminal kinase (JNK), extracellular signal–regulated kinases (ERK), p38 mitogen-activated protein kinase (MAPK), and nuclear factor-kappa B (NF-κB) signaling pathways. Although the TGF-β/Smad signaling has also been shown to promote cardiac hypertrophy development ^{22, 23, 24, 25}, little is known about its involvement in the antihypertrophic effect of puerarin.

In the present study, we isolated primary neonatal rat cardiomyocytes (NRCMs) from the heart of neonatal Wistar rats to explore the role of puerarin in TGF-β₁-induced hypertrophic changes and the underlying mechanism. Our results suggest that deactivation of the Smad signaling might be involved in the protective role of puerarin against TGF-β₁-induced hypertrophic changes in NRCMs, providing basic dosage information and possible targets for puerarin treatment in animal models.

2. Materials and Methods

Neonatal Wistar rats (1-to2-day-old) were obtained from the Medical Laboratory Animal Center of Sun Yat-Sen University (Guangzhou, China). All animal care and procedures were approved by the Ethics Committee of Guangzhou Medical University (Guangzhou, China). All animal experiments were performed in accordance with the guidelines for the use of laboratory animals of Guangzhou Medical University (EC Directive 86/609/EEC).

Neonatal Wistar rats were euthanized via hypothermia. The hearts were immediately excised, and the ventricles were dissected. NRCMs were isolated using 0.25% trypsin (Gibco, Maryland, USA) and purified as previously described ²⁶. The cells were resuspended in Dulbecco’s modified Eagles’s medium (DMEM, Gibco) containing 20% fetal calf serum and 1% penicillin-streptomycin and plated in tissue culture dishes for 2 h to remove noncardiomyocyte cells. In the first part the isolated NRCMs were cultured in DMEM supplemented with 10% FBS and 0.1 mM bromide oxy-uridine and were seeded at a low density (1×10⁵ cells/mL) for TGF-β₁ treatment or a high density (1×10⁶ cells/mL) for other experiments. After a 24-h starvation, NRCMs were treated with TGF-β₁ (0.1, 1, 3, or 10 μg/L; Calbiochem, San Diego, CA, USA), followed by analyses described below.

Then in the second part Puerarin (0.1, 1, or 5 g/L; Great Sun Biochemical Pharmaceutical Co., Ltd., Guangzhou, China), or Smad2siRNA (0.6, 1.0, or 1.4 μL; Invitrogen, Carlsbad, CA, USA) were individually or in combination treated to NRCMs with TGF-β₁ 3μg/L, followed by analyses described below.

NRCMs (10⁶ cells/well) were plated in a 6-well plate on day 1 and transfected with different amounts of Smad2siRNA (0.6, 1.0, or 1.4 μL) on day 4 using Lipofectamin^TM2000 (Invitrogen) according to the manufacturer’s protocol. At 24 h after transfection, cells were treated with TGF-β₁ (3 μg/L) alone or in combination with puerarin (1 g/L). Cells were collected at 24 h after treatment for gene expression detection.

Cellular RNA was stained with RNA-sensitive fluorescence PI (Thermo Fisher Scientific, MD, USA) after DNase treatment ^{27, 28}. Briefly, the NRCMs were treated with TGF-β₁ (0, 0.1, 1, 3, or 10 μg/L) in the first part, or TGF-β₁ 3μg/L in combination with puerarin (0.1, 1, or 5 g/L) for 24 h in the second part, followed by Hanks' balanced salt solution (HBSS) rinses and 75% ethanol fixation for 60 min. The ethanol-fixed cells were rinsed with HBSS and incubated in a solution containing DNase (1 mg/mL), sucrose (0.25 mol/L), MgCl₂ (5 mmol/L), and Tris-HCL (20 mmol/L, pH 6.5) at 36 ℃ for 40 min. After incubation, 1 mL of HBSS containing PI (0.05 mg/mL) was added to each well and incubated for 30 min. The RNA contents were determined by measuring the fluorescent signals in 5 randomly selected fields in each well using a fluorescence conversion microscope (1500X/CC; Leica, Germany).

[³H]-leucine incorporation was measured as previously described ²⁸. NRCMs were seeded at a density of 10⁶ cells/well in 24-well plates and cultured overnight. After a 24- h starvation, NRCMs were treated with different concentrations of TGF-β₁ (0, 0.1, 1, 3, or 10 μg/L) alone or TGF-β₁ 3μg/L in combination with puerarin (0.1, 1, or 5 g/L) for 24 h and incubated with one microcurie per-milliliter [³H]-leucine (Chinese Atomic Research Institute, Beijing, China) in leucine-free medium for additional 12 h. The radioactivity incorporated into the trichloroacetic acid precipitable component was determined by β-scintillation counting.

Total RNA was isolated using Trizol reagent (Invitrogen) and subjected to qRT-PCR with an Access RT-PCR system (Invitrogen). The qRT-PCR reactions were carried out using a Quanti-Tect^TM SYBR Green PCR Kit (Qiagen, Hilden, Germany). The PCR primers were as follows: sense primer for ANF, 5′-AAA GCA AAC TGA GGG CT-3′; antisense primer for ANF, 5′-GGG ATC TTT TGC GAT CT-3′; sense primer for β-MHC, 5′-TGC AGT TAA AGG TGA AGG C-3′; antisense primer for β-MHC, 5′-CAG GGC TTC ACA GGC AT-3′; sense primer for Smad2, 5′ -GGG AAG TGT TTT GCC GAG TG-3′; antisense primer for Smad2, 3′ -TAG GGT GGT CCG ACA TTG GA-5′; sense primer for GAPDH, 5′- CGG AGT CAA CGG ATT TGG TGG TAT-3′; antisense primer for GAPDH, 5′-AGC CTT CTC CAT GGT GGT GAA GAC-3′.

NRCMs were lysed in lysis buffer (Cell Signaling Technology, Danvers, MA, USA) on ice.After centrifugation for 2 min at 12,000 g, the protein concentration was measured using a Bio-Rad protein assay kit (Bio-Rad, Hercules, CA, USA). Equal amounts of proteins were subjected to 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis, and the proteins were transferred to PVDF membranes. The membranes were blocked by 3% BSA in TBST (10 mm Tris, 0.1 M NaCl, 0.1% Tween-20, pH 7.4) for 1.5 h. Following the incubation with monoclonal anti-Smad2, anti-pSmad2 (Ser465/467), or anti-Smad2/3 antibody (Cell Signaling Technology), the membranes were washed 3 times for 10 min with TBST and then incubated with appropriate horseradish peroxidase-labeled secondary antibodies (1:1000; Cell Signaling Technology) for 1 h at room temperature. Following three 10-min washes with TBST, the protein bands were detected by chemiluminescence (Sigma, St. Louis, MO, USA).

Data are presented as mean ± standard deviation. Statistical analysis was performed using Analysis of Variances (ANOVA) with SPSS 18.0 software (IBM Corporation, Armonk, NY, USA). Levene’s tests for equal variances were applied before one-way ANOVAs. When equal variances were assumed, a one-way ANOVA with Bonferroni post- hoc testing was used for multiple group comparison. If equal variances were not assumed, Tamhane's T2 test was used for multiple comparison tests. All tests were two-tailed. Differences with a P value less than 0.05 were considered statistically significant.

3. Results

To optimize the dose of TGF-β₁ to induce hypertrophic responses, we stimulated NRCMs with TGF-β₁ for 24 h. As shown in Figure 1A–G, 3 μg/L of TGF-β₁ resulted in significantly increased PI fluorescence intensity and [³H]-leucine incorporation in NRCMs compared with all other groups (P < 0.01 vs. other groups). Similar results were observed in the mRNA levels of fetal genes ANF and β-MHC (Figure 1H and 1I). Therefore, 3 μg/L TGF-β₁ that generates the maximum effect in inducing hypertrophic responses in NRCMs was applied in the following experiment.

Because the TGF-β/Smad signaling promotes cardiac hypertrophy ^{22, 23, 24, 25}, we hypothesized that puerarin might protect the heart from TGF-β₁-induced hypertrophy by suppressing the Smad signaling. To test our hypothesis, we explored whether TGF-β₁ can induce the Smad signaling in NRCMs. As shown in Figure 2, TGF-β₁ (3 μg/L) significantly induced upregulation of mRNA and protein expression of Smad2, pSmad2, and total Smad2/3 in NRCMs starting at 30, and 15 min and peaking at 1 h, 2 h, and 30 min, respectively, after treatment, indicating a chronological activation of the Smad signaling in NRCMs exposed to TGF-β₁.

Next, we investigated the effect of puerarin or knockdown of Smad2 on TGF-β₁-induced hypertrophic responses in NRCMs. We treated NRCMs with different doses of puerarin in the presence of TGF-β₁ (3 μg/L) or transfected NRCMs with different amounts of Smad2 siRNA (Figure 3) prior to TGF-β₁ exposure. As shown in Figure 4A and 4B, both puerarin and Smad2 siRNA could significantly reverse TGF-β₁-enhanced [³H]-leucine incorporation in NRCMs (all P< 0.05), but puerarin (1 g/L) exhibited a higher potency than different amounts of Smad2 siRNA (P<0.05). Consistently, puerarin (1 g/L) also significantly countered TGF-β₁-enhanced mRNA expression of ANF and β-MHC, with a greater effect than Smad2 siRNA (Figure 4C; all P< 0.05). Taken together, these results suggest that the Smad2 signaling is required for TGF-β₁-induced protein synthesis and fetal gene expression in NRCMs and that puerarin could effectively reverse these effects of TGF-β₁ in NRCMs.

Next, we sought to explore whether puerain can affect Smad protein expression in the presence of TGF-β₁. We determined the mRNA and protein expression of Smad2 in NRCMs exposed to TGF-β₁ alone or in combination with puerarin. As shown in Figure 5A–C, like Smad2 siRNA, puerarin could significantly abrogate TGF-β₁-enhanced expression of Smad2, pSmad2, and total Smad2/3 in NRCMs (all P< 0.01 vs. TGF-β₁ group), suggesting that puerarin protects NRCMs against TGF-β₁-induced hypertrophic changes possibly via deactivation of the Smad signaling.

4. Discussion

In the present study, we investigated the effect of puerarin on TGF-β₁-induced hypertrophic changes and Smad protein alterations in NRCMs. Our results showed that puerarin could effectively reverse TGF-β₁-induced hypertrophic changes and significantly abrogate TGF-β₁-induced Smad signaling activation in NRCMs, providing Smad proteins as possible targets for puerarin treatment in animal models.

Cardiac hypertrophy is characterized by enlargement of cardiomyocytes, enhancement of protein synthesis, and reactivation of fetal genes ^{1, 2, 3}. Our results showed that TGF-β₁ could significantly promote total RNA and protein synthesis and fetal gene transcription in NRCMs, suggesting a prohypertrophic effect of TGF-β₁ in NRCMs, consistent with previous studies ^{22, 25, 29, 30}. TGF-β₁ is secreted by cardiac fibroblasts, cardiomyocytes, and endothelial cells, mediating intracellular signaling via the Smad family ^{31, 32}. Our results showed that significant upregulation of total Smad2/3 began at 15 min and peaked at 30 min, whereas that of Smad2 and pSmad2 began and peaked at later time points post TGF-β₁ stimulation, indicating a chronological activation of total Smad2/3, Smad2, and pSmad2 in TGF-β₁-stimulated NRCMs. It has been reported that pharmacological inhibition of the Smad2 signaling can attenuate aortic banding-induced cardiomyocyte hypertrophy and restore cardiac function in mice ³³, suggesting that targeting the Smad2 signaling is a promising therapeutic strategy in the treatment of cardiac hypertrophy. Consistently, in the present study, we found that knockdown of Smad2 could effectively reverse TGF-β₁-induced hypertrophic changes in NRCMs, indicating that the Smad signaling is required in TGF-β₁-induced hypertrophic changes. Importantly, puerarin treatment exhibited similar but stronger effects than Smad2siRNA in inhibiting TGF-β₁-stimulated protein synthesis and fetal gene transcription. Based on these findings, we speculate that puerarin might attenuate TGF-β₁-induced hypertrophic responses in NRCMs possibly and partially via suppressing the Smad2 signaling. Furthermore, we found that, like Smad2 siRNA, puerarin significantly abrogated TGF-β₁-enhanced expression of Smad2, pSmad2, and total Smad2/3 in NRCMs, suggesting an involvement of Smad signaling suppression in the antihypertrophic role of puerarin in NRCMs.

Previous studies have suggested other signaling pathways underlying the antihypertrophic effect of puerarin. For example, in spontaneously hypertensive rats, puerarin reduces mRNA expression of TGF-β₁ and Smad3 while increasing that of Smad7 in myocardium ³⁴. In addition, puerarin retards the progression of cardiac hypertrophy by blocking the PI3K/Akt and JNK signaling pathways ¹⁹. Our previous study also suggests that puerarin can protect against cardiomyocyte hypertrophy by restoring autophagy through the AMPK/mTOR signaling ³⁵. These findings collectively suggest that puerarin may ameliorate cardiac hypertrophy via modulating multiple signaling pathways, explaining why puerarin exhibited greater inhibition in TGF-β₁-induced protein synthesis in NRCMs compared with Smad2 knockdown.

In conclusion, our results demonstrated inhibitory effects of puerarin on TGF-β₁-induced hypertrophic responses and Smad signaling activation in NRCMs, providing new insights into the mechanism underlying the antihypertrophic effect of puerarin. Future studies will further investigate the antihypertrophic role of puerarin and the involvement of the Smad signaling in an anrimal model.

Acknowledgments

All authors declare that they have no conflict of interests.

Funding

The study was supported by Guangdong administration of traditional Chinese medicine (20162096).

References