Role of caspase-9 in the effector caspases and genome expressions, and growth of bovine skeletal myoblasts

Authors


Abstract

Caspase-9 has been reported as the key regulator of apoptosis, however, its role in skeletal myoblast development and molecular involvements during cell growth still remains unknown. The current study aimed to present the key role of caspase-9 in the expressions of apoptotic caspases and genome, and cell viability during myoblast growth using RNA interference mediated silencing. Three small interference RNA sequences (siRNAs) targeting caspase-9 gene was designed and ligated into pSilencer plasmid vector to construct shRNA expression constructs. Cells were transfected with the constructs for 48 h. Results indicated that all three siRNAs could silence the caspase-9 mRNA expression significantly. Particularly, the mRNA expression level of caspase-9 in the cells transfected by shRNA1, shRNA2 and shRNA3 constructs were reduced by 37.85%, 68.20% and 58.14%, respectively. Suppression of caspase-9 led to the significant increases in the mRNA and protein expressions of effector caspase-3, whereas the reduction in mRNA and protein expressions of caspase-7. The microarray results showed that the suppression of caspase-9 resulted in significant upregulations of cell proliferation-, adhesion-, growth-, development- and division-regulating genes, whereas the reduction in the expressions of cell death program- and stress response-regulating genes. Furthermore, cell viability was significantly increased following the transfection. These data suggest that caspase-9 could play an important role in the control of cell growth, and knockdown of caspase-9 may have genuine potential in the treatment of skeletal muscle atrophy.

Introduction

Muscle tissue possesses the post-natal growth and intrinsic regenerative capacity (Bischoff 1986). Comprehensive understanding of the myoblasts involvement in postnatal myogenesis, myofiber regeneration and skeletal muscle hypertrophy are noteworthy issues for fundamental agricultural reasons. Aspartate-specific cysteine proteases (caspases) are expressed as pro-enzymes containing three domains, including an NH2 terminal, a large subunit (approximately 20 kDa) and a small subunit (approximately 10 kDa). To date, 14 different caspases have been identified, which play distinct roles in apoptosis and inflammation in a number of different tissues and cell types (Wolf & Green 1999; Nakagawa & Yuan 2000; Philchenkov 2004). Out of these caspases, caspase-3,-7 and-9 are involved apoptosis. These apoptotic caspases can be further subdivided into initiator caspases such as caspase-9 and the effector caspases such as caspase-3 and -7, depending on their position in the cell death pathway (Earnshaw et al. 1999).

Previous studies have demonstrated caspase-9 as a key regulator of apoptosis triggered in a response to stimuli that damage mitochondria (Luo et al. 1998; Morishima et al. 2002; Robertso et al. 2002; Würstle et al. 2012). The caspase-9 involves the release of cytochrome c from mitochondria, which then binds and activates apoptotic protease-activating factor-1 (Apaf-1) and then activates procaspase-9. The active caspase-9 at the apoptosome subsequently activates procaspase-3 and caspase-7 to execute cell death program (Grutter 2000; Würstle et al. 2012). However, the role of caspase-9 in skeletal muscle cell development and molecular involvements during growth still remain unknown. A recent study by Lakhani et al. (2006) found that the apoptotic caspases-lacking mice die shortly after birth, suggesting that the apoptotic caspases may also play some important role in cell growth.

More to the point, in recent years RNA interference (RNAi) mediated by small interfering RNA (siRNA) has been widely used in gene function studies (Shan et al. 2010; Liu et al. 2011). The RNAi method usually uses short hairpin RNA (shRNA) expressed from plasmid vectors, and the shRNA-expressing construct is integrated into the genome, it becomes a sustainable source of siRNA and produces sustained RNAi effects. Thus, the RNAi strategy, which can inhibit caspase-9 specifically, would be a powerful tool to clarify physiological functions.

Furthermore, satellite cells in postnatal skeletal muscle are mononucleated myogenic precursors located beneath the basal lamina of myofibers (Bischoff 1986). These cells are normally quiescent, but can be activated to regulate the postnatal muscle growth and muscle repair (Allen & Rankin 1990; Cornelison & Wold 1997). In recent years, there has been a growing interest in the agricultural community to apply myoblasts for studying the cellular regulation of meat-animal muscle growth. Therefore, understanding the role of particular genes involved in cell growth and apoptosis is noteworthy for maintaining myoblast proliferation during muscle growth as well as control of cell death in muscle atrophy. To the best of our knowledge, no scientific information regarding the role of caspase-9 in cell proliferation and its effect on the expressions of effector caspases as well as the genomic system in bovine skeletal myoblasts during growth is available. Thus, the current work was designed to study the biological role of caspae-9 in the expressions of effector caspases and genomic system, and cell proliferation.

Materials and methods

Cell preparation and culture

The myoblasts were isolated from 24-month-old Korean native cattle (Hanwoo) following the Dodson et al. (1987) method. The entire work involving the use of animals was approved by an Institutional Animal Care and Use Committee. Briefly, semitendinosus muscle (500 g) was excised from the animal immediately after slaughter at a commercial abattoir (located in Jeonju province, South Korea). The connective tissue and most of the fat was trimmed off and discarded. The muscle was cut into small fragments (about 3 mm3). After enzymatic digestion with 1% pronase solution (Sigma, St. Louis, MO, USA) at 37°C for 60 min, single cells were separated from the fragments by repeated centrifugation at 1000 g for 10 min at room temperature. The primary muscle cells were cultured in Dulbecco's Modified Eagle's Medium (DMEM, Gibco) supplemented with 15% fetal bovine serum (FBS), 100 IU/mL penicillin, and 100 μg/mL streptomycin (Sigma) in a humidified incubator at 37°C with 5% CO2. To isolate myoblasts from the primary muscle cells, the cells were applied to a magnetic cell sorting system (AutoMACS, Milteny Biotech, Germany). Particularly, when the cells reached 80% confluence, they were collected and re-suspended in phosphate-buffered saline (1× PBS, Gibco) supplemented with 0.5% bovine serum albumin (BSA) and 2 mmol/L ethylenediaminetetraacetic acid (EDTA). After centrifugation (400 g for 5 min), the cell pellet was re-suspended in PBS (100 μL) containing 10 μg anti-M-cadherin antibodies and then incubated with 20 μL of anti-mouse IgG1 microbeads at 4°C for 30 min. Finally, cell suspension (approximately 107 cells in 2 mL PBS) was loaded into a magnetic cell sorting system to isolate myoblasts. Furthermore, to confirm whether the isolated cells are really myoblasts, the positive cells were also cultured in myogenic differentiation medium (DMEM containing 2% horse serum) for 7 days to check the myotube formation. The isolated cells were cultured in growth medium as mentioned above, sub-cultured when they reached confluence (approximately 80%), and cells from the fourth passage were used for the present study.

Designing of siRNA and construction of plasmid vector

Three siRNA sequences against caspase-9 gene were designed as per the siRNA designing program: http://www.ambion.com/techlib/misc/siRNA_design.html. The siRNAs were selected on the basis of ranking criteria of Reynolds et al. (2004). The siRNAs were converted to shRNAs by using the siRNA target finder program for the pSilencer vector on the web page http://www.ambion.com/techlib/misc/psilencer_converter.html. The top strand oligonucleotides encoding the shRNA contains the following structural features: a nucleotide sequence derived from the caspase-9 (sense strand), a short space of loop sequence, a nucleotide sequence that is the reverse complement of the caspase-9 (antisense strand) and following four continuous thymines as terminate signal, and two restriction enzyme sites (BamH I and Hind III) located at two termination ends. The three target shRNA sequences for constructing pSilencer vector against caspase-9 were named as shRNA1 top: (5′-GATCCGTGAGCGAGGTGATGAAGCTTTCAAGAGAAGCTTCATCACCTCGCTCATTTTTTGGAAA-3′) and shRNA1 bottom: (5′-AGCTTTTCCAAAAAATGAGCGAGGTGATGAAGCTTCTCTTGAAAGCTTCATCACCTCGCTCACG-3′); shRNA2 top: (5′-GATCCGGACGAGAACTACGACCTGTTCAAGAGACAGGTCGTAGTTCTCGTCCTTTTTTGGAAA-3′) and shRNA2 bottom: (5′-AGCTTTTCCAAAAAAGGACGAGAACTACGACCTGTCTCTTGAACAGGTCGTAGTTCTCGTCCG-3′); shRNA3 top: (5′-GATCCGGAGAGCTTCGAGAACTACTTCAAGAGAGTAGTTCTCGAAGCTCTCCTTTTTTGAAA-3′) and shRNA3 bottom: (5′-AGCTTTTCCAAAAAAGGAGAGCTTCGAGAACTACTCTCTTGAAGTAGTTCTCGAAGCTCTCCG-3′). These oligo sequences were annealed and the resulting annealed shRNAs were ligated into pSilencer hygro vector by T4 DNA ligase between the BamH I and Hind III restriction sites according to the manufacturer's instructions. The ligated product (shRNA expression constructs) was transformed into Escherichia coli GC competent cells (Sigma) following the manufacturer's protocol. After amplification, the ligated product was isolated using a GenElute Plasmid Miniprep Kit (Sigma) and then was digested by endonuclease BamH I and Hind III restriction enzymes. The digested products (shRNA and plasmid vectors) were visualized by 1.8% agarose gel electrophoresis and then used for the transfection.

Electroporation of shRNA expression constructs

The cells (2 × 106 cells) at fourth passage suspended in 100 μL of opti-MEM (Gibco) were transferred to a 4-mm electroporation cuvette (BTX, Holliston, MA, USA). Ten micrograms of each shRNA expression constructs or pSilencer hygro vector negative control was added to the cuvette and electroporated with two pulses at 100V, 100Ω and 1500 μF using an ECM630 Electroporator (BTX, San Diego, MA, USA). After electroporation, the transfected cells were collected and cultured in growth medium (DMEM containing 15% FBS, 100 IU/mL penicillin, and 100 μg/mL streptomycin) in a humidified incubator at 37°C with 5% CO2. The day following transfection, the cells were washed once with 1 × PBS and replaced with the fresh growth medium. The cells were cultured for a further 24 h and then were used to investigate the transfection efficiency, expressions of effector caspases and genome, morphology and cell viability.

Confocal scanning laser microscope

For the confocal analysis, after electroporation the transfected cells at density of 2 × 105 cells/well were transferred to a 6-well plate with 2 mL of growth medium/well and incubated for 48 h as described above in the present study. The cells were then stained with the contents of live/dead cytotoxicity kit (Invitrogen) following our previously established method (Amna et al. 2013). Finally the stained cells were observed under the fluorescence microscope.

Quantitative real time polymerase chain reaction

Following 48 h shRNA transfection, the transfected cells were lysed using RNA extraction reagent kit (Trizol, Invitrogen) and total RNA was extracted according to the manufacturer's protocol. The first-strand cDNA was synthesized from 1 μg of the total RNA using the reverse transcriptase with the anchored oligo d(T)12-18 primer (Gene Link). Real-time polymerase chain reaction (PCR) was performed using a cDNA equivalent of 10 ng of total RNA from each sample with primers specific for bovine caspase-3, caspase-7, caspase-9 and a housekeeping gene glyceraldehyde 3-phosphate dehydrogenase (GAPDH). Primers specific for bos taurus bovine caspase-3, caspase-7, caspase-9 and a housekeeping gene GAPDH used in this research were as follows: caspase-3 (forward 5′-GTTCATCCAGGCTCTTTG-3′ and reverse 5′-TCTATTGCTACCTTTCG-3′); caspase-7 (forward 5′-GAATGGGTGTCCGCAACG-3′ and reverse 5′-TTGGCACAAGAGCAGTCGTT-3′); caspase-9 (forward 5′-CGCCACCATCTTCTCCCTG-3′ and reverse 5′-CCAACGTCTCCTTCTCCTCC-3′); and a housekeeping gene GAPDH (forward 5′-CACCCTCAAGATTGTCAGC-3′ and reverse 5′-TAAGTCCCTCCACGATGC-3′). PCR was performed using 20 μL reaction volumes, containing 10 μL SsoFast EvaGreen Supermix (Bio-Rad), 0.5 μL each (10 pmol/μL) of forward and reverse primers, 1 μL of cDNA (100 ng/μL) and 8 μL of nuclease-free water. The PCR thermal cycle reactions consisted of initial annealing at 57°C for 2 min, denaturation at 95°C for 3 min followed by 40 cycles of denaturation and annealing at 95°C/10 s and 60°C/10 s, respectively. The relative expression of transcripts present for the measured genes in the myoblasts was normalized to GAPDH transcripts. Relative ratios were calculated using the method as discribed by Pfaffl (2001).

Preparation of cell lysates and western blot

After 48 h of transfection the transfected cells were harvested and protein content was extracted using CelLytic M kit (Sigma) according to the manufacturer's instructions. The protein concentration was determined using Bio-Rad protein assay kit (Bio-Rad). Whole protein (50 μg) of each sample was separated on 12.5% acrylamide with 4% acrylamide stacking gels. The proteins from gels were transferred to Hybond-P PVDF membrane (GE Healthcare, Amersham, UK) for 60 min at 200 mA. Membranes were blocked in 20 mL of blocking solution (20 mmol/L Tris-HCl, 137 mmol/L NaCl, 5 mmol/L KCl and 0.05% Tween 20) for 60 min and then incubated for 60 min at room temperature with either caspase-3 (1:1000), caspase-7 (1:1000) or actin (1:1000) primary antibodies in blocking solution. Subsequently, the bound primary antibodies were then labeled with alkaline phosphate-conjugated rabbit anti mouse IgG secondary antibody in blocking solution for 60 min at room temperature. The bound protein-antibodies were visualized by incubating the membrane with alkaline phosphate-conjugate substrate (Bio-Rad).

Microarray analysis

The total RNA was extracted from the transfected cells for microarray analysis. The synthesis of target cRNA probes and hybridization were performed using Agilent's LowInput QuickAmp Labeling Kit (Agilent Technology) according to the manufacturer's instructions. The fragmented cRNA was then re-suspended with 2× hybridization buffer and directly pipetted onto assembled bovine (V2) gene expression 4 × 44K microarray (Agilent Technologies). The hybridization images were analyzed by Agilent DNA microarray Scanner (Agilent Technology) and the data quantification was performed using Agilent Feature Extraction software 10.7 (Agilent Technology). All data normalization and selection (fold-changed) were performed using GeneSpringGX 7.3.1 (Agilent Technology). Reliable genes were filtered by flag as following the Agilent manual. The average of normalized ratio was calculated by dividing the average of control normalized signal intensity by the average of test normalized signal intensity. Functional categorization of gene families over-represented was performed using the program from the National Institute of Allergy and Infectious Disease (NIAID) web site (http://david.abcc.ncifcrf.gov/summary.jsp).

Cell viability assay

To assay the impact of caspase-9 suppression on cell viability in cell proliferation, the cell viability was determined by using CCK-8 colorimetric assay (Sigma). Briefly, cells were transfected with shRNA expression constructs or pSilencer hygro vector negative control using electroporation technique as described above in the present study. After transfection, cells at density of 2 × 104 cells/well were transferred to 96-well plates with 100 μL growth medium. Afterward, the cells were incubated in a humidified incubator at 37°C with 5% CO2 as described above in the present study. After incubation of 48 h, 10 μL of CCK-8 solution was added to each well and incubated for 4 h at 37°C in a humidified incubator. The light absorption was measured at 450 nm with a microplate reader (Bio-Rad).

Statistical analysis

The experiments were performed in triplicates. Analysis of variance (anova) test was applied to test the significance of difference between the cells transfected by three different shRNA expression constructs with control cells. All mean values were compared using the Duncan's multiple-range test at the 5% level of significance (SAS Institute).

Results

Confirmation of shRNA constructs for RNAi study and shRNA transfection efficiency

The shRNA consists of a sense strand, a short loop sequence, an antisense strand and following four continuous thymines RNA polymerase III terminator. All of the annealed shRNA template inserts were ligated between the BamH I and Hind III sites of the pSilencer plasmid vector. Sixteen hours after the transformation of the constructs into the E. coli GC competent cells, transformed cells produced distinct colonies on LB-agar plate containing ampicillin (50 μg/mL) as selective media. The expected CANP1-siRNAs and plasmid vector digested with BamH I and Hind III was visualized in 1.8% agarose gel electrophoresis (Fig. 1A).

Figure 1.

Knockdown of caspase-9 in myoblasts analyzed at mRNA level. (A) the expected colony plasmid vector and shRNAs digested with BamH I and Hind III was visualized in 1.8% agarose gel electrophoresis; (B) the mRNA expression of caspase-9 following shRNA transfection was assayed by reverse transcription–polymerase chain reaction RT–PCR. Gene expression was normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) expression and denoted as a gene/GAPDH ratio. *< 0.05; (C): The expression of caspase-9 gene was analyzed by 1.5% agarose gel.

In order to get effective siRNA sequences, three shRNA expression constructs were constructed. Following the transfection, qRT–PCR were used to assess the inhibition level of caspase-9 expression achieved by all used siRNA sequences. Transfection with a plasmid encoding caspase-9 siRNAs resulted in significant downregulation of caspase-9 mRNA level in myoblasts. In particular, the relative mRNA expression level of caspase-9 gene in the cells transfected by shRNA1, shRNA2 and shRNA3 constructs were reduced by 37.85%, 68.20% and 58.14%, respectively, when compared with the control cells (Fig. 1B,C).

Expression of effector caspase-3 and -7

In order to determine the involvement of initiator caspase-9 in expressions of effector caspase-3 and -7 in bovine skeletal myoblast during growth, we have explored the mRNA and protein expressions of these caspases following 48 h tranfection by using qRT–PCR and western blot, respectively. Caspase-3 mRNA expression in the cells transfected by the shRNA1, shRNA2 and shRNA3 constructs increased to 2.48-, 3.15- and 2.66-fold, respectively, when compared with control cells (Fig. 2A). Similarly, the protein expression levels of caspase-3 in the transfected cells were also significantly increased (Fig. 2C,D). In contrast to the caspase-3 expression, the caspase-7 mRNA expression in the cells transfected by the shRNA1, shRNA2 and shRNA3 constructs reduced to 0.29-, 0.25-, and 0.35-fold, respectively. Also, the caspase-7 protein expressions were found to significantly decrease when compared with the control cells (Fig. 3A–D).

Figure 2.

Effect of caspase-9 suppression on the caspase-3 expression in myoblasts. (A) The mRNA expression of caspase-3 was assayed by quantitative real-time polymerase chain reaction (PCR) following 48 h transfection with either shRNA expression constructs or pSilencer hygro vector negative control (control). Values expressed as mean ± standard error of mean (SEM), calculated from three independent experiments. Gene expression was normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) expression and denoted as a gene/GAPDH ratio, *< 0.05; (B) The expression caspase-3 gene was analyzed by 1.5% agarose gel; (C) Western blot graph of caspase-3; (D) The relative densitometric values of caspase-3 western blot. Western blot with anti-actin antibody was used as loading control.

Figure 3.

Effect of caspase-9 suppression on the caspase-7 expression in myoblasts. (A) The mRNA expression of caspase-7 was assayed by quantitative real-time polymerase chain reaction (PCR) following 48 h transfection with either shRNA expression constructs or pSilencer hygro vector negative control (control). Values expressed as mean ± standard error of mean (SEM), calculated from three independent experiments. Gene expression was normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) expression and denoted as a gene/GAPDH ratio, *< 0.05; (B) The expression caspase-3 gene was analyzed by 1.5% agarose gel; (C) Western blot graph of caspase-7; (D) The relative densitometric values of caspase-7 western blot. Western blot with anti-actin antibody was used as loading control.

Even though qRT–PCR with specific primers confirmed that caspase-9 gene was silenced at the mRNA level, and caspase-3 upregulated, whereas caspase-7 downregulated in qRT–PCR analysis, we were interested to double-check the expression of these genes by using the microarray technique. As indicated by the present study results, the shRNA2 silenced caspase-9 the most, therefore we chose the shRNA2-transfected cells for the microarray analysis. Similarly, the results of microarray analysis showed that caspase-9 and caspase-7 mRNA expressions were reduced to 0.41- and 0.53-fold, respectively, whereas caspase-3 mRNA expression was increased to 1.03-fold relative to control cells (Fig. 4). The results of microarray analysis were consistent with the results obtained in the RT–PCR.

Figure 4.

Microarray analysis of caspase-9, -7 and -3 gene expressions in myoblasts which were transfected with the shRNA2 for 48 h. The mRNA expression level of transfected cells was calculated based on the respective control values.

Gene expression profile

In order to understand the involvements of caspase-9 in the expressions of other genes we further studied the expressions of genome in the shRNA2-transfected cells following 48 h tranfection by using microarray technique. Interestingly, we observed that 3901 genes were upregulated, whereas 4312 genes were downregulated following the transfection. On the other hand, we compared the gene expression profile in the shRNA2-transfected cells and the control cells. In the scatter plot (Fig. 5) the position of each gene on the plot is determined by its expression level in both shRNA2-transfected cells (x-axis) and control cells (y-axis). Genes with identical or similar expression are depicted by a black line, whereas the gene clusters around the red line depict the ≥2-fold upregulation. The green solid line demonstrates the ≤2-fold downregulation. The red spots at the top of red line indicate ≥3-fold upregulation, whereas the blue spots at the bottom of the green line illustrate ≤3-fold downregulation, respectively. We identified 435 genes (1.76% of 24 681 genes examined) whose expression level in shRNA2-transfected cells was ≥2-fold higher than those of the control cells, and 407 genes (1.64% of 24 681 genes examined) whole expression level was ≤2-fold lower than those in the control cells.

Figure 5.

Expression profiling of genes in shRNAs-transfected cells and control cells. The gene expression profiles of transfected cells or controls were assessed using microarray technique. The position of each gene on the plot is determined by its expression levels in control (x-axis) and shRNAs-transfected cells (y-axis).

By comparing the shRNA2-transfected cells with the control cells, we identified 44 genes, which were significantly downregulated (<5-fold) after transfection by the shRNA2. These downregulated genes were further grouped according to the biologically functional categories including: inhibition of cellular biosynthetic process (nine genes), regulation of programmed cell death (19 genes), stress response (eight genes), inhibition of cell proliferation (four genes) and negative regulation of multicellular organismal process (four genes) (Table 1). On the other hand, we also found 78 genes which were significantly upregulated (>5-fold) after transfection by the shRNA2. These upregulated genes were also grouped according to the biologically functional categories including; regulation of cell proliferation (17 genes), inhibition of apoptosis (14 genes), cell adhesion (23 genes), regulation of cell division (four genes), regulation of cell development (nine genes) and regulation of cell growth (11 genes) (Table 2).

Table 1. A total of 41 genes are significantly downregulated following the shRNA2 transfection
Functional categoryGene IDGene nameSpecies
Inhibition of cellular biosynthetic process NM_001037463 Domain containing E3 ubiquitin protein ligase 1 Bos taurus
NM_001128497 Amino-terminal enhancer of split Bos taurus
NM_001075155 Death-domain associated protein Bos taurus
NM_181010 Endothelin 1 Bos taurus
NM_181811 Myogenic factor 6 (herculin) Bos taurus
NM_001046257 Nitric oxide synthase trafficker Bos taurus
NM_181024 Peroxisome proliferator-activated receptor gamma Bos taurus
NM_001076907 Retinoblastoma 1 Bos taurus
NM_001099023 Zinc finger protein 639 Bos taurus
Regulation of programmed cell death NM_001015544 Bcl2-like 14 (apoptosis facilitator) Bos taurus
NM_001040472 CD3 g molecule, gamma (CD3-TCR complex) Bos taurus
NM_173899 CD5 molecule Bos taurus
NM_174678 PROP paired-like homeobox 1 Bos taurus
NM_001105340 TNF receptor-associated factor 5 Bos taurus
NM_001075129 Calcium channel, voltage-dependent, P/Q type, alpha 1A subunit Bos taurus
NM_001144104 Caspase 2, apoptosis-related cysteine peptidase Bos taurus
NM_001035419 Caspase 6, apoptosis-related cysteine peptidase Bos taurus
NM_001077111 Caspase recruitment domain family, member 9 Bos taurus
NM_001083449 Cell death-inducing DFFA-like effector a Bos taurus
NM_001099017 Distal-less homeobox 1 Bos taurus
NM_001015602 Serine/threonine kinase 4 Bos taurus
NM_001166609 Heat shock 60 kDa protein 1 (chaperonin) Bos taurus
NM_001077988 Neurotrophin 3 Bos taurus
NM_174435 Protein kinase C, alpha Bos taurus
NM_174176 Secretogranin II (chromogranin C) Bos taurus
NM_001012673 Signal transducer and activator of transcription 5A Bos taurus
NM_001038681 Xeroderma pigmentosum, complementation group A Bos taurus
NM_174435 Protein kinase C, alpha Bos taurus
Stress response NM_001075302 Heat shock 105 kDa/110 kDa protein 1 Bos taurus
NM_001167895 Heat shock 70 kDa protein 1-like Bos taurus
NM_001079637 Heat shock protein 90 kDa alpha (cytosolic), class B member 1 Bos taurus
NM_001038530 General transcription factor IIH, polypeptide 2, 44 kDa Bos taurus
NM_001102258 Polymerase (DNA directed) kappa Bos taurus
NM_001079637 Heat shock protein 90 kDa alpha (cytosolic), class A member 1 Bos taurus
NM_174550 Heat shock 70 kDa protein 1A Bos taurus
NM_174345 Heat shock 70 kDa protein 8 Bos taurus
Inhibition of cell proliferation NM_001076804 GATA binding protein 3 Bos taurus
NM_001001601 Cadherin 5, type 2 (vascular endothelium) Bos taurus
NM_001077903 Cyclin-dependent kinase inhibitor 1C (p57, Kip2) Bos taurus
NM_174067 Ghrelin/obestatin prepropeptide Bos taurus
Negative regulation of multicellular organismal process NM_173877 Coagulation factor II (thrombin) Bos taurus
NM_174067 Ghrelin/obestatin prepropeptide Bos taurus
NM_181037 Nitric oxide synthase 3 (endothelial cell) Bos taurus
NM_174474 Troponin T type 1 (skeletal, slow) Bos taurus
Table 2. A total of 80 genes were significantly upregulated following the shRNA2 transfection
Functional categoryGene IDGene nameSpecies
Positive regulation of cell proliferation NM_001038072 BMI1 polycomb ring finger oncogene Bos taurus
NM_181004 CD28 molecule Bos taurus
NM_001099000 GLI family zinc finger 1 Bos taurus
NM_001101097 H2.0-like homeobox Bos taurus
NM_001128499 TGFB-induced factor homeobox 1 Bos taurus
NM_174002 Calcium-sensing receptor Bos taurus
NM_173909 Erythropoietin Bos taurus
NM_178325 Growth hormone releasing hormone Bos taurus
NM_001077828 Insulin-like growth factor 1 (somatomedin C) Bos taurus
NM_180997 Interleukin 2 Bos taurus
NM_001100324 Interleukin 34 Bos taurus
NM_173923 Interleukin 6 (interferon, beta 2) Bos taurus
NM_173924 Interleukin 7 Bos taurus
NM_001046143 CDKN2A interacting protein Bos taurus
NM_001035379 RAS guanyl releasing protein 4 Bos taurus
NM_001103097 Coagulation factor II (thrombin) receptor Bos taurus
NM_001007820 Cocaine and amphetamine responsive transcript Bos taurus
Inhibition of apoptosis NM_173894 BCL2-associated X protein Bos taurus
NM_001046043 Angiopoietin-like 4 Bos taurus
NM_001035386 Catalase Bos taurus
NM_174006 Chemokine (C-C motif) ligand 2 Bos taurus
NM_174027 Colony stimulating factor 2 (granulocyte-macrophage) Bos taurus
NM_174289 Crystallin, alpha A Bos taurus
NM_174290 Crystallin, alpha B Bos taurus
NM_001098958 Cyclin-dependent kinase inhibitor 1A (p21, Cip1) Bos taurus
NM_001037464 Eukaryotic translation elongation factor 1 alpha 2 Bos taurus
NM_001083674 Glutamate-cysteine ligase, catalytic subunit Bos taurus
NM_174076 Glutathione peroxidase 1 Bos taurus
NM_180997 Interleukin 2 Bos taurus
NM_199445 Prokineticin 2 Bos taurus
NM_001012673 Signal transducer and activator of transcription 5A Bos taurus
Cell adhesion NM_001075310 BCL2-like 11 (apoptosis facilitator) Bos taurus
NM_174010 CD36 molecule (thrombospondin receptor) Bos taurus
NM_176661 CD97 molecule Bos taurus
NM_001103335 Protein NGX6 Bos taurus
NM_001034635 Actinin, alpha 2 Bos taurus
NM_181002 Amine oxidase, copper containing 3 (vascular adhesion protein 1) Bos taurus
NM_001080345 Angiopoietin-like 3 Bos taurus
NM_001035373 Annexin A9 Bos taurus
NM_174741 Basal cell adhesion molecule (Lutheran blood group) Bos taurus
NM_001015550 Cadherin 16, KSP-cadherin Bos taurus
NM_001099181 Cadherin 19, type 2 Bos taurus
NM_001166492 Cadherin 2, type 1, N-cadherin (neuronal) Bos taurus
NM_001015642 Carboxypeptidase X (M14 family), member 1 Bos taurus
NM_001166517 Cartilage oligomeric matrix protein Bos taurus
NM_001102110 Catenin (cadherin-associated protein), alpha 2 Bos taurus
NM_001145508 Coagulation factor VIII, procoagulant component Bos taurus
NM_001076831 Collagen, type III, alpha 1 Bos taurus
NM_001083388 Collagen, type XVIII, alpha 1 Bos taurus
XM_599315 Collagen, type XXII, alpha 1 Bos taurus
NM_001102496 Integrin, alpha D Bos taurus
NM_174679 Integrin, beta 5 Bos taurus
NM_174698 Integrin, beta 6 Bos taurus
NM_174348 Intercellular adhesion molecule 1 Bos taurus
Regulation of cell division NM_001075956 Bardet-Biedl syndrome 4 Bos taurus
NM_001035386 Catalase Bos taurus
NM_174092 Interleukin 1, alpha Bos taurus
NM_173950 Placental growth factor Bos Taurus
Regulation of cell development NM_173894 BCL2-associated X protein Bos taurus
NM_001099130 ISL LIM homeobox 1 Bos taurus
NM_001046443 NK2 transcription factor related, locus 5 (Drosophila) Bos taurus
NM_001128499 TGFB-induced factor homeobox 1 Bos taurus
NM_001098099 Atonal homolog 1 (Drosophila) Bos taurus
NM_001102026 Meteorin, glial cell differentiation regulator Bos taurus
NM_174121 Neurofilament, light polypeptide Bos taurus
NM_001098062 Ring finger protein (C3H2C3 type) 6 Bos taurus
NM_001113233 Tubulin tyrosine ligase Bos taurus
Regulation of cell growth NM_001046143 CDKN2A interacting protein Bos taurus
NM_001046095 Chemokine (C-X-C motif) ligand 16 Bos taurus
NM_001098958 Cyclin-dependent kinase inhibitor 1A (p21, Cip1) Bos taurus
NM_001076012 Discoidin domain receptor tyrosine kinase 1 Bos taurus
NM_174681 Growth differentiation factor 9 Bos taurus
NM_174555 Insulin-like growth factor binding protein 2, 36 kDa Bos taurus
NM_001105322 Keratin 17 Bos taurus
NM_001035025 Myosin, light chain 2, regulatory, cardiac, slow Bos taurus
NM_001046501 Proline/serine-rich coiled-coil 1 Bos taurus
NM_001098062 Ring finger protein (C3H2C3 type) 6 Bos taurus
NM_001113233 Tubulin tyrosine ligase Bos taurus

Cell viability

The CCK-8 assay was used to assay the effect of caspase-9 knockdown on the viable cells in cell proliferation. Figure 6A shows CCK-8 data of three different shRNA expressions constructs at 48 h post-transfection when the trasfected cells reached 100% confluent. The results showed a significant increase in the cell viability following the shRNA transfections compared with control cells. Particularly, the cell viability in the cells transfected by shRNA1, shRNA2 and shRNA3 expression constructs were increased by 16.22%, 19.88% and 13.12%, respectively, when compared with the control cells. Further, the cell morphology after transfections was also studied using confocal scanning laser microscopy. The CSLM provides information about cell behavior in a wet environment. Figure 6D,E shows the representative CSLM images of myoblast following 48 h transfection. One can observe that no dead cells were observed in the shRNA2-transfected cells as well as the control cells.

Figure 6.

Effect of caspase-9 suppression on myoblast growth. (A) The viability of cells following transfection as indicated was assayed by the CCK-8 assay. Data represent the mean ± standard error of mean (SEM) of triplicates. Representative images of transfected cells that were taken by electronic microscope at 40 ×  magnification, (B) represents the control cells and (C) represents the shRNA2-transfected cells at 48 h; (D) represents the confocal scanning laser microscope (CSLM) image of control cells; (E) represents the CSLM image of shRNA2- transfected cells at 48 h. The cells were stained by DiOC18 (3)/PI. Dead cells are labeled by PI and have red nuclei. Live cells are labeled by DiOC18 (3) and have green nuclei.

Discussion

In this study, the primary myoblasts are isolated directly from semitendinosus muscle of 24-month-old Korean native cattle. The in vitro properties exhibited by primary cultures of myoblasts more closely reflect their in vivo properties than those exhibited by transformed cell lines (Allen 1987). Primary cell cultures, however, do possess non-myogenic cells (Bischoff 1986; Dodson et al. 1996). To decrease the presence of non-myogenic cells in primary cultures, in the present study the cell suspensions were loaded onto a magnetic cell sorting system (AutoMACS) to isolate the myoblasts. Under aseptic and our standardized culture conditions as aforementioned, cells proliferated until day 7, where they begin to fuse into myotubes.

In this study, three siRNA sequences that silence caspase-9 mRNA of cattle skeletal myoblasts with the highest scores were selected on the basis of ranking criteria of Reynolds et al.(2004). The silencing efficiency of caspase-9 gene in cells was recorded with varying efficiencies among three different shRNA expression constructs. Earlier workers (Golding et al. 2006) have also reported the efficient silencing of specific genes by the RNAi approach using plasmid vectors to transfect into the cell lines.

In the present study to understand how the initiator caspase-9 involved in the expressions of effector caspase-3 and -7 in myoblasts during cell growth we have explored the mRNA and protein expressions of these caspases following the caspase-9 suppression. Interestingly, it was observed that suppression of caspase-9 significantly increased caspase-3 expression, meaning that activation of caspase-9 may inactivate caspase-3 expression. In contrast to our results many authors have reported that the activation of caspase-9 led to the activation of the effector caspases including caspase-3 (Maria 2012; Würstle et al. 2012). However, this is probably due to in the previous studies, the experiments were conducted and performed under the apoptosis-induced conditions that led to the activation of the apoptotic caspase system, whereas in the present study, the cells were cultured under the normal growth condition. McNeish et al. (2003) reported that inhibition of caspase-9 subsequently reduced the caspase-3 activation in ovarian cells and it is possible that differences in caspases level between different cell types could be responsible for different effector caspases executions. Additionally, a study of Gee et al. (2002) indicated that the caspase-3 is not necessarily essential for the execution of apoptosis in MCF-7 breast carcinoma cells under apoptosis-induced conditions. On other hand, numerous reports have found the existence of caspase-independent pathways leading to cell death (Bello et al. 2004; Chipuk & Green 2005; Eguchi et al. 2009). Thus, the result indicating the suppression of caspase-9 increased caspase-3 expression in our work suggests that the caspase-3/caspase-9 interaction is a complex process and it may play some important role in the proliferation of cattle skeletal myoblasts.

A number of studies have found that the caspase-7 play an important role in the execution of apoptosis and the activation of initiator caspase-9 under the apoptotic conditions that led to the increase in the expression of effector caspase-7 (Lamkanfi & Kanneganti 2010). In the present study, the suppression of caspase-9 led to the significant reduction in caspase-7 expression. This means that the caspase-9 activation may activate downstream of caspase-7, and thus the caspase-9/caspase-7 interaction may be responsible for the execution of apoptosis in the cattle skeletal myoblasts. On the other hand, to date there are at least two major cross-talking pathways in cells involving apoptosis: (i) the mitochondrion-initiated pathway (intrinsic pathway), and (ii) the cell surface death receptors pathway (extrinsic pathway) (Slee et al. 2000; Strasser et al. 2000). Previous study showed that treatment of ovarian carcinoma cells with caspase-9 inhibitor reduced caspase-3 activation (McNeish et al. 2003). This evidence and our results both supported the idea of a caspase-dependent pathway (Kischkel et al. 1995; Shi 2001).

In the present study, to elucidate the molecular involvement of caspase-9 during cell growth, the gene expression profile was analyzed using a microarray technique. It is interesting to note that the suppression of caspase-9 significantly increased the expressions of proliferation-, adhesion- and division-regulating genes, whereas it downregulated the expressions of apoptosis- and stress response-regulating genes. These results suggest that caspase-9 not only plays an important role in regulation of apoptosis but also in the control of cell proliferation, adhesion and division. Previous studies have found that the apoptotic caspases also play a positive role in the regulation of lymphocytes, differentiation of human and murine erythroblasts, and other cellular events (Christian & Klaus 2003). Furthermore, we also examined the effect of caspase-9 suppression on cell viability in cell proliferation and morphology; the results show that the cell viability was increased following the shRNA transfection. The reason why the suppression of caspase-9 led to the increase in the cell viability remains to be answered; however, it is probably that the suppression of caspase-9 may reduce the impact from environmental factors, such as stress due to the transfection, and thus activates the cell division and proliferation. Overall, the caspases in general and caspase-9 in particular play important roles in initiating apoptosis execution in cells that need to be eliminated during early developmental stages, and to remove the damaged cells to suppress proliferative diseases during the entire lifetime. However, the apoptosis under the regulation of these caspases can participate in muscle atrophy or muscle wasting by leading to loss of myofibers or loss of myofiber segments (hypotrophy). Therefore, understanding the pathways that lead to apoptosis and identifying strategies to regulate this pathway may have important implications in the control of cell proliferation as well as prevention of skeletal muscles from atrophy under the atrophic conditions.

In summary, the present study demonstrated that caspase-9 plays an important role in the expressions of caspase-3, caspase-7 and other genes in the genomic system. Additionally, targeted suppression of caspase-9 led to the reduction of caspase-7 activation, suggesting that the caspase-9/caspase-7 interaction could undertake the apoptosis execution in cattle skeletal myoblasts through the intrinsic pathway. Furthermore, cell viability assay indicated that suppression of caspase-9 significantly increased cell viability. Also, microarray results indicated that suppression of caspase-9 significantly increased the expressions of cell proliferation-, adhesion- and division-regulating genes, whereas it reduced the expression of cell death program- and stress response-regulating genes. From the present study results, it is possible to suggest that caspase-9 could play an important role in myoblast proliferation during muscle growth.

Acknowledgments

This work was supported by the research funds of Chonbuk National University, South Korea in 2012; the research grants for the FTA issue project (No. PJ907055), RDA and the research grants for the FTA issue project (No. PJ008525), RDA, Republic of Korea.

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