<i>Chlorella vulgaris</i> Modulates Genes and Muscle-Specific microRNAs Expression to Promote Myoblast Differentiation in Culture

Zainul Azlan, Nurhazirah; Mohd Yusof, Yasmin Anum; Alias, Ekram; Makpol, Suzana

doi:https://doi.org/10.1155/2019/8394648

Evidence-Based Complementary and Alternative Medicine

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Research Article | Open Access

Volume 2019 | Article ID 8394648 | https://doi.org/10.1155/2019/8394648

Chlorella vulgaris Modulates Genes and Muscle-Specific microRNAs Expression to Promote Myoblast Differentiation in Culture

Nurhazirah Zainul Azlan,^1,2Yasmin Anum Mohd Yusof,¹Ekram Alias,¹and Suzana Makpol¹

Academic Editor: Carmen Mannucci

Received20 Apr 2019

Revised12 Jun 2019

Accepted03 Jul 2019

Published21 Jul 2019

Abstract

Background. Loss of skeletal muscle mass, strength, and function due to gradual decline in the regeneration of skeletal muscle fibers was observed with advancing age. This condition is known as sarcopenia. Myogenic regulatory factors (MRFs) are essential in muscle regeneration as its activation leads to the differentiation of myoblasts to myofibers. Chlorella vulgaris is a coccoid green eukaryotic microalga that contains highly nutritious substances and has been reported for its pharmaceutical effects. The aim of this study was to determine the effect of C. vulgaris on the regulation of MRFs and myomiRs expression in young and senescent myoblasts during differentiation in vitro. Methods. Human skeletal muscle myoblast (HSMM) cells were cultured and serial passaging was carried out to obtain young and senescent cells. The cells were then treated with C. vulgaris followed by differentiation induction. The expression of Pax7, MyoD1, Myf5, MEF2C, IGF1R, MYOG, TNNT1, PTEN, and MYH2 genes and miR-133b, miR-206, and miR-486 was determined in untreated and C. vulgaris-treated myoblasts on Days 0, 1, 3, 5, and 7 of differentiation. Results. The expression of Pax7, MyoD1, Myf5, MEF2C, IGF1R, MYOG, TNNT1, and PTEN in control senescent myoblasts was significantly decreased on Day 0 of differentiation (p<0.05). Treatment with C. vulgaris upregulated Pax7, Myf5, MEF2C, IGF1R, MYOG, and PTEN in senescent myoblasts (p<0.05) and upregulated Pax7 and MYOG in young myoblasts (p<0.05). The expression of MyoD1 and Myf5 in young myoblasts however was significantly decreased on Day 0 of differentiation (p<0.05). During differentiation, the expression of these genes was increased with C. vulgaris treatment. Further analysis on myomiRs expression showed that miR-133b, miR-206, and miR-486 were significantly downregulated in senescent myoblasts on Day 0 of differentiation which was upregulated by C. vulgaris treatment (p<0.05). During differentiation, the expression of miR-133b and miR-206 was significantly increased with C. vulgaris treatment in both young and senescent myoblasts (p<0.05). However, no significant change was observed on the expression of miR-486 with C. vulgaris treatment. Conclusions. C. vulgaris demonstrated the modulatory effects on the expression of MRFs and myomiRs during proliferation and differentiation of myoblasts in culture. These findings may indicate the beneficial effect of C. vulgaris in muscle regeneration during ageing thus may prevent sarcopenia in the elderly.

1. Introduction

Muscle weakness and atrophy occur in ageing due to multifactorial degenerative processes. It is impacted by cellular ageing biology or primary ageing besides environmental and behavioral factors [1]. The term “sarcopenia” is derived from Greek words “sarx” and “penia”, which mean flesh poverty. It was first introduced by Rosenberg in 1988, in referring to the lean body mass loss in ageing [2, 3]. The diagnosis of sarcopenia can be based on the presence of both low muscle mass and low gait speed [3]. Sarcopenia is also defined as the age-related decline of muscle mass, function, and/or strength, with high prevalence in ageing [4]. It has been reported that muscle strength loss at 2.5% to 3.0% per year in women and 3.0% to 4.0% per year in men, while muscle mass loss at 0.64% to 0.70% per year in women and 0.80% to 0.98% per year in men, in people aged 75 years [5].

Myogenesis is the process of myoblast cells generation which occurs during skeletal muscle tissue synthesis [6]. The satellite cell is important as it is the source of myogenic cells required for myofiber growth and repair throughout life. Proliferation of satellite cell gives rise to satellite cell-derived myoblasts that can differentiate and form multinucleated myotubes [7]. The myogenic regulatory factors (MRFs) are the member of the basic helix-loop-helix family of transcription factors that control the differentiation of skeletal muscle cells during regeneration of muscle [8]. The MRFs consist of four muscle-specific proteins that include Myf5, MyoD, Myogenin, and Myogenic Regulatory Factor 4 (MRF4). These proteins act at multiple points in the muscle lineage to cooperatively establish the skeletal muscle phenotype. It regulates precursor cell proliferation and cell cycle arrest and activates sarcomeric and muscle-specific genes to facilitate differentiation and sarcomere assembly [9].

The satellite cells remain in quiescent state under normal condition. However, when muscle is damaged, the satellite cells are activated and reenter cell cycle phases to produce muscle progenitor cells that are able to regenerate new muscle fibers. In ageing, the satellite cells as well as the systemic and niche environment undergo changes that affect the regenerative functions of the muscle. Aged satellite cells lose their reversible quiescent state due to the upregulation of gene encoding for , a regulator protein of cellular senescence. Furthermore, the disruption of FGF2-Spryl1 signaling and delocalization of β1-integrin in old satellite cells leads to the disturbance of quiescence state and induction of which provokes a switch of senescent-like state in becoming presenescent cells. This process will consequently impair the regeneration of skeletal muscle which includes activation, proliferation, and self-renewal of myoblasts [10–12].

MicroRNAs (miRNAs) are evolutionary conserved, small RNAs with about 17 to 22 nucleotides. It plays vital roles in gene regulatory networks by binding and repressing the activity of specific target mRNAs. MicroRNA has shown its potential as biomarkers in diseases such as cancer [13, 14]. miRNAs are also vital in the posttranscriptional control of gene expression [15, 16]. Several types of miRNAs are highly expressed in different types of tissue and cell, with most of them being tissue-specific. A previous study reported that mature miRNA expression is 20-fold higher than the mean expression of miRNA in other tissues indicating its specificity. Moreover, the expression of miRNA in tissue-enriched with mature miRNA is higher than in other tissues but less than 20-fold [17].

The miRNAs are divided into two categories: miRNAs that are expressed specifically in muscle and not in other tissues known as myomiRs, and (2) miRNA that are expressed in nonmuscle tissue or broadly expressed in other cell types. The myomiRs are important in controlling skeletal muscle development and function as it may affect several biological pathways involved in myoblast proliferation, differentiation, and muscle regeneration [14, 18]. Subsets of myomiRs can be categorized as striated muscle-specific (miR-1, miR-133a, miR-133b, miR-206, miR-208a, miR-208b, and mir-499) or muscle-enriched (miR-486). Some of the myomiRs are expressed in skeletal muscle as well as in cardiac muscle. However, miR-133b and miR-206 are skeletal muscle-specific, and miR-208a is cardiac muscle-specific [16].

World Health Organization (2015) reported that there will be an increase in the number of older population with age between 70 to 80 years in 2050 [19]. And among the major problems in the elderly, sarcopenia is of importance. Taking a natural remedy such as microalgae may be beneficial as one of the approaches in the prevention and management of this condition. Microalgae are believed to be first consumed by human 2000 years ago when the Chinese used blue-green algae, Nostoc to survive during the scarcity of food [20]. Another alga, Chlorella, has been used in variety of biotechnology applications such as biodiesel production and biosorption of heavy metals [21–23]. Chlorella, a genus of unicellular green algae, is the most popular photosynthetic microalgae being studied and investigated currently.

A Dutch researcher, Martinus Willem Beijerinck, discovered C. vulgaris 129 years ago and described it as coccoid green algal “balls” with a well-defined nucleus [23, 24]. The natural components found in C. vulgaris may be responsible for its pharmacological actions. Several studies reported C. vulgaris pharmacological effects, and this includes its antidiabetic [25, 26] and anticancer actions [27–29]. It also possesses highly nutritive contents such as carbohydrates, proteins, nucleic acids, chlorophylls, vitamins, and minerals [30] besides having β-carotene, lutein, chlorophyll-a, chlorophyll-b, ascorbic acid, tocopherol, riboflavin, and retinol [31, 32]. In this study, we aimed to determine the effects of Chlorella vulgaris on the expression of genes and myomiRs involved in muscle differentiation of young and senescent myoblast cells in an attempt to elucidate the mechanism involved during myoblasts differentiation. The finding of this study may provide current information regarding the properties of C. vulgaris that can be used for the prevention of sarcopenia.

2. Materials and Methods

2.1. Cell Culture

Human Skeletal Muscle Myoblasts (HSMM) from 20 years old female Caucasian donor were purchased from Lonza (Walkersville, MD, USA). The skeletal muscle myoblasts were cultured in Skeletal Muscle Basal Medium (SkBM) with fetal bovine serum (FBS), L-glutamine, human epidermal growth factor (hEGF), dexamethasone, and gentamicin/amphotericin-B as supplements to the media (Lonza, Walkersville, MD, USA). Cells were cultivated at 37°C with 5% CO₂ atmosphere. The skeletal muscle myoblast cell then underwent serial passaging until it reached cellular senescence. The population doubling (PD) of the cell was calculated for each passage according to the formula ln (N/n)/ln 2 as N is the number of cells at harvest stage and n is the number of cells at seeding stage [33]. The starting PD for this research was PD 8. The skeletal muscle myoblast cells reached cellular senescence when the cells were unable to proliferate in culture, even with consecutive replenishment. Myoblast cells were considered young at PD 14 and senescent at PD 21.

2.2. Preparation of Chlorella vulgaris for Cell Treatment

Stock of C. vulgaris Beijerinck (strain 072) was obtained from the University of Malaya Algae Culture Collection (UMACC, Malaysia). The stock was grown in Bold’s Basal Media (BBM) with a 12 h dark and 12 h light cycle. The algae were then harvested by centrifugation at 1000 rpm and dried by using freeze dryer. Later, the algae were mixed in distilled water at a concentration of 10% (w/v) and boiled at 100°C for 20 min by reflux method. The alga was centrifuged and lyophilized using a freeze dryer to obtain C. vulgaris in powder form [29, 30, 34]. Myoblast cells which were cultured in T25 culture flask were treated with C. vulgaris at concentrations of 10 and 100 μg/ml [34]. After plating the myoblasts for 24 h and the cells reached 70 to 80% confluency, the culture medium was replaced with C. vulgaris containing medium and left for 24 h in a CO₂ incubator at 37°C. The culture media were then replaced with differentiation medium and parameters were measured on Days 0, 1, 3, 5, and 7 of differentiation induction.

2.3. Immunofluorescence Staining of Myoblasts

Immunofluorescence staining on myoblasts in culture was performed using an antibody specific for desmin, at a dilution of 1:50 (D33; DAKO, Glostrup, Denmark). Skeletal myoblast cells were washed with phosphate-buffered saline (PBS) and fixed in 100% cold ethanol, followed by incubation in 1% fetal bovine serum (FBS). The skeletal myoblast cells were washed again with PBS and incubated consequently with anti-desmin monoclonal antibody (D33, DAKO, Glostrup, Denmark) and Alexa Flour 488 goat anti-mouse in dark environment (Life Technologies, Carlsbad, CA, USA) with PBS washing in between the two antibodies incubation. Cell nuclei were visualized by Hoechst 33342 (Life Technologies, Carlsbad, CA, USA). The desmin staining was viewed under EVOS FL Digital Inverted Fluorescence Microscope (Life Technologies, Carlsbad, CA, USA). The morphological changes of the myoblast cell were observed throughout the differentiation days.

2.4. Induction of Myogenic Differentiation

For induction of muscle cell differentiation, the proliferation medium SkBM was replaced with a differentiation medium; DMEM: F12 (Lonza, Walkersville, MD, USA) with supplementation of 2% horse serum (ATCC, Baltimore, USA). The differentiation medium was changed every two days until the desired day of differentiation for RNA extraction.

2.5. RNA Extraction

The extraction of RNA was carried out on Days 0, 1, 3, 5, and 7 of differentiation. Total RNA was isolated using TRI® reagent (Molecular Research Center Inc, Ohio, USA) on Days 0, 1, 3, 5, and 7 of differentiation. A total of 2 ml TRI reagent was added to 25 cm² culture flask of seeding cells and left for 2 min prior to aliquot into microcentrifuge tube. Chloroform was added at 200 μl and vortex before being left at room temperature for 15 min, followed by vortex at 12000 g and 4°C, for 15 min. The resultant clear layer of RNA was transferred into another new tube prior adding 500 μl isopropanol and 5 μl poly acryl carrier and left at room temperature for 5 min before centrifuged at 12000 g and 4°C for 8 min. The supernatant was removed and the RNA pellet was cleaned with 1 ml of 75% ethanol, followed by centrifuged at 7500 g and 4°C, for 5 min. Supernatant was further removed and RNA pellet was left to dry for about 20 min by directing the open tube to the laminar flow chamber. The dried pellet was later dissolved in 20 μl RNase-free water (Life Technologies, New York, USA) and vortexed to be further kept at −80°C.

2.6. RNA Purity and Concentration

The purity of the extracted RNA was determined by Nanodrop 2000c Spectrophotometer (Thermo Scientific, USA). The extracted RNA with A260/280 value in a range of 1.8 to 2.0 was considered pure and used in gene and myomiRs expression analysis.

2.7. Primer Design

The primer of desired genes was designed from the National Center for Biotechnology Information (NCBI) gene bank. All of the primers used in gene expression analysis were synthesized by Integrated DNA Technologies, Inc. (Illinois, USA) except the primers for Pax7, which was synthesized from Bio Basic Inc. (Markham, ON, Canada) (Table 1). The target sequences for myomiRs expression analysis were synthesized by Life Technologies Corporation (Texas, USA) (Table 2).

2.8. Gene Expression Analysis

Gene expression analysis was carried out by using KAPA SYBR® FAST Bio-Rad iCycler® One-Step RT-qPCR Kit (Kapa Biosystems Pty. Ltd., Boston, Massachusetts, USA) according to manufacturer’s instructions. The PCR mixture was prepared by mixing 0.3 μl RT mix, 9 μl master mix, 12.7 μl RNase-free water, 1.0 μl of 100 μM forward primer, and 1.0 μl of 100 μM reverse primer of desired gene. A volume of 24 μl PCR mixture was added to the desired well of 96-well PCR plate and 1 μl of total RNA sample was also added. The 96-well plate was placed in CFX96 Touch™ Real-Time PCR Detection System (Biorad, California, USA). The protocol of one-step RT-qPCR was cDNA synthesis at 42° for 5 min, inactivation of reverse-transcriptase (RT) at 95°C for 4 min, followed by 40 cycles of 95°C for 3 sec and 53°C for 20 sec. The melting curve and data analysis were then generated at 95°C for a minute, 95°C for 30 sec, 55°C for a min, and 81 cycles of 55°C to 95°C for 10 sec. Primer specificity was then determined from the melting curve produced. All values of the threshold cycle (Ct value) obtained were normalized with the reference gene GAPDH. Relative expression value (REV) was calculated based on the relative quantitative method [35] by (1):

2.9. MyomiRs Expression Analysis

The determination of miRNA expression was carried out by using TaqMan® Advanced miRNA Assays (Life Technologies Corporation, Texas, USA) based on the manufacturer’s protocol. The poly (A) tailing reaction was prepared firstly by mixing 2 μl of total RNA sample and poly (A) reaction mix in reaction tubes, consisting of 0.5 μl 10X poly (A) buffer, 0.5 μl ATP, 0.3 μl poly (A) enzyme, and 1.7 μl RNase-free water, which was then incubated in the thermal cycler for polyadenylation at 37°C for 45 min and stop reaction at 65°C for 10 min. Then, adaptor ligation reaction was performed by adding the ligation reaction mix which contains 3 μl 5X DNA ligase buffer, 4.5 μl 50% PEG 8000, 0.6 μl 25X ligation adaptor, 1.5 μl RNA ligase, and 0.4 μl RNase-free water, into the reaction tubes containing poly (A) tailing reaction product and incubated for ligation at 16°C for 60 min. This is followed by RT reaction by adding the RT reaction mix, consisting of 6 μl 5X RT buffer, 1.2 μl dNTP mix, 1.5 μl 20X universal RT primer, 3 μl 10X RT enzyme mix, and 3.3 μl RNase-free water, into the reaction tubes which contain the adaptor ligation reaction product and incubated for RT reaction at 42°C for 15 min and stop reaction at 85°C for 5 min. The miR-Amp reaction was then continued by mixing miR-Amp reaction mix containing 25 μl 2X miR-Amp master mix, 2.5 μl 20X miR-Amp primer mix, and 17.5 μl RNase-free water and RT reaction product and incubated for enzyme activation at 95°C for 5 min, 14 cycles of denaturation at 95°C for 3 sec, and annealing/extension at 60°C for 30 sec and stop reaction at 99°C for 10 min. The miR-Amp reaction product was further subjected to RT-PCR reaction. A total of 5 μl diluted cDNA template was mixed with 10 μl Taqman® Fast Advanced Master Mix, 1 μl Taqman® Advanced miRNA Assay, and 4 μl RNase-free water and run for enzyme activation at 95°C for 20 sec and 40 cycles of denaturation at 95°C for 3 sec and annealing/extension at 60°C for 30 sec. Data of the threshold cycle (Ct value) acquired was normalized with the reference endogenous control hsa-miR-191-5p. The relative expression value (REV) was then calculated based on the relative quantitative method .

2.10. Statistical Analysis

Data obtained were expressed as mean ± SD and statistical analysis was carried out using SPSS software version 23. Data were analysed using one-way ANOVA followed by Tukey’s post hoc test for multiple comparison. Value of p<0.05 was considered statistically significant.

3. Results

3.1. Immunofluorescence Staining of Myoblasts

Myoblast cells at young stage were spindle-shaped with less branches and underwent changes in morphology at senescent stage on Day 0 of differentiation. Senescent myoblast cells were larger and flatter with the presence of prominent intermediate filaments and single nuclei for each myoblast (Figure 1). Throughout the differentiation days, C. vulgaris-treated young and senescent myoblast cells were observed to differentiate, as notably indicated by the fusion of nuclei per myotube on Day 3 of differentiation and more formation of myotubes on Day 7 compared to Day 0 of differentiation. However, the differentiation of senescent myoblasts was observed to be less efficient as compared to young myoblasts for both the untreated and C. vulgaris-treated cells.

3.2. Gene Expression at Day 0 of Differentiation

On Day 0 of differentiation, Pax7, MyoD1, Myf5, MEF2C, IGF1R, MYOG, TNNT1, and PTEN were significantly downregulated in control senescent myoblasts as compared to control young myoblasts (p<0.05) (Figures 2(a)–2(i)). However, Pax7 was significantly upregulated on Day 0 of differentiation with 100 μg/ml C. vulgaris treatment in young myoblasts (p<0.05) (Figure 2(a)). A similar increase was observed in the expression of Pax7 (Figure 2(a)) and Myf5 (Figure 2(c)) in senescent myoblasts treated with 10 μg/ml and 100 μg/ml C. vulgaris (p<0.05). Decreased MyoD1 (Figure 2(b)) and Myf5 (Figure 2(c)) were observed in young myoblasts treated with both 10 μg/ml and 100 μg/ml C. vulgaris (p<0.05). Treatment with 100 μg/ml C. vulgaris in senescent myoblasts significantly increased the expression of MEF2C (Figure 2(d)) and MYOG (Figure 2(f)) (p<0.05). MYOG was also significantly upregulated in young myoblasts with both 10 μg/ml and 100 μg/ml C. vulgaris treatment (p<0.05) (Figure 2(f)). C. vulgaris treatment at 10 μg/ml in senescent myoblast significantly increased the expression of IGF1R (Figure 2(e)) and PTEN (Figure 2(h)) compared to control senescent myoblasts (p<0.05). The expression of TNNT1 and MYH2 however was not affected by C. vulgaris treatment in both young and senescent myoblasts (Figures 2(g)–2(i)).

Figure 2

Effects of C. vulgaris treatment on the gene expression on Day 0 of differentiation (a-i) and Days 1, 3, 5, and 7 of differentiation (j-r). The expression of gene was determined in control and C. vulgaris-treated young and senescent myoblasts. The data are presented as the means ± SD, n = 3. p<0.05 significantly different compared to control young on respective day, p<0.05 significantly different compared to control senescent on respective day, p<0.05 significantly different compared to control young on Day 0, p<0.05 significantly different compared to young treated with 10 μg/ml C. vulgaris on Day 0, p<0.05 significantly different compared to young treated with 100 μg/ml C. vulgaris on Day 0, p<0.05 significantly different compared to control senescent on Day 0, p<0.05 significantly different compared to senescent treated with 10 μg/ml C. vulgaris on Day 0, and p<0.05 significantly different compared to senescent treated with 100 μg/ml C. vulgaris on Day 0, with a post hoc Tukey HSD test.

3.3. Gene Expression on Days 1, 3, 5, and 7 of Differentiation

Pax7 was significantly upregulated in control young myoblasts on Days 5 and 7 of differentiation (Figure 2(j)) while MyoD1 in control young myoblasts was significantly upregulated on Day 5 of differentiation as compared to Day 0 (p<0.05) (Figure 2(k)). The expression of Myf5 was decreased in control young myoblasts on Days 1, 3, 5, and 7 of differentiation as compared to Day 0 (Figure 2(l)) while MEF2C in control young myoblasts was upregulated on Day 7 of differentiation (p<0.05) (Figure 2(m)). No significant change was observed on the expression of IGF1R (Figure 2(n)) and MYH2 (Figure 2(r)) on Day 1 till Day 7 of differentiation in control young myoblasts compared to Day 0 of differentiation. The expression of MYOG (Figure 2(o)) and PTEN (Figure 2(q)) in control young myoblasts was significantly increased on Days 3, 5, and 7 of differentiation while TNNT1 in control young myoblasts was significantly upregulated on Day 3 of differentiation as compared to Day 0 of differentiation (p<0.05) (Figure 2(p)).

In control senescent myoblasts, an upregulation of Pax7 (Figure 2(j)), TNNT1 (Figure 2(p)), and MYH2 (Figure 2(r)) was observed on Days 5 and 7 of differentiation while MyoD1 (Figure 2(k)) and Myf5 (Figure 2(l)) were upregulated on Days 1, 3, 5, and 7 of differentiation, and MEF2C (Figure 2(m)), MYOG (Figure 2(o)), and PTEN (Figure 2(q)) were upregulated on Days 3, 5, and 7 of differentiation compared to Day 0 (p<0.05). No significant change was observed in the expression of IGF1R in control senescent myoblasts on Day 1 till Day 7 of differentiation compared to Day 0 (Figure 2(n)).

Treatment with 10 μg/ml and 100 μg/ml C. vulgaris was found to upregulated Pax7 in young myoblasts on Days 3, 5, and 7 of differentiation and in senescent myoblasts on Day 7 of differentiation (p<0.05) (Figure 2(j)); upregulated MyoD1 in both young and senescent myoblasts on Day 5 of differentiation (p<0.05) (Figure 2(k)); upregulated Myf5 in senescent myoblasts on Day 1 and 3 of differentiation (p<0.05), (Figure 2(l)); upregulated MEF2C in both young and senescent myoblasts on Day 5 of differentiation (p<0.05) (Figure 2(m)); upregulated IGF1R in both young and senescent myoblasts on Days 5 and 7 of differentiation (p<0.05) (Figure 2(n)); upregulated MYOG in young myoblasts on Days 3 and 5 of differentiation and in senescent myoblasts on Days 5 and 7 of differentiation (p<0.05) (Figure 2(o)); upregulated TNNT1 in young myoblasts on Days 3, 5, and 7 (p<0.05) (Figure 2(p)); upregulated PTEN in young myoblasts on Days 5 and 7 of differentiation and in senescent myoblasts on Days 1, 3, 5, and 7 of differentiation (p<0.05) (Figure 2(q)); downregulated MYH2 in young myoblasts on Day 1 of differentiation and upregulated MYH2 in young myoblasts on Days 3 and 7 of differentiation as compared to control (p<0.05) (Figure 2(r)).

3.4. MyomiRs Expression on Day 0 of Differentiation

The expression of miR-133b, miR-206, and miR-486 in control senescent myoblasts was significantly decreased as compared to control young myoblasts (p<0.05) (Figures 3(a)–3(c)). A similar decrease in the expression of miR-133b, miR-206, and miR-486 in young myoblasts was observed with C. vulgaris treatment at 10 μg/ml and 100 μg/ml compared to control young (p<0.05). In senescent myoblasts however, treatment with 10 μg/ml C. vulgaris was found to increase the expression of miR-133b, miR-206, and miR-486 compared to control senescent (p<0.05) (Figures 3(a)–3(c)).

Figure 3

Effects of C. vulgaris treatment on the myomiRs expression at Day 0 of differentiation (a-c) and Days 1, 3, 5, and 7 of differentiation (d-f). The expression of myomiRs was determined in control and C. vulgaris-treated young and senescent myoblasts. The data are presented as the means ± SD, n = 3. p<0.05 significantly different compared to control young on respective day, p<0.05 significantly different compared to control senescent on respective day, p<0.05 significantly different compared to control young on Day 0, p<0.05 significantly different compared to young treated with 10 μg/ml C. vulgaris on Day 0, p<0.05 significantly different compared to young treated with 100 μg/ml C. vulgaris on Day 0, p<0.05 significantly different compared to control senescent on Day 0, p<0.05 significantly different compared to senescent treated with 10 μg/ml C. vulgaris on Day 0, and p<0.05 significantly different compared to senescent treated with 100 μg/ml C. vulgaris on Day 0, with a post hoc Tukey HSD test.

3.5. MyomiRs Expression on Days 1, 3, 5, and 7 of Differentiation

In control young myoblasts, the expression of miR-133b was significantly decreased on Days 1, 3, 5, and 7 of differentiation compared to Day 0 (p<0.05) (Figure 3(d)). However, no significant change was observed in the expression of miR-133b in control senescent myoblasts on Day 1 till Day 7 of differentiation. A similar reduction in the expression of miR-206 in control young myoblasts was observed on Days 1, 3, 5, and 7 of differentiation as compared to Day 0 (p<0.05) (Figure 3(e)) with no significant change being observed in control senescent myoblasts. The expression of miR-486 was not significantly changed in both control young and control senescent myoblasts (Figure 3(f)). Treatment with 10 μg/ml C. vulgaris significantly increased the expression of miR-133b and miR-206 in senescent myoblasts on Day 3 of differentiation compared to control (p<0.05) (Figures 3(d)-3(e)). Expression of miR-206 in young and senescent myoblasts was increased on Day 1 of differentiation with 100 μg/ml C. vulgaris treatment compared to control (p<0.05) (Figure 3(e)). 100 μg/ml C. vulgaris treatment also increased mir-206 expression in senescent myoblasts on Day 3 of differentiation compared to control (p<0.05). No significant change was observed in the expression of miR-486 with C. vulgaris treatment in young and senescent myoblasts (Figure 3(f)).

4. Discussion

Several mechanisms may contribute to sarcopenia and decrease the ability to reverse muscle atrophy in ageing. This includes decreased protein synthesis, reduction in neural function, hormonal deficit, chronic inflammation, oxidative stress, loss of mitochondrial function, inappropriate signaling in muscle due to inadequate nutrition, nuclear apoptosis, and reduction in satellite cell function [36]. In aged and sarcopenic muscles, satellite cell proliferation and differentiation may be diminished and consequently cause the decline in the regenerative potential. Modification in satellite cell function may affect differentiation and fusion of myoblast cells. It has been reported that, in senescent satellite cells, the level of MyoD protein and its DNA-binding activity were significantly reduced and delayed compared to young cells [33, 36] indicating the effect of ageing on myogenic regulatory factors (MFRs). Thus, elucidation of the mechanism involved is vital so that targeted intervention can be proposed.

The MRFs genes such as myogenic differentiation 1 protein (MyoD1), myogenic factor 5 (Myf5), myogenin (MYOG), and muscle-specific regulatory factor 4 (Mrf4) are myogenic specific which act for the activation of satellite cells [36]. The paired box protein (PAX7) controls the regulation of MYF5 and MYOD1 protein expression upon activation of quiescent satellite cells. The results of this study showed that the expression of IGF1R, MEF2C, Myf5, MyoD1, MYOG, Pax7, TNNT1, and PTEN was significantly decreased in control senescent myoblast cells as compared to control young myoblast cells on Day 0 of differentiation indicating downregulation of these genes during replicative senescence or cellular senescence of myoblast cells. The decrease in regenerative potential of aged muscles was reported to be associated with the reduction in satellite cell function and declined Pax7 pool of myogenic stem cells [36]. Another study reported that Pax3, Pax7, Myf5, MyoD, and MYOG were significantly downregulated in the myoblasts extracted from elderly subjects compared to young subjects [37]. However, the C. vulgaris-treated senescent myoblast cells demonstrated a significantly increased expression of IGF1R, MEF2C, Myf5, MYOG, Pax7, and PTEN on Day 0 of differentiation as compared to control senescent myoblast cells indicating its role in the regulation of MRFs expression.

Our previous study showed that C. vulgaris improves the regenerative capacity of young and senescent myoblasts and promotes myoblast differentiation. Treatment with C. vulgaris resulted in decreased percentage of senescent myoblast cells stained positive for SA-β-gal. Additionally, C. vulgaris treatment improved the formation of myotubes in senescent myoblast cells as branched and multinucleated myotubes were observed. Fusion index and maturation index were also significantly higher on Day 5 of differentiation for C. vulgaris-treated young myoblast cells and Day 7 of differentiation for C. vulgaris-treated senescent myoblast cells as compared to control myoblasts. The percentage of young and senescent myoblast cells positively stained for myogenin was also observed to be significantly increased on Day 3 of differentiation compared to control myoblasts [34]. Thus, the promotion of myogenic differentiation by C. vulgaris was indicated and its potential in promoting muscle regeneration was further proven in the gene expression observed in this study. However, no significant difference was observed on MYH2 gene expression on Day 0 of differentiation in control and C. vulgaris-treated myoblasts for both young and senescent myoblasts. This observation may indicate that this gene is not affected in senescence and its expression is not regulated by C. vulgaris.

Besides MRFs and their regulators, microRNAs have been demonstrated to be involved in myogenic differentiation by modulating the expression of MRFs family in myogenesis. The MRFs, such as MyoD, Myf5, and MYOG, will activate the expression of a collection of myogenic microRNAs. This includes miR-1, miR-133, and miR-206, which are also known as myomiRs [37, 38]. The findings of this study showed that the expression of miR-133b, miR-206, and miR-486 was significantly decreased in control senescent myoblast cells compared to control young myoblast cells on Day 0 of differentiation indicating the involvement of these microRNAs in cellular senescence of myoblasts. However, a significant increase was observed in C. vulgaris-treated senescent myoblast cells on Day 0 of differentiation which may results in the promotion of myoblasts proliferation. A previous study reported that myoblast extracted from elderly subjects showed downregulation of miR-133b [37].

After limited rounds of proliferation, the majority of satellite cells enter the myogenic differentiation program and begin to fuse with each other to form new myofibers [38]. The paired box 7 (Pax7) is one of the earliest markers during myogenesis. It has been reported that Pax7 regulates muscle marker genes such as Myf5 and MyoD towards differentiation [39]. Proliferating myoblasts continue to express Pax7. However, in contrast to their quiescent progenitors, it also expresses MyoD. A reduction in Pax7 along with the induction of muscle-specific transcription factor MYOG marks myoblast that have entered the differentiation phase and initiate cell cycle withdrawal [7]. The Pax7 was also reported to play a dual role in myogenesis regulation by activating commitment to the myogenic program and simultaneously prevent terminal differentiation [40]. Upon commitment to terminal differentiation, upregulation of MYOG will directly or indirectly downregulate Pax7. A high ratio of Pax7 to MyoD was observed in quiescent satellite cells to maintain satellite cells in their quiescent state. An intermediate ratio of Pax7 and MyoD will allow the satellite cells to proliferate, but not differentiate. However, satellite cells with a low Pax7 to MyoD ratio will begin to differentiate and a further reduction in Pax7 level was observed with activation of MYOG [38]. In this study, a similar finding on Pax7 and MyoD ratio was observed. The expression of Pax7 was increased in early days of differentiation but later decreased towards the end of differentiation for both young and senescent myoblasts while the expression of MyoD and MYOG was increased. However, with C. vulgaris treatment, the expression of Pax7 was increased compared to its untreated control on each day of differentiation, in both young and senescent myoblasts indicating the promotion of proliferation and differentiation of myoblast by the alga.

Coinciding with or occurring soon after the upregulation of MYOG, differentiating myoblasts will initiate the expression of various genes encoding for structural proteins, such as sarcomeric myosin which fuse into myotubes [7]. Proliferation of satellite cells leads to the formation of new stem cells, which is maintained in undifferentiated state and myogenic precursor cells that express MRFs for muscle differentiation, which includes MyoD, Myf5, and MYOG. The conversion from quiescent state to the activated state is rapidly followed by muscle differentiation, with myosin heavy chains (MyHCs) expression and myoblast fuse with each other for the formation of myotubes. The activated satellite cells will begin to express either MyoD or Myf5. However, most cells will express both Myf5 and MyoD simultaneously, in which MYOG will be expressed by the cells followed by the expression of both MyoD and Myf5. Many cells will ultimately express all MRFs simultaneously. The activation of satellite cells followed by proliferation and fusion will occur after one day of injury and up to seven days [41]. In another study, it was described that MyoD and Myf5 were quickly upregulated in young myoblasts during the first hours of differentiation, followed by the expression of cell cycle regulator p57 and MYOG [33]. However, in senescent myoblast, both delay expression and downregulation of MyoD were observed resulting in failure of Myf5 activation. The delayed expression of p57 and MYOG was also observed in senescent myoblast.

The results of our study showed that the expression of MyoD, Myf5, and MYOG was increased in both young and senescent myoblasts with the increasing number of differentiation days. The expression of these transcription factors was also significantly increased with the treatment of C. vulgaris, in both young and senescent myoblasts, thus confirming its potential in promoting myoblast differentiation. A previous study reported that myoblast coexpressing both Myf5 and MyoD exhibits the intermediate growth and differentiation propensities, with the expression of MyoD peaks in mid G₁ and Myf5 expression is maximal at the G₀ and G₂ phases of the cell cycle [38].

A previous study reported that serum response factor (SRF) positively regulates the expression of MyoD in proliferating myoblasts by binding to serum response element (SRE) within the MyoD regulatory region. However, SRF only drives low levels of MyoD expression due to its activity which is hindered by cyclin D1 induced dependent kinase 4 (Cdk4). Induction of myocyte enhancer factor-2 (MEF2) expression prior to differentiation enables MEF2 to outcompete SRF for the SRE binding site and consequently resulted in high levels of MyoD expression and initiation of differentiation [38]. The MEF2 family of human transcription factors consists of four proteins, MEF2A, MEF2B, MEF2C, and MEF2D, in which the expression of MEF2C was determined in this study. Our finding showed that the expression of MEF2C was increased in both young and senescent myoblasts throughout the differentiation day. Treatment with C. vulgaris was found to significantly increase the expression of MEF2C as compared to its control on respective day of differentiation. MEF2C is involved in the regulation of cytoskeletal structures and loss of MEF2C in skeletal muscle will result in improper sarcomere organization. Its isoform, MEF2Cα2, is predominantly expressed in skeletal muscle and promotes muscle-specific gene expression and myogenic differentiation [42, 43]. Thus upregulation of MEF2C observed in this study may indicate promotion of myogenic differentiation by C. vulgaris throughout the differentiation day.

Insulin-like growth factors (IGFs) exist as two isoforms, IGF-I and IGF-II that play vital roles in the regulation of satellite cell activity. The IGF exerts pleiotropic functions, such as anti-inflammation, cell migration, and stimulation of both proliferation and differentiation in satellite cells, which is mediated by the binding of IGF-I to IGF-I receptor (IGF1R), a ligand-activated receptor tyrosine kinase. The expression level of IGF1R is critical for the regulation of muscle development due to its function in directly regulating the intracellular responsiveness of muscle cell to the extracellular IGF signal. The activation of IGF1R in satellites cells will promote the expression of MRFs, such as MYOG. The IGF1R activates two primary signaling pathways: upon ligand binding, IGF1R becomes auto phosphorylated and induces the phosphatidylinositol 3-kinase (PI3K)/Akt pathway which is involved in myoblast differentiation and (2) IGF1R activates the Ras/Raf/extracellular response kinases (ERKs) cascade, resulting in the activation of other protein kinases and transcription factors for satellite cell proliferation [38, 44]. The results of this study showed that the expression of IGF1R was maintained throughout the differentiation in both young and senescent myoblast cells. However, with C. vulgaris treatment, the expression of IGF1R was significantly increased in young and senescent myoblasts suggesting promotion of myoblast differentiation by C. vulgaris.

Troponin T (TnT) is a central player in the calcium regulation of actin thin filament function and is essential for the contraction and relaxation of striated muscles. TnT exists as three isoforms, TNNT1 and TNNT3 which are specifically expressed in slow and fast twitch skeletal muscle fibers and TNNT2 which is expressed specifically in cardiac muscle [45]. The deletion of phosphatase and tensin homologue (PTEN) in quiescent satellite cells will cause spontaneous activation of satellite cells and undergo premature differentiation without proliferation, resulting in satellite cells depletion and defective regenerative function of skeletal muscle in response to injury. PTEN is vital for satellite cell homeostasis and the deletion of PTEN in embryonic myogenic progenitors resulted in defective and depleted satellite cells and consequently causes failure to myoblast regeneration [46, 47].

The myofibers consist of repeated actin and myosin myofibrils forming sarcomere, which is the basic functional unit of skeletal muscle which is involved in muscle contraction. Skeletal muscle fibers can be categorized into a slow-contracting/fatigue-resistant type and fast-contracting/fatigue-susceptible type muscle fibers. It differs in terms of their myosin heavy chains (MyHC) isoforms and types of metabolism [38]. Various forms of MyHCs are encoded by a large family of sarcomeric MYH genes expressed in striated muscles. The MYH2 gene produces MyHC-2A protein with fast type 2A fibers [48]. In this study, the expression of TNNT, PTEN, and MYH2 was significantly increased in both young and senescent myoblast cells during differentiation. The expressions of TNNT1, PTEN, and MYH2 were significantly increased with C. vulgaris treatment indicating its potential in the promotion of muscle differentiation and regeneration.

The microRNAs play an important role during muscle proliferation and differentiation by regulating the expression of a number of transcription factors and signaling molecules required for myogenesis [49]. miR-133b and miR-206 are located at human chromosome 6p12.2 with tissue specificity towards skeletal muscle, while mir-486 is located at human chromosome 8p11.21 with tissue specificity towards heart and skeletal muscle [16]. The function of each myomiR is summarized as follows: miR-133b promotes myoblasts differentiation and fusion, regeneration, alternative splicing regulation, chromatin remodeling, cell fate regulation, and proapoptotic; miR-206 promotes myoblast differentiation and regeneration, regeneration of neuromuscular synapses, chromatin remodeling, antiangiogenic, proapoptotic, oxidative stress control, and antimigration; and miR-486 promotes myoblast differentiation and fusion, alternative splicing regulation, antiapoptotic, and promigration [50]. Both miR-133b and miR-206 are required for skeletal muscle differentiation.

Generally, the cell signaling pathways targeted by miR-206 tend to have opposing functions to the regulatory pathways targeted by miR-133b, in which miR-206 acts to promote myogenic differentiation and miR-133b maintains the undifferentiated state and promotes cell growth; thus coexpression of myomiRs will help in the maintenance of homeostasis under normal cellular conditions. Since the downregulation of myostatin permits expression of miR-133b/-206 and MyoD, and MYOG binds to miR-206 promoter, it was suggested that miR-133b/-206 expression in the muscle may also be partly controlled by MyoD and MYOG [51]. In this study, the expression of miR-133b in young myoblasts was significantly increased during differentiation as compared to Day 0. A similar increase in miR-133b was not observed in senescent myoblasts during differentiation suggesting slower regenerative capacity in ageing. Treatment with C. vulgaris however was found to increase the expression of miR-133b in senescent myoblasts on Day 3 of differentiation. C. vulgaris treatment also significantly increased the expression of miR-206 in young myoblasts on Day 1 and Day 3 of differentiation while the expression of miR-206 in senescent myoblasts was increased only on Day 3 of differentiation. These findings may indicate the potential of C. vulgaris in promoting myogenic differentiation. A previous study reported that increased miR-206 expression in proliferating myoblasts was initiated by the proximal (PROX) promoter. miR-206 is located within the intron of linc-MD1 and is transcribed autonomously under the control of its own PROX promoter [51]. This microRNA is upregulated by MyoD and targets Pax3 and Pax7 mRNA for myogenic differentiation regulation. Thus, through this miR-206 mediated negative feedback mechanism, MyoD facilitates progression of myoblasts towards terminal differentiation [52, 53].

Figure 4 summarizes the myogenesis process which is controlled by a group of myogenic regulatory factors (MRFs) that command the progression from quiescence to activation, proliferation, and differentiation of muscle satellite cells (MSCs). This process will lead to the transformation of individual satellite cell into a syncytial contractile myofibers [38]. Quiescent muscle satellite cells (MSCs) are characterized by the expression of transcription factor Pax7. These MSCs will exit from the quiescent state and will be activated in the presence of stimuli such as damaged to the environment of satellite cells [38, 54]. The proliferating MSCs will later express MyoD1 and Myf5 for its activation and promotion of entry into the cell cycle, during muscle fiber development and muscle regeneration [54]. It will be further promoted to differentiate by inducing MyoD1 expression. The MYOG expression will initiate terminal differentiation and fusion in immature myotubes and the expression of myosin heavy chain (MYH2) in mature myotubes will activate muscle-specific structural and contractile genes, thereby impacting muscle function [38, 54]. Pax7 will be downregulated upon the activation of myogenic regulatory genes such as Myf5 and MyoD1 during the initiation of myogenesis [55]. Thus, the activated satellite cells will return to quiescence state. This is important to maintain the satellite cell pool for long term muscle integrity [38].

The phosphatidylinositol-3-kinases (PI3K)/Akt signaling pathway is the major signaling pathway which regulates muscle protein synthesis by modulating IGF-1 and insulin expression [4, 56] that promote protein synthesis and muscle hypertrophy by interacting with their respective tyrosine kinase receptors to phosphorylate the insulin receptor substrate (IRS-1). PI3K/Akt will then activate and stimulate mTOR to further promote protein synthesis [4] (Figure 4). The presence of low levels of reactive oxygen species (ROS) plays an important role in inducing the upregulation of growth factors such as IGF-1 [57].

The IGF-1 targets miR-206 and miR-133 while the PI3K/Akt pathway inhibits FoxO3 expression for the upregulation of miR-206 which resulted in the promotion of myoblast differentiation [50]. The miR-133 has a conserved and functional binding site in the 3′′-UTR of IGF-1R, which consequently declined in IGF-1R abundance [14]. The miR-133 will promote myoblast proliferation by targeting the mRNA for serum response factor (SRF) for miR-486 activation [54]. The miR-486 will be highly upregulated during muscle differentiation by targeting Pax7 and consequently accelerated myoblast differentiation [58]. While the miR-133b will suppress myoblast proliferation and promote myoblast differentiation via mitogen-activated protein kinase (MAPK) regulation, by downregulating its transducers, which includes fibroblast growth factor receptor 1 (FGFR1) and protein phosphatase 2A catalytic subunit (PP2AC) [59]. The MAPK is also responsible in ensuring sufficient myoblast accumulation for fusion and myotubes formation. Thus, MAPK negatively regulates miR-133b expression and therefore maintaining proliferation until differentiation induction by a significant upregulation of miR-133b occurs [50].

The expression of myomiRs is dependent on the expression of myogenic regulatory factors such as MyoD1, MYOG, and Mef2c which regulate the balance between myoblast proliferation and differentiation [55]. The Mef2c is also responsible for the promotion of miR-206 [50]. The inhibition of Pax7 by miR-486 will result in MyoD1 upregulation and further enhancement of its own expression since miR-486 is directly regulated by MyoD1 and SRF [58]. The PTEN is downregulated by miR-486, thereby increasing pAkt, which in turn phosphorylates FoxO resulting in its activation. A previous study reported that muscle wasting is limited by the inactivation of FoxO [14]. The downregulation of miR-486 in normal myoblasts also leads to impaired migration and myoblast fusion [50].

5. Conclusion

C. vulgaris may promote myogenic differentiation as indicated by the upregulation of MRFs and myomiRs expression in culture. The modulatory effects of C. vulgaris on the expression of MRFs and myomiRs may indicate its potential in promoting muscle regeneration and ameliorating sarcopenia.

Data Availability

The data used to support the findings of this study are included within the article.

Conflicts of Interest

The authors declare no conflict of interest.

Funding

This work was financially supported by the Ministry of Education (MOE) Malaysia [grant number: FRGS/2/2014/SKK01/UKM/01/1]; and the Universiti Kebangsaan Malaysia [grant number: UKM-FF-2016-318].

Acknowledgments

The authors are grateful to the University of Malaya Algae Culture Collection (UMACC, Malaysia) for supplying the stock of C. vulgaris. We are also thankful for the contribution from all researchers and staffs of Department of Biochemistry and Department of Anatomy, Faculty of Medicine, Universiti Kebangsaan Malaysia Medical Centre.

References

G. D. Cartee, R. T. Hepple, M. M. Bamman, and J. R. Zierath, “Exercise promotes healthy aging of skeletal muscle,” Cell Metabolism, vol. 23, no. 6, pp. 1034–1047, 2016.
View at: Publisher Site | Google Scholar
I. H. Rosenberg, “Sarcopenia: origins and clinical relevance,” Journal of Nutrition, vol. 127, no. 5, pp. 990S–991S, 1997.
View at: Publisher Site | Google Scholar
M. Muscaritoli, S. D. Anker, J. Argilés et al., “Consensus definition of sarcopenia, cachexia and pre-cachexia: joint document elaborated by Special Interest Groups (SIG) “cachexia-anorexia in chronic wasting disease” and “nutrition in geriatrics”,” Clinical Nutrition, vol. 29, no. 2, pp. 154–159, 2010.
View at: Publisher Site | Google Scholar
M. M. Ziaaldini, E. Marzetti, A. Picca, and Z. Murlasits, “Biochemical pathways of sarcopenia and their modulation by physical exercise: A narrative review,” Frontiers in Medicine, vol. 4, no. 167, pp. 1–8, 2017.
View at: Google Scholar
W. K. Mitchell, J. Williams, P. Atherton, M. Larvin, J. Lund, and M. Narici, “Sarcopenia, dynapenia, and the impact of advancing age on human skeletal muscle size and strength; a quantitative review,” Frontiers in Physiology, vol. 3, article 260, pp. 1–18, 2012.
View at: Publisher Site | Google Scholar
D. Costamagna, P. Costelli, M. Sampaolesi, and F. Penna, “Role of inflammation in muscle homeostasis and myogenesis,” Mediators of Inflammation, vol. 2015, Article ID 805172, 14 pages, 2015.
View at: Publisher Site | Google Scholar
Z. Yablonka-Reuveni, “The skeletal muscle satellite cell: still young and fascinating at 50,” Journal of Histochemistry & Cytochemistry, vol. 59, no. 12, pp. 1041–1059, 2011.
View at: Publisher Site | Google Scholar
J. M. Hernández-Hernández, E. G. García-González, C. E. Brun, and M. A. Rudnicki, “The myogenic regulatory factors, determinants of muscle development, cell identity and regeneration,” Seminars in Cell & Developmental Biology, vol. 72, pp. 10–18, 2017.
View at: Publisher Site | Google Scholar
M. Buckingham and P. Rigby, “Gene regulatory networks and transcriptional mechanisms that control myogenesis,” Developmental Cell, vol. 28, no. 3, pp. 225–238, 2014.
View at: Publisher Site | Google Scholar
M. Li and J. C. I. Belmonte, “Genetic rejuvenation of old muscle,” Nature, vol. 506, article 7488, pp. 304-305, 2014.
View at: Publisher Site | Google Scholar
P. Sousa-Victor, S. Gutarra, L. García-Prat et al., “Geriatric muscle stem cells switch reversible quiescence into senescence,” Nature, vol. 506, pp. 1–16, 2014.
View at: Publisher Site | Google Scholar
E. Bengal, E. Perdiguero, A. L. Serrano, and P. Muñoz-Cánoves, “Rejuvenating stem cells to restore muscle regeneration in aging,” F1000Research, vol. 6, pp. 1–10, 2017.
View at: Google Scholar
K. Felekkis, C. T. E. Stefanou, and C. Deltas, “Micrornas: anewly described class of encoded molecules that play a role in health and disease,” Hippokratia, vol. 14, no. 4, pp. 236–240, 2010.
View at: Google Scholar
X. H. Wang, “MicroRNA in myogenesis and muscle atrophy,” Current Opinion in Clinical Nutrition & Metabolic Care, vol. 16, no. 3, pp. 258–266, 2013.
View at: Publisher Site | Google Scholar
M. Ha and V. N. Kim, “Regulation of microRNA biogenesis,” Nature Reviews Molecular Cell Biology, vol. 15, pp. 509–524, 2014.
View at: Publisher Site | Google Scholar
J. Siracusa, N. Koulmann, and S. Banzet, “Circulating myomiRs: a new class of biomarkers to monitor skeletal muscle in physiology and medicine,” Journal of Cachexia, Sarcopenia and Muscle, vol. 9, no. 1, pp. 20–27, 2018.
View at: Publisher Site | Google Scholar
H. S. Lee, H. J. Park, and M. K. Kim, “Effect of Chlorella vulgaris on lipid metabolism in Wistar rats fed high fat diet,” Nutrition Research and Practice, vol. 2, no. 4, pp. 204–210, 2008.
View at: Publisher Site | Google Scholar
G. P. Diniz and D. Z. Wang, “Regulation of skeletal muscle by micrornas,” Comprehensive Physiology, vol. 6, no. 3, pp. 1279–1294, 2011.
View at: Google Scholar
WHO, World Report on Ageing And Health, World Health Organization, 2015.
P. Spolaore, C. Joannis-Cassan, E. Duran, and A. Isambert, “Commercial applications of microalgae,” Journal of Bioscience and Bioengineering, vol. 101, no. 2, pp. 87–96, 2006.
View at: Publisher Site | Google Scholar
S. S. Ahluwalia and D. Goyal, “Microbial and plant derived biomass for removal of heavy metals from wastewater,” Bioresource Technology, vol. 98, no. 12, pp. 2243–2257, 2007.
View at: Publisher Site | Google Scholar
C. F. Gao, Y. Zhai, Y. Ding, and Q. Wu, “Application of sweet sorghum for biodiesel production by heterotrophic microalga Chlorella protothecoides,” Applied Energy, vol. 87, no. 3, pp. 756–761, 2010.
View at: Publisher Site | Google Scholar
L. Krienitz, V. A. Huss, and C. Bock, “Chlorella: 125 years of the green survivalist,” Trends in Plant Science, vol. 20, no. 2, pp. 67–69, 2015.
View at: Publisher Site | Google Scholar
C. Safi, B. Zebib, O. Merah, P. Pontalier, and C. Vaca-Garcia, “Morphology, composition, production, processing and applications of Chlorella vulgaris: A review,” Renewable & Sustainable Energy Reviews, vol. 35, pp. 265–278, 2014.
View at: Publisher Site | Google Scholar
H. Jeong, H. J. Kwon, and M. K. Kim, “Hypoglycemic effect of Chlorella vulgaris intake in type 2 diabetic Goto-Kakizaki and normal Wistar rats,” Nutrition Research and Practice, vol. 3, no. 1, pp. 23–30, 2009.
View at: Publisher Site | Google Scholar
O. Aizzat, S. Yap, H. Sopiah et al., “Modulation of oxidative stress by Chlorella vulgaris in streptozotocin (STZ) induced diabetic Sprague-Dawley rats,” Advances in Medical Sciences, vol. 55, no. 2, pp. 281–288, 2010.
View at: Publisher Site | Google Scholar
W. Z. W. Ngah and Y. A. M. Yusof, “Chemopreventive effect of chlorella vulgaris in choline deficient diet and ethionine induced liver carcinogenesis in rats,” International Journal of Cancer Research, vol. 2, no. 3, pp. 234–241, 2006.
View at: Publisher Site | Google Scholar
E. S. M. Azamai, S. Sulaiman, S. H. M. Habib et al., “Chlorella vulgaris triggers apoptosis in hepatocarcinogenesis-induced rats,” Journal of Zhejiang University SCIENCE B, vol. 10, no. 1, pp. 14–21, 2009.
View at: Publisher Site | Google Scholar
N. A. Mukti, S. Sulaiman, S. M. Saad, and H. Basari, “Chlorella vulgaris menunjukkankesanantioksidandan antitumor terhadapkanserhepardalamkajian in vivo dan in vitro,” SainsMalaysiana, vol. 38, no. 3, pp. 773–784, 2009.
View at: Google Scholar
S. M. Saad, Y. A. M. Yusof, and W. Z. W. Ngah, “Comparison between locally produced Chlorella vulgaris and Chlorella vulgaris from Japan on proliferation and apoptosis of liver cancer cell line, HepG2,” Malaysian Journal of Biochemistry and Molecular Biology, vol. 13, no. 1, pp. 32–36, 2006.
View at: Google Scholar
K. Kitada, S. Machmudah, M. Sasaki et al., “Supercritical CO2 extraction of pigment components with pharmaceutical importance from Chlorella vulgaris,” Journal of Chemical Technology and Biotechnology, vol. 84, no. 5, pp. 657–661, 2009.
View at: Publisher Site | Google Scholar
Y. Panahi, B. Pishgoo, H. R. Jalalian et al., “Investigation of the effects of Chlorella vulgaris as an adjunctive therapy for dyslipidemia: Results of a randomised open-label clinical trial,” Nutrition & Dietetics, vol. 69, no. 1, pp. 13–19, 2012.
View at: Publisher Site | Google Scholar
A. Bigot, V. Jacquemin, F. Debacq-Chainiaux et al., “Replicative aging down-regulates the myogenic regulatory factors in human myoblasts,” Biology of the Cell, vol. 100, no. 3, pp. 189–199, 2008.
View at: Publisher Site | Google Scholar
N. Z. Azlan, Y. A. M. Yusof, E. Alias, and S. Makpol, “Chlorella vulgaris improves the regenerative capacity of young and senescent myoblasts and promotes muscle regeneration,” Oxidative Medicine and Cellular Longevity, vol. 2019, Article ID 3520789, 16 pages, 2019.
View at: Publisher Site | Google Scholar
T. D. Schmittgen and K. J. Livak, “Analyzing real-time PCR data by the comparative CT method,” Nature Protocols, vol. 3, no. 6, pp. 1101–1108, 2008.
View at: Publisher Site | Google Scholar
S. E. Alway, M. J. Myers, and J. S. Mohamed, “Regulation of satellite cell function in sarcopenia,” Frontiers in Aging Neuroscience, vol. 6, article 246, pp. 1–15, 2014.
View at: Publisher Site | Google Scholar
E. S. Di Filippo, R. Mancinelli, T. Pietrangelo et al., “Myomir dysregulation and reactive oxygen species in aged human satellite cells,” Biochemical and Biophysical Research Communications, vol. 473, no. 2, pp. 462–470, 2016.
View at: Publisher Site | Google Scholar
H. Yin, F. Price, and M. A. Rudnicki, “Satellite cells and the muscle stem cell niche,” Physiological Reviews, vol. 93, no. 1, pp. 23–67, 2013.
View at: Publisher Site | Google Scholar
W. Izumi, Y. Takuma, R. Ebihara, W. Mizunoya, R. Tatsumi, and M. Nakamura, “Paired box 7 inhibits differentiation in 3T3-L1 preadipocytes,” Animal Science Journal, vol. 89, no. 8, pp. 1214–1219, 2018.
View at: Publisher Site | Google Scholar
H. C. Olguín and A. Pisconti, “Marking the tempo for myogenesis: Pax7 and the regulation of muscle stem cell fate decisions,” Journal of Cellular and Molecular Medicine, vol. 16, no. 5, pp. 1013–1025, 2012.
View at: Publisher Site | Google Scholar
S. Ciciliot and S. Schiaffino, “Regeneration of mammalian skeletal muscle. Basic mechanisms and clinical implications,” Current Pharmaceutical Design, vol. 16, no. 8, pp. 906–914, 2010.
View at: Publisher Site | Google Scholar
M. J. Potthoff, M. A. Arnold, J. McAnally, J. A. Richardson, R. Bassel-Duby, and E. N. Olson, “Regulation of skeletal muscle sarcomere integrity and postnatal muscle function by Mef2c,” Molecular and Cellular Biology, vol. 27, no. 23, pp. 8143–8151, 2007.
View at: Publisher Site | Google Scholar
J. R. Pon and M. A. Marra, “MEF2 transcription factors: developmental regulators and emerging cancer genes,” Oncotarget, vol. 7, no. 3, pp. 2297–2312, 2016.
View at: Publisher Site | Google Scholar
M.-B. Huang, H. Xu, S.-J. Xie, H. Zhou, and L.-H. Qu, “Insulin-like growth factor-1 receptor is regulated by microRNA-133 during skeletal myogenesis,” PLoS ONE, vol. 6, no. 12, Article ID e29173, 2011.
View at: Publisher Site | Google Scholar
B. Wei and J.-P. Jin, “TNNT1, TNNT2, and TNNT3: Isoform genes, regulation, and structure-function relationships,” Gene, vol. 582, no. 1, pp. 1–13, 2016.
View at: Publisher Site | Google Scholar
F. Yue, P. Bi, C. Wang, J. Li, X. Liu, and S. Kuang, “Conditional loss of Pten in myogenic progenitors leads to postnatal skeletal muscle hypertrophy but age-dependent exhaustion of satellite cells,” Cell Reports, vol. 17, no. 9, pp. 2340–2353, 2016.
View at: Publisher Site | Google Scholar
F. Yue, P. Bi, C. Wang et al., “Pten is necessary for the quiescence and maintenance of adult muscle stem cells,” Nature Communications, vol. 8, no. 1, pp. 1–13, 2017.
View at: Publisher Site | Google Scholar
S. Schiaffino, “Muscle fiber type diversity revealed by anti‐myosin heavy chain antibodies,” FEBS Journal, vol. 285, no. 20, pp. 3688–3694, 2018.
View at: Publisher Site | Google Scholar
T. Sadkowski, A. Ciecierska, J. Oprządek, and E. Balcerek, “Breed-dependent microRNA expression in the primary culture of skeletal muscle cells subjected to myogenic differentiation,” BMC Genomics, vol. 19, no. 109, pp. 1–14, 2018.
View at: Publisher Site | Google Scholar
M. Horak, J. Novak, and J. Bienertova-Vasku, “Muscle-specific microRNAs in skeletal muscle development,” Developmental Biology, vol. 410, no. 1, pp. 1–13, 2016.
View at: Publisher Site | Google Scholar
K. R. Mitchelson and W.-Y. Qin, “Roles of the canonical myomiRs miR-1, -133 and -206 in cell development and disease,” World Journal of Biological Chemistry, vol. 6, no. 3, p. 162, 2015.
View at: Publisher Site | Google Scholar
J.-F. Chen, Y. Tao, J. Li et al., “microRNA-1 and microRNA-206 regulate skeletal muscle satellite cell proliferation and differentiation by repressing Pax7,” The Journal of Cell Biology, vol. 190, no. 5, pp. 867–879, 2010.
View at: Publisher Site | Google Scholar
H. Hirai, M. Verma, S. Watanabe, C. Tastad, Y. Asakura, and A. Asakura, “MyoD regulates apoptosis of myoblasts through microRNA-mediated down-regulation of Pax3,” The Journal of Cell Biology, vol. 191, no. 2, pp. 347–365, 2010.
View at: Publisher Site | Google Scholar
D. Salvatore, W. S. Simonides, M. Dentice, A. M. Zavacki, and P. R. Larsen, “Thyroid hormones and skeletal muscle - New insights and potential implications,” Nature Reviews Endocrinology, vol. 10, no. 4, pp. 206–214, 2014.
View at: Publisher Site | Google Scholar
C. G. Crist and M. Buckingham, “microRNAs gain magnitude in muscle,” Cell Cycle, vol. 8, no. 22, pp. 3627-3628, 2014.
View at: Publisher Site | Google Scholar
S. K. Powers, A. B. Morton, B. Ahn, and A. J. Smuder, “Redox control of skeletal muscle atrophy,” Free Radical Biology & Medicine, vol. 98, pp. 208–217, 2016.
View at: Publisher Site | Google Scholar
E. Barbieri and P. Sestili, “Reactive oxygen species in skeletal muscle signaling,” Journal of Signal Transduction, vol. 2012, Article ID 982794, 17 pages, 2012.
View at: Publisher Site | Google Scholar
B. K. Dey, J. Gagan, and A. Dutta, “miR-206 and -486 induce myoblast differentiation by downregulating Pax7,” Molecular and Cellular Biology, vol. 31, no. 1, pp. 203–214, 2011.
View at: Publisher Site | Google Scholar
Y. Feng, L. Niu, W. Wei et al., “A feedback circuit between miR-133 and the ERK1/2 pathway involving an exquisite mechanism for regulating myoblast proliferation and differentiation,” Cell Death & Disease, vol. 4, no. 11, p. e934, 2013.
View at: Google Scholar

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Copyright © 2019 Nurhazirah Zainul Azlan et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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