APOBEC2 negatively regulates myoblast differentiation in muscle regeneration

The International Journal of Biochemistry & Cell Biology 85 (2017) 91–101

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Research paper

APOBEC2 negatively regulates myoblast differentiation in muscle regeneration Hideaki Ohtsubo a , Yusuke Sato a,b , Takahiro Suzuki a,c,1 , Wataru Mizunoya a , Mako Nakamura d , Ryuichi Tatsumi a,∗ , Yoshihide Ikeuchi a,∗ a

Department of Animal and Marine Bioresource Sciences, Graduate School of Agriculture, Kyushu University, Hakozaki, Fukuoka 8128581, Japan Department of Bio-Productive Science, Utsunomiya University, Utsunomiya, Tochigi 3218505, Japan c Department of Molecular and Developmental Biology, Kawasaki Medical School, Kurashiki, Okayama 7010192, Japan d Graduate School of Agriculture, Kyushu University, Hakozaki, Fukuoka 8128581, Japan b

a r t i c l e

i n f o

Article history: Received 21 November 2016 Received in revised form 6 February 2017 Accepted 7 February 2017 Available online 12 February 2017 Keywords: APOBEC2 Muscle regeneration Myoblast differentiation Myogenin Myotube

a b s t r a c t Recently we found that the deficiency of APOBEC2, a member of apoB mRNA editing enzyme, catalytic polypeptide-like family, leads to a diminished muscle mass and increased myofiber with centrally-located nuclei known as dystrophic phenotypes. APOBEC2 expression is predominant in skeletal and cardiac muscles and elevated exclusively at the early-differentiation phase of wild-type (WT) myoblast cultures; however the physiological significance is still un-known. Here we show that APOBEC2 is a key negative regulator of myoblast differentiation in muscle regeneration. APOBEC2-knockout (A2KO) mice myoblast cultures displayed a normal morphology of primary myotubes along with earlier increase in fusion index and higher expression levels of myosin heavy chain (MyHC), myogenin and its cooperating factor MEF2C than WT myoblasts. Similar response was observable in APOBEC2-knockdown cultures of WT myoblasts that were transfected with the specific siRNA at the differentiation phase (not proliferation phase). Importantly, cardiotoxin-injured A2KO gastrocnemius muscle provided in vivo evidence by showing larger up-regulation of neonatal MyHC and myogenin and hence earlier regeneration of myofiber structures with diminished cross-sectional areas and minimal Feret diameters. Therefore, the findings highlight a promising role for APOBEC2 in normal progression of regenerative myogenesis at the early-differentiation phase upon muscle injury. © 2017 Elsevier Ltd. All rights reserved.

1. Introduction Satellite cells are resident myogenic stem cells and are normally found in a quiescent state in adult animals (Mauro, 1961; Snow, 1997). In response to muscle injury, the cells are quickly activated in a hepatocyte growth factor/NO radical-dependent manner; once activated, they proliferate into myoblasts, differentiate, and fuse

Abbreviations: APOBEC, apoB mRNA editing enzyme catalytic polypeptide-like; CTX, cardiotoxin; DMEM, Dulbecco’s modified Eagle’s medium; FBS, fetal bovine serum; HS, horse serum; MEF2C, myocyte enhancer factor 2C; MyHC, myosin heavy chain. ∗ Corresponding authors at: Ryuichi Tatsumi or Yoshihide Ikeuchi, Department of Animal and Marine Bioresource Sciences, Graduate School of Agriculture, Kyushu University, Hakozaki 6-10-1, Higashi-ku, Fukuoka 8128581, Japan. E-mail addresses: [email protected] (R. Tatsumi), [email protected] (Y. Ikeuchi). 1 Present address: Cell and Tissue Biology Lab., Research Faculty of Agriculture, Hokkaido University, Sapporo, Hokkaido 0608589, Japan. http://dx.doi.org/10.1016/j.biocel.2017.02.005 1357-2725/© 2017 Elsevier Ltd. All rights reserved.

to damaged myofibers (or form new fibers) (Tatsumi et al., 1998; Anderson, 2000; Hawke and Garry, 2001; Tatsumi and Allen, 2008; Tatsumi, 2010). These sequential events are highly regulated by a variety of intracellular and extracellular molecules including transcription factors, cell-membrane and nuclear receptors, growth factors, cytokines, and hormones (Bentzinger et al., 2010; Turner and Badylak, 2012). The mechanisms that coordinate these various molecules are not yet fully understood, particularly for the fascinating and dramatic steps of myoblast differentiation and fusion in regenerative myogenesis. The family of myogenic regulatory factor (MRF) proteins, also called basic helix-loop-helix (bHLH) transcription factors, includes four MyoD, Myf5, myogenin, and MRF4; together they play crucial roles in myoblast differentiation and fusion. MyoD expression peaks in the mid-G1 phase of the cell cycle; MyoD expression permits myoblast differentiation and ultimately leads to cell-cycle arrest, while Myf5 expression begins shortly after MyoD expression begins and helps determine the cell fate toward the myogenic lineage (Ishibashi et al., 2005; De Falco and De Luca, 2006; Gayraud-

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Morel et al., 2007; Ustanina et al., 2007). Importantly, myogenin is up-regulated at the early-differentiation phase and regulates the fusion of myoblasts into myotubes (immature myofibers) that is followed by MRF4-dependent myotube maturation (Venuti et al., 1995; Zhang et al., 1995; Zanou and Gailly, 2013). APOBEC2 is a member of the apolipoprotein B mRNA editing enzyme, catalytic polypeptide-like family of proteins having zinc-dependent cytidine deaminase activity (Liao et al., 1999; Navaratnam and Sarwar, 2006; Conticello, 2008). Recently, we found that a deficiency of APOBEC2 leads to a diminished mouse muscle mass and increased myofiber with centrally-located nuclei known as dystrophic phenotypes (Sato et al., 2009, 2010). Experiments also demonstrated that APOBEC2 expression is predominant in skeletal and cardiac muscles and in cultured wild-type (WT) myoblasts, its expression rises exclusively in the earlydifferentiation phase along with its intracellular translocation from the nucleus to the cytoplasm of myoblasts. Additional information was provided by a Zebrafish study to demonstrate that APOBEC2 protein is localized at the Z-line region of sarcomeres along the myofibril and that APOBEC2 knockdown leads to a myopathic phenotype (Etard et al., 2010). These reports encourage a model in which APOBEC2 mediates differentiation/fusion of myoblasts responsible for the formation of myotubes during muscle regeneration from injury, even though there is a limited number of references on APOBEC2 biology, including some recent studies to show that APOBEC2 may participate in the left-right axis determination in Xenopus embryogenesis (Vonica et al., 2011) and retina regeneration independent on cytidine deaminase activity in zebrafish (Powell et al., 2012, 2014). Thus it is important to clarify the biological and physiological roles of APOBEC2 in muscle regeneration. The present study was designed to examine the effect of APOBEC2 knockout in primary satellite cell cultures and in vivo in a model of muscle injury. Results clearly showed that APOBEC2 deficiency accelerates the formation of primary myotubes in culture and increases the up-regulation of differentiation markers including myosin heavy chain (MyHC), myogenin, and myocyte enhancer factor 2C (MEF2C) which cooperates with myogenin. The enhanced differentiation and fusion observed in culture after APOBEC2 knockout may explain the accelerated restoration of myofiber structures in regeneration of muscle after in vivo injury. Therefore, APOBEC2 functions as a negative regulator in regeneration of wild-type muscle, and its role centers on modulating processes of myoblast differentiation and fusion. 2. Materials and methods 2.1. Animal care and use APOBEC2-deficient mice (A2KOs; C57BL/6 as the background strain) were generated by Dr. Neuberger (Medical Research Council Laboratory of Molecular Biology, United Kingdom) and bred in our laboratory (Mikl et al., 2005). Inbred C57BL/6 mice were used as wild-type (WT) controls. All animal experiments were conducted in strict accordance with the Guidelines for Proper Conduct of Animal Experiments published by the Science Council of Japan and ethics approvals from the Kyushu University Institutional Review Board (approval nos. 20–12, 23–62, A22–218, A24–075, A26–078, and A28–090). 2.2. Satellite cell isolation and primary culture Satellite cells were isolated from body muscles of 4-monthold WT and A2KO mice according to Jackson et al. (1999) with a slight modification and used for all culture experiments (except

for APOBEC2-knockdown experiments shown in Fig. 3). Briefly, muscle was treated 0.2% collagenase type II (LS0041 from Worthington, Lakewood, NJ) for 45 min followed by 0.1% trypsin (15400, Thermo Fisher Scientific, Waltham, MA) for 45 min at 37 ◦ C. Cells were collected by centrifugation and plated on a non-coated dish for 2 h. Non-adherent cells were collected and plated on plastic dishes and 8-well chamber slides (for measuring fusion index) coated with Matrigel (356234, BD Bioscience, Bedford, MA) and maintained in Dulbecco’s modified Eagle’s medium (DMEM) containing 20% fetal bovine serum (FBS; Invitrogen, Grand Island, NY), 1% antibiotic-antimycotic mixture (15240-062, Thermo Fisher Scientific) and 0.5% gentamicin (15710-64) (proliferation medium) until about 70% confluence. Subsequently, cell differentiation was induced by serum starvation by altering the culture medium to DMEM containing 5% horse serum (HS; 16050-122, Invitrogen), 1% antibiotic-antimycotic and 0.5% gentamicin (differentiation medium). Myotube formation was evaluated by the measurement of fusion indices (the number of myonuclei as a percentage of total nuclei, visualized with 4 ,6-diamidino-2-phenylindole, DAPI) at five random fields for each well (at least 5000 nuclei counted per well, n = 3 wells per group). Companion satellite-cell cultures, prepared at the same time, were immunostained for the presence of Pax7 at 48 h post-plating, using a monoclonal anti-Pax7 antibody (clone PAX7, Developmental Studies Hybridoma Bank, Iowa City, IA) and Alexa Fluor 594-labeled anti-mouse IgG secondary antibody (1:500 dilution; A11005 of Thermo Fisher Scientific), in order to determine the percentage of myogenic cells present; cultures with less than 95% Pax7-positive cells were not used for experiments. 2.3. Muscle injury Mice were anesthetized by intraperitoneal injection (ip) with a mixture of agents (0.3 ␮g medetomidine hydrochloride, 4 ␮g midazolam, and 5 ␮g butorphanol tartrate per g body weight) and then had an intramuscular injection of 50 ␮l of 10 ␮M cardiotoxin (CTX; the peptidic myotoxin isolated from venom of the Naja mossambica mossambica; Sigma-Aldrich, St. Louis, MO) into each medial and lateral head of gastrocnemius muscle as optimized previously (Sakaguchi et al., 2014). CTX-injected muscles were dissected at day 0 (before injection), or 4, 7, 14, and 21 days after injection and frozen in isopentane (26405-65, Nacalai Tesque, Kyoto, Japan) cooled with liquid nitrogen. Muscle cryosections (mid-belly portions, about 7-␮m thickness) were stained with hematoxylin (8650; Sakurai Finetek Japan, Tokyo, Japan) and eosin (8659; Sakurai Finetek Japan) and dehydrated with ethanol. Myofiber cross-sectional area and minimal Feret diameter (at least 800 fibers per group, n = 3) of muscles from WT and A2KO mice were measured using ImageJ 1.34 s software (originally developed by Dr. Wayne Rasband, National Institutes of Health, Bethesda, MD, USA). 2.4. Immunohistochemistry Primary cultures of satellite cells were fixed with 4% paraformaldehyde (09154-85, Nacalai Tesque) and permeabilized with 0.1% Triton X-100 in phosphate-buffered saline (PBS). Subsequently, cells were blocked with 3% bovine serum albumin (BSA; A8022, Sigma-Aldrich) in 0.1% polyethylene sorbitan monolaurate (Tween 20)-Tris buffered saline (TTBS) at room temperature for 1 h before incubation with monoclonal anti-pan MyHC antibody (1:500 dilution; clone MF20, IC4470F from R&D Systems, Minneapolis, MN) and polyclonal anti-MyoD antibody (1:200 dilution; sc-760 from Santa Cruz Biotechnology, Santa Cruz, CA) and with Alexa Fluor 594-labeled anti-mouse secondary antibody (1:500 dilution) and Alexa Fluor 488 anti-rabbit secondary antibody (1:500 dilution; A-21441, Invitrogen), respectively. Cells were counter-stained with

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DAPI (1:1000 dilution; D523 of DOJINDO Laboratories, Kumamoto, Japan). Muscle sections (mid-belly portions) were fixed by incubation in PBS at 90 ◦ C for 10 min and blocked with normal donkey blocking solution (2% donkey serum, 1% BSA, 0.1% cold fish skin gelatin, 0.1% Triton X-100, 0.05% Tween 20, 0.05% sodium azide, 0.01% avidin and 100 mM mannose in PBS). Subsequently, sections were incubated in a mixture of monoclonal anti-developmental MyHC (1:100 dilution; clone RNMy2/9D2, NCL-MHCd from Leica Biosystems, Newcastle, United Kingdom; recognizes both embryonic and neonatal MyHC isoforms) and polyclonal anti-laminin antibodies (1:500; L9393 from Sigma-Aldrich). Alexa Fluor 594-labeled antimouse IgG (1:500 dilution; A21201, Invitrogen) and Alexa Fluor 488 anti-rabbit IgG (1:500 dilution; A-21441, Invitrogen) were used as secondary antibodies, respectively. Sections were counter-stained with DAPI and observed under a Leica DMI6000B fluorescence microscope equipped with a DFC365FX digital camera and LAS AF 3.1.0 software.

Table 1 PCR primer sets for mouse target genes and the reference.

2.5. Western blotting

2.7. APOBEC2-knockdown cultures

Whole-cell lysates from cell cultures were applied to 10% polyacrylamide gels for electrophoresis under reducing conditions and transferred to nitrocellulose membranes as described previously (Tatsumi et al., 2009). Blots were blocked with 5% powdered milk in 0.1% Tween 20-Tris buffered saline (TTBS) before incubation overnight with antigen-affinity-purified primary antibodies against pan MyHC (1:5000 dilution; IC4470F, MF20 monoclonal, R&D Systems), myogenin (1:1000 dilution; sc-12732, F5D monoclonal, Santa Cruz Biotechnology), APOBEC2 (1:500; HPA017957, polyclonal, Sigma-Aldrich), fast MyHC (1:1000; M4276, MY-32 monoclonal, Sigma-Aldrich) and slow MyHC (1:1000; M8421, NOQ7.5.4D monoclonal, Sigma-Aldrich) in CanGetSignal solution 1 (NKB-101; Toyobo, Osaka, Japan) containing 0.05% sodium azide. Membranes were subsequently treated for 1 h with affinitypurified anti-mouse IgG antibody conjugated to horseradish peroxidase (P0447, Dako cytomation, Glostrup, Denmark) or antirabbit IgG antibody (P0399, Dako cytomation) diluted 1:5000 in CanGetSignal solution 2; immuno-reactive bands were visualized with ECL Select solution (RPN2235, GE Healthcare, Little Chalfont, UK) for APOBEC2 or with ECL solution (RPN2106, GE healthcare) for other target proteins.

C57BL/6J mouse-derived myoblasts (Ojima et al., 2004, 2007, 2015), a kind gift from Dr. Koichi Ojima, NARO Institute of Livestock and Grassland Science (Tsukuba, Ibaraki, Japan), were used only for APOBEC2-knockdown experiments (Fig. 3). Myoblasts were plated on dishes coated with 20% Matrigel-DMEM and maintained for 24 h in 20% FBS-Ham’s F10 Nutrient Mixture medium (11550043; Invitrogen). Subsequently, cell differentiation was induced by replacing the medium with differentiation medium (5% HS-DMEM) that additionally contained 100 nM small interfering RNA (siRNA) in DharmaFECT siRNA Transfection Reagent 1 (T-2001-01; Thermo Fisher Scientific). The APOBEC2 siRNAs used here were Stealth RNAiTM duplexes (Invitrogen) that were designed with BLOCK-iT RNAi Designer and formulated specifically to block mouse target transcripts (Table 2); a cocktail of three siRNAs (427, 738, and 765; 33.3 nM each) was added to the differentiation medium. Cultures transfected with Allstars Negative Control siRNA (1027281; Qiagen) served as the control group. Cells were harvested at 2 days after combined differentiation and siRNA-lipofection and analyzed for target-gene expression by real-time RT-qPCR and Western blotting.

2.6. Real-time RT-qPCR

Student’s t-test was employed for statistical analysis of experimental results using Microsoft Excel X for Macintosh. Data are represented as mean ± SE for three cultures or three mice per treatment. For analysis of mRNA expression levels, data were expressed relative to the mean of the same measure for WT controls. The level of significance was set to P < 0.05 and statistically significant differences from the control group at P < 0.05 and P < 0.01 are indicated by (*) and (**), respectively. Results are representative examples of at least three independent experiments.

Total RNA was isolated from cell cultures and gastrocnemius muscles (proximal-half part) using an RNeasy Micro kit (74,004; Qiagen, Hilden, Germany) and Trizol reagent (Thermo Fisher Scientific), respectively, according to the manufacturer’s recommendations. cDNA was synthesized from total RNA by a reversetranscriptase SuperScript III (180,808-44; Invitrogen) using oligo(dT) primer. The expression of mRNA for myogenin (NCBI RefSeq accession no. NM 031189.2), MyoD (NM 010866.2), myf5 (NM 008656.5), MRF4 (NM 008657.2), MEF2C (NM 001170537.1), APOBEC2 (NM 009694.3), neonatal MyHC (NM 177369.3), and embryonic MyHC (NM 001099635.1) was monitored by realtime quantitative PCR using Roche LightCycler 1.5 (Mannheim, Germany) run under the TaqMan probe detection format standardized with hypoxanthine guanine phosphoribosyl transferase (HPRT; accession no. NM 013556.2). Primer sets were designed by ProbeFinder (version 2.35 for mice, Roche) with an intron-spanning assay for mouse myogenin (amplicon 94 nt), MyoD (75 nt), myf5 (67 nt), MRF4 (77 nt), MEF2C (135 nt), APOBEC2 (71 nt), neonatal MyHC (63 nt), embryonic MyHC (69 nt), and HPRT (90 nt) as shown in Table 1. Annealing temperature was set to 60 ◦ C in all cases.

Gene

Forward (5 –3 )

Reverse (5 –3 )

APOBEC2 myogenin MyoD Myf5 MRF4 embryonic MyHC neonatal MyHC HPRT

ggagaagttggcagacatcc ccttgctcagctccctca cccctacactgtatgctgagg acaatgggacattccaggag gcacgccagtgcttcttc ttttcggccaaagtgaaca gaggagagggcggacatt tcctcctcagaccgctttt

tctgagtggcagcaggtaaa tgggagttgcattcactgg agaaaggagggcatgaatcc aaagactggcgctgctca catgctgctgtctgaaggtc cacccgcattgacgttct actcttcattcaggcccttg cctggttcatcatcgctaatc

Table 2 Stealth siRNA strands for mouse APOBEC2. siRNA

Sense (5 –3 )

Antisense (5 –3 )

Stealth 427 Stealth 738 Stealth 765

aaagcuggcaggaugguguuaaaga aguauaggaaguucuccugaauguc gaggagaaguuggcagacauccuga

ucuuuaacaccauccugccagcuuu gacauucaggaagaacuuccuauacu ucaggaugucugccaacuucuccuc

2.8. Statistical analyses

3. Results 3.1. Myoblast differentiation and fusion accelerated by A2KO The purpose of this study was to clarify a possible role of APOBEC2 in myoblast differentiation; therefore experiments began by examining the effect of A2KO on myogenic differentiation and fusion of myoblasts in primary culture (over 95% Pax7positive at 48 h post-plating). Myoblasts were isolated from wild type (WT) and A2KO mice and monitored for the activity of myotube formation in differentiation medium (DMEM-5% HS) by immunofluorescence staining with MF20 anti-MyHC at 0–8 days

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Fig. 1. Enhanced differentiation and fusion of A2KO myoblasts. Primary cultures of satellite cells from WT and A2KO mice (over 95% Pax7-positive at 48 h post-plating) were maintained in proliferation medium (DMEM-20% FBS) until 70% confluence and then transferred to differentiation medium (DMEM-5% HS) for the next 8 days and analyzed for fusion index and the level of differentiation marker proteins. (A) Representative phase-contrast images (upper panel) and immunofluorescence images (lower panel) for WT and A2KO cultures of anti-pan MyHC-positive cells (red, MF20-positive) counter-stained with DAPI (blue). Scale bars, 150 ␮m. (B) A bar graph of the fusion index measured at prescribed time-points after differentiation. Bars depict the mean ± SE for three cultures per group and significant differences from WT myoblast cultures (open bars) at the same time-point at P < 0.01 are indicated by (**). (C, D) Representative images of cell cultures from each group containing DAPI-positive and MyoD-positive nuclei (arrowheads indicate MyoD-negative/DAPI-positive cell nuclei) (C) and a bar graph of total cell number (per mm2 , DAPI-positive) and the percentage of MyoD-positive cells (about 80% positive in both cultures) (D) at Day-0 in WT and A2KO cultures, just prior to switching to differentiation medium. Scale bars, 100 ␮m. NS, no significant difference. (E) Level of two differentiation-marker proteins in WT and A2KO cultures, pan MyHC (MF20-positive) and myogenin. Whole-cell lysates were analyzed at prescribed timepoints after differentiation by ECL-Western blotting, referenced to ␤-actin. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

post-differentiation. As shown in Fig. 1A (upper panel of phase contrast images), A2KO myoblasts formed myotubes with normal appearance. Notably, myotube formation appeared to progress faster in A2KO myoblast cultures than WT myoblast cultures, as revealed by the immunofluorescence detection of more myotubes in A2KO myoblast cultures at 2 days post-differentiation (Fig. 1A, lower panel of fluorescence images). Subsequent experiments showed that A2KO myoblasts had a higher fusion index (percentage of nuclei in MyHC-positive myotubes) than WT myoblasts at day-1 and day-2 post-differentiation, with no significant difference at the later time-points (Fig. 1B). These findings, and the observa-

tion that there was no significant difference in total cell number or the percentage of MyoD-positive cells between WT and A2KO myoblast cultures just before induction of differentiation at day0 (Fig. 1C, D), are interpreted to show that cultures had similar myogenicity until the time of differentiation. These results were supported by Western blotting analysis of proteins from wholecell lysates showing that the expression levels of total MyHC and myogenin were higher in A2KO myoblasts at 2 days and 2–8 days post-differentiation, respectively (Fig. 1E), consistent with greater up-regulation of mRNA expression of myogenin and MEF2C (a myogenin-cooperating factor) (Fig. 2A, C). Other muscle-specific

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B 1.8 MyoD

**

1000

WT A2KO

800 **

600 400 200 0

0

1

2

3

MyoD / HPRT (relative unit)

Myogenin / HPRT (relative unit)

A1200 myogenin

1.6

1.4 1.2 1.0 0.8 0.6 0.4 0.2 0

4

Days post-Differentiation 60 50

**

40 *

30 20 10 0

0

1

2

3

4

Days post-Differentiation

MRF4 / HPRT (relative unit)

E

25

MRF4

0

1

2

3

4

1

2

3

4

Days post-Differentiation

D 1.4 myf5

MEF2C

Myf5 / HPRT (relative unit)

MEF2C / HPRT (relative unit)

C

95

1.2 1.0 0.8 0.6 0.4 0.2 0

0

Days post-Differentiation

P=0.054

20 15 P=0.075

10 5 0

0

1

2

3

4

Days post-Differentiation Fig. 2. Expression levels of MRF-family and MEF2C transcripts in WT and A2KO myoblast cultures. WT and A2KO myoblasts were cultured as shown in Fig. 1 and assayed by real-time RT-qPCR standardized with HPRT (TaqMan Probe assay) for mRNA expression of myogenin (A), MyoD (B), MEF2C (C), and myf5 (D), as well as MRF4 (E) at prescribed time-points after differentiation. Bars depict the mean and SE for three cultures per group relative to the mean for Day-0 WT myoblasts; significant differences (P < 0.05 and P < 0.01) from the WT mean at the same time point (open bars) are indicated by (*) and (**), respectively.

bHLH transcription factors, MyoD and myf5, showed comparable mRNA expression between WT and A2KO myoblast cultures (Fig. 2B, D) and a trend toward lower MRF4 mRNA expression in A2KO cultures at all time-points (Fig. 1E, P < 0.1 at day-1, 2), hence served as the negative control groups in the RT-qPCR assays. In order to further clarify the role of APOBEC2 in differentiation/fusion of myoblasts, an additional in vitro experiment on WT myoblast cultures was conducted to knockdown APOBEC2 at the differentiation phase (about 95% of the knockdown efficiency; see Fig. 3A left column). Proliferating myoblasts were changed to differentiation medium (DMEM-5% HS) containing APOBEC2specific siRNA or control siRNA for 48 h and evaluated for myogenic differentiation-maker expression by real-time RT-qPCR and Western blotting (Fig. 3). Results clearly showed that the siRNAs prevented APOBEC2 expression and that during APOBEC2 knockdown, there was increased expression of myogenin and total MyHC (adult and two developmental MyHCs) in message and protein

levels (Fig. 3A–C). These observations are consistent with the accelerated myogenic phenotype observed in A2KO myoblast cultures, as shown in Figs. 1 and 2. Adult MyHC protein expression (detected with a mixture of anti-fast and anti-slow-type MyHC antibodies) was also elevated by APOBEC2 knockdown, demonstrating an earlier progression of myotube formation in the absence of APOBEC2 expression (Fig. 3B first row and Fig. 3C). Notably, the expression levels of APOBEC2 in WT myoblast cultures were remarkably upregulated 1–4 days post-differentiation, with a maximum at 1 day (about 15-fold, relative to the baseline at Day-0, just before differentiation) (Fig. 3D), consistent with the previous report of a drastic increase in APOBEC2 protein expression in response to serum starvation (Sato et al., 2010). These results indicate that APOBEC2 may function at the early-differentiation phase as a negative regulator of myoblast differentiation and the resulting myotube-formation.

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Fig. 3. WT myoblast differentiation was enhanced by in vitro APOBEC2 knockdown. WT myoblasts were cultured in proliferation medium (F10-20% FBS) for 24 h before transfection with control siRNA or a mixture of 3 APOBEC2-specific siRNAs (100 nM) in differentiation medium (DMEM-5% HS) followed by evaluation for differentiation marker-gene expression at Day-2 post-transfection/differentiation by RT-qPCR standardized with HPRT (TaqMan Probe assay) (A) and by ECL-Western blotting referenced to ␤-actin (B, C) (n = 3 wells per treatment; panel C, densitometry analysis of the blots normalized to ␤-actin). Bars depict the mean and SE expressed relative to the mean of control-siRNA cultures (open bars); significant differences from control at P < 0.05 and P < 0.01 are indicated by (*) and (**), respectively. (D) Time-course of APOBEC2 mRNA expression during differentiation in WT derived myoblast cultures; note that APOBEC2 expression increased drastically at Day-1 post-differentiation (P < 0.01), and gradually declined over 4 days while remaining significantly higher than at Day-0 before transfection/differentiation. Significant differences from Day-0 at P < 0.01 are indicated by (**).

3.2. In vivo demonstration The physiological significance of the in vitro findings was explored using in vivo muscle-injury experiments in which WT and A2KO mice were evaluated for the myogenic regenerative potential after cardiotoxin (CTX) injection of gastrocnemius muscle (Figs. 4–6). The CTX-injury model is used to ensure high reproducibility of muscle-damage specific to muscle, without affecting vascular or nervous tissue as reported for the crush-injury model (McIntosh et al., 1994; Anderson and Vargas, 2003; Tatsumi et al., 2009; Shono et al., 2013; Sakaguchi et al., 2014), as it synchronously affects the whole muscle and thus provides a qualitative advantage for simpler analysis of the progressive myoblast differentiation and myotube formation. To monitor the time-course of muscle damage and regeneration at 0, 4, 7, 14, and 21 days post-injury, cryo-sections were stained with hematoxylin and eosin or with anti-laminin and anti-developmental MyHCs (since embryonic and neonatal MyHCs are known as regenerating myofiber markers). As shown in Fig. 4, histological observations of gastrocnemius muscle sections (mid-belly portion) showed many mono-nucleated cells and centrally nucleated muscle fibers as well as remnant areas of disrupted mature muscle fibers at 4–14 days post-injury in WT and A2KO mice. Importantly, the regenerative response was earlier in A2KO mice, as revealed by the smaller extent of areas with remnant fibers and higher incidence of developmental MyHC-

positive myotubes at 7 days post-CTX. Observation of newly formed myotubes was supported by RT-qPCR results which showed that the mRNA expression of neonatal MyHC was significantly higher in A2KO mice at 7 days post-injury (Fig. 5B). There was also a consistently higher level of myogenin mRNA expression at all time-points examined (P < 0.01 at days 4–21) (Fig. 5C), similar to findings from A2KO myoblast cultures shown in Figs. 1E and 2A, C. These results indicate that the loss of APOBEC2 enhanced myoblast differentiation and fusion in vivo, although it is still not clear if early progression of myogenesis affects the outcome of muscle-regeneration. This important question was addressed by measuring myofiber cross-sectional area and minimal Feret diameter (Fig. 6). Results demonstrated that mean cross-sectional area was significantly lower in regenerating gastrocnemius muscle in A2KO than WT controls at day-14 after CTX injury (678.85 ± 83.84 vs. 1268.85 ± 85.85 ␮m2 (mean ± SE), respectively) and at day21 (1087.82 ± 66.08 vs. 1470.24 ± 111.09 ␮m2 , A2KO and WT, respectively) (Fig. 6A–C). The validity of the findings of reduced fiber cross-sectional area was confirmed by the lower minimal Feret diameter of myofibers in A2KO than WT mice at day-14 (21.69 ± 1.54 vs. 31.24 ± 0.29 ␮m (mean ± SE), A2KO and WT, respectively, P < 0.01) and the trend to be lower at day-21 (27.83 ± 1.37 vs. 32.50 ± 3.27 ␮m, P = 0.17) as shown in Fig. 6D–F. Before injury (day-0), fiber size was significantly smaller in A2KO (1223.81 ± 55.13 vs. 2173.21 ± 108.92 ␮m2 in cross-sectional area

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Fig. 4. Histochemical detection of the regenerative phenotype in CTX-injured muscles of WT and A2KO mice. Gastrocnemius muscle was injured by intramuscular injection of CTX and collected at prescribed time-points after injury. (A) Micrographs (cross-sections at mid-belly portion) represent the progression of muscle regeneration as seen by hematoxylin and eosin staining (top row, WT; bottom row, A2KO mice). Note that there were more myotubes in regenerating muscle of A2KO than WT mice at Day-4 and -7 (arrow heads). (B) The panel (top row, WT; bottom row, A2KO) shows micrographs from double-immunostaining with anti-developmental MyHC (red; marking regenerating myotubes) and anti-laminin (green; marking the basal lamina) and nuclei were counter-stained with DAPI (blue). Note that regenerating muscle in A2KO mice showed more myotubes positive for developmental MyHC at 7 days post-CTX compared to WT regenerating muscle. Scale bars, 100 ␮m. The middle portion of gastrocnemius muscle (exclusive of the medial and lateral heads), known as fast IIb fiber-abundant area, was observed. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

and 32.75 ± 1.37 vs. 44.32 ± 1.52 ␮m in minimal Feret diameter) in agreement with our previous finding that APOBEC2 deficiency leads to a diminished muscle mass (Sato et al., 2010). Together these results indicate that APOBEC2 expression may contribute to normal myofiber formation by delaying progression of myoblast differentiation and fusion upon muscle injury. 4. Discussion We previously showed that APOBEC2 protein is expressed predominantly in striated muscles and expression of APOBEC2 is drastically increased upon differentiation of mouse myoblast cultures (Sato et al., 2010). The present study further explored features of myogenic differentiation in vitro, and extended the focus to an examination of the role of APOBEC2 in in vivo regenerative myogenesis after CTX injury. Results demonstrated that in vitro APOBEC2 deficiency enhances early progression of myogenic differentiation and fusion of myoblasts along with earlier up-regulation of MyHC, myogenin and its co-operating factor, MEF2C. Importantly, similar results were observed in APOBEC2-knockdown experiments in vitro, to show that reduced APOBEC2 during differentiation increased expression of MyHC (total and adult MyHCs) and myogenin in WT myoblast cultures. Interestingly, Vonica et al. (2011) reported the contrary result that myogenin was up-regulated in C2C12 myoblast cultures lipofected with APOBEC2 siRNA in growth medium for 6 h and analyzed 2 days later in differentiation medium without siRNA. These results indicate that APOBEC2 functions not in the proliferation phase but early in differentiation as a negative regulator of myoblast differentiation and myotube formation (see a model shown in Fig. 7). The phase-dependent specificity of the impact of APOBEC2 deficiency is further supported by two additional results, as follows. 1) There was no difference in cell proliferation, as characterized by total cell number and the percentage

of MyoD-positive cells in WT vs. A2KO myoblast preparations after plating at the same cell density and assayed just before inducing differentiation (Day-0, Fig. 1C, D). 2) The levels of expression of MyoD and myf5 were not affected by A2KO throughout the period of differentiation (Fig. 2B, D). MyoD and myf5 are crucial MRF-family members up-regulated exclusively during myoblast proliferation and known to mediate cell commitment toward subsequent differentiation (Ishibashi et al., 2005; De Falco and De Luca, 2006; Gayraud-Morel et al., 2007; Ustanina et al., 2007). In contrast to the lack of change in MyoD and myf5 expression and up-regulation of myogenin, expression of MRF4 showed a trend to decrease at the early-differentiation phase in response to APOBEC2 deficiency (P = 0.075 and 0.054 at Day-1 and -2, respectively; see Fig. 2E). Considering that the duration of MRF4 expression is longer than for other MRF-family members during differentiation and mediates the shift to myotube maturation (Zhang et al., 1995), the impact of APOBEC2 deficiency can be interpreted as enhancing primary myotube formation and also may suppress myotube growth. This possible insight may be supported by fusion index measurements getting comparable levels between A2KO and WT myoblasts at Day-3, 4 in culture (Fig. 1B) and also by the prolonged significant decrease in mean myofiber cross-sectional area 14 and 21 days after CTX-injury in vivo (Fig. 6B, C). Discussion of APOBEC2 as a negative regulator of myoblast differentiation/fusion and the impact on myotube formation and growth in muscle regeneration requires context from understanding details of this member of the AID/APOBEC cytidine deaminase family of proteins that edit DNA and RNA (Liao et al., 1999). Recent studies report that APOBEC2 may participate in DNA demethylation through its cytidine deaminase activity (Rai et al., 2008); the biological functions are not yet clear because APOBEC2-demethylation targets are still not known. The importance of DNA methylation is well established; it is one of the key epigenetic modifications

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eMyHC/HPRT (relative unit)

A

eMyHC

70 60

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50 40 30 20 10 0

0

neoMyHC / HPRT (relative unit)

B 160

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C

4 7 14 21 Days after CTX Injection

neoMyHC **

140 120 100 80 60 40 20 0

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4 7 14 Days after CTX Injection

21

myogenin

40 35

**

30

**

25 20 15 10

**

5 0

0

**

4 7 14 21 Days after CTX Injection

Fig. 5. Regenerative phenotype revealed by enhanced expression of gene transcripts for neonatal MyHC and myogenin at different time points after muscle injury. CTXinjured gastrocnemius muscle (proximal half) was assayed by RT-qPCR standardized with HPRT (TaqMan Probe assay), for the level of expression of a developmental MyHC (panel A, embryonic MyHC (eMyHC); B, neonatal MyHC (neoMyHC)) and myogenin (C) at prescribed time-points after injury. Bars depict the mean and SE relative to the mean at Day-0 of expression in the WT group and significant differences (P < 0.01) between WT (open bars) and A2KO groups (closed bars) at same time point are indicated by (**).

and masks DNA promoter regions to suppress gene transcription. Hence, demethylation of these regions normally will allow gene expression to initiate specific biological events (Morgan et al., 2005; Dilworth and Blais, 2011). Regulation of this gene-modification event is important for proper cell differentiation, as revealed by previous iconic experiments to show that treatment of a DNA methylation inhibitor 5-azacytidine disturbs fibroblast differentiation into the muscle lineage and C2C12 cells into the smooth muscle lineage (Taylor and Jones, 1979; Lee and Kim, 2007). Notably, the level of methylation is drastically reduced by induction of differentiation in vitro (Jost et al., 2001), and some reports showed that reduction of DNA methylation are observed in various events

including muscle regeneration and exercise-dependent muscle remodeling (Acharyya et al., 2010; Palacios et al., 2010; Dilworth and Blais, 2011; Barrès et al., 2012). These reports indicate that DNA methylation/demethylation is a key event that regulates neonatal muscle development as well as adult muscle status. APOBEC2deficient mice showed various phenotypes including changes in the fast: slow fiber-type proportion and an atrophic/dystrophic appearance in muscle (Sato et al., 2010); therefore it is tempting to speculate that these phenotypes may be generated by the dysfunction of DNA demethylation of the genes responsible for these changes in myogenesis. An inhibitory transcription factor for myogenin gene expression would be a reasonable candidate for receiving the APOBEC2-promoted demethylation postulated here, because myogenin expression was drastically up-regulated by APOBEC2 deficiency. There is not yet evidence to address this particular idea and further study is necessary to elucidate roles of APOBEC2 in muscle regeneration. The physiological significance of the APOBEC2-promoting negative regulation of myoblast differentiation in muscle regeneration is also important for discussion in context of current findings. APOBEC2 deficiency leads to diminished muscle mass (measured at 15-week-old, see Ref. Sato et al., 2010) and mean myofiber cross-sectional area after muscle injury (observed here); these new results are evidence that APOBEC2 deficiency may disturb normal muscle growth and regeneration despite enhancing myoblast differentiation and fusion. The muscle-regeneration process is divided into three phases including destruction/inflammation, repair, and remodeling (Turner and Badylak, 2012). At the destruction/inflammation phase, damaged myofibers undergo necrosis and inflammatory cells such as M1 and M2 macrophages invade the site of injury to perform the important function of phagocytosing fiber debris and secreting a variety of cytokines (Arnold et al., 2007; Tidball and Villalta, 2010; Kharraz et al., 2013; Mounier et al., 2013). At the same time, activated satellite cells migrate within and to the site of injury and fuse with each other (particularly in the CTX model of injury) to form new myofibers during the repair phase. This is followed by the remodeling phase characterized by fiber maturation including reconstruction of the extracellular matrix (ECM) and time-coordinated progression of regenerative moto-neuritogenesis and angiogenesis. The present results indicate that loss of APOBEC2 may “fast-forward” or accelerate the repair phase by up-regulating myoblast differentiation and hence shorten the duration of the destruction/inflammation phase. This effect has potential to lead to insufficient recovery of muscle function along with reduced myofiber diameter after CTX injury. In fact, dysfunction of phagocytosis and cytokine secretion by inflammatory cells in this early phase is known to impair normal muscle regeneration. A recent study on human hepatocytes showed that APOBEC2 expression was induced by treatment with tumor necrotic factor-␣ (TNF-␣) which is secreted predominantly by activated macrophages during inflammation (Matsumoto et al., 2006; Chen et al., 2007), which would be consistent with the idea that APOBEC2 deficiency could impair functional recovery from CTX injury despite accelerating the timing of myoblast differentiation and fusion into myotubes in vivo and in vitro. Finally, in association with the DNA demethylation hypothesis mentioned earlier, it is worth reiterating that APOBEC2 is downregulated during denervation-induced muscle atrophy (Sato et al., 2009) and that APOBEC2 deficiency induces up-regulation of myogenin expression in myogenic regeneration. Recent reports showed that in response to muscle denervation, myogenin expression is drastically up-regulated and promotes an increase in the expression of atrogin-1 and MuRF-1, known as E3 ubiquitin ligases that directly promote ubiquitin proteolysis (Tang et al., 2009; Moresi et al., 2010; Macpherson et al., 2011). These results suggest that myogenin may be a key factor in regulating denervation-induced

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Fig. 6. Decrease in myofiber size of gastrocnemius by A2KO. Cryo-sections (cross-sections of med-belly portion) were stained with anti-laminin at Day-0 (without injury) and at 14, and 21 days post-CTX injury to measure the distribution of myofiber cross-sectional area (% of fibers) (A–C) and the minimal Feret diameter means (D-F) of regenerating gastrocnemius muscle in WT (open bars) and A2KO mice (closed bars). The middle portion of gastrocnemius muscle (exclusive of medial and lateral heads), known as fast IIb fiber-abundant area, was observed. Black and white arrows indicate the mean fiber cross-sectional area of myofibers in A2KO and WT groups, respectively. Significant differences between A2KO and WT muscles at P < 0.05 and P < 0.01 are indicated by (*) and (**), respectively.

muscle atrophy and hence that APOBEC2 deficiency would also lead to muscle atrophy due to the activation of myogenin-atrogin-1MuRF-1 signaling. This possibility may account for the observations that loss of APOBEC2 leads to diminished muscle mass and lower fiber area in context of an incomplete inflammatory phase of phagocytosis and cytokine-secretion, described previously. Interestingly, Jiang et al. (2013) observed that increases in atrogin-1 and MuRF-

1 expression stimulate myoblast differentiation by degrading an inhibitory factor of myoblast differentiation in normal muscle regeneration; this report and our new findings advance the hypothesis that APOBEC2 plays an essential role in balancing the two directions of myoblast fate in muscle regeneration: myoblast differentiation and apoptosis via the ubiquitin proteasome system.

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Fig. 7. A possible model for the role of APOBEC2 in regenerative myogenesis. APOBEC2 protein (cytidine deaminase) is expressed exclusively in the early-differentiation phase upon muscle injury and may function as a key negative regulator of differentiation and fusion by myoblasts (upper flow). Hence, APOBEC2 deficiency enhances early progression of myogenic differentiation and fusion of myoblasts along with earlier up-regulation of myosin, myogenin and its co-operating factor, MEF2C, resulting in the insufficient maturation of myofibers (lower flow). This possibility may account for the observations that loss of APOBEC2 leads to diminished muscle mass and lower fiber area/diameter in context of (i) an incomplete inflammatory phase of phagocytosis and cytokine-secretion, (ii) the activation of myogenin-atrogin-1-MuRF-1 signaling responsible for muscle atrophy (not depicted here), and (iii) the dysfunction of epigenetic DNA/RNA demethylation of the genes responsible for these phenotypes in myogenesis (although APOBEC2-demethylation targets are still not known). Additionally, it is tempting to speculate that APOBEC2 may be an important mediator in the “self-renewal” functions of satellite cells, namely in the re-establishment of quiescent status after activation and proliferation; it is possible that APOBEC2 regulates a competitive balance between two trajectories of proliferated myoblasts: a return to cell quiescence and their differentiation and fusion which results in myotube formation (see Fig. 1 in Ohtsubo et al., submitted to Data in Brief).

Most studies to date on APOBEC2 biology have been conducted predominantly using zebrafish (Etard et al., 2010; Powell et al., 2012, 2014), with very little literature on APOBEC2 functions in mammals (Mikl et al., 2005; Sato et al., 2009, 2010). Powell et al. (2012, 2014) reported that APOBEC2 interacts with 3 proteins, the ubiquitin-conjugating enzyme 9 (Ubc9); the topoisomerase I-binding, arginine/serine-rich, E3 ubiquitin protein ligase (Toporsa); and the POU class 6 homeobox 2 protein (Pou6f2), to regulate retina regeneration in zebrafish, independent of its cytidine deaminase activity. APOBEC2 also controls left-right axis determination through a TGF-␤-signaling pathway in Xenopus embryogenesis (Vonica et al., 2011). The present study showed that APOBEC2 may function as a negative regulator of differentiation and fusion by mouse myoblasts, and thus provides novel insight into the function of APOBEC2 in the early-differentiation phase, so important during muscle regeneration. We are now testing an additional hypothesis that APOBEC2 may be an important mediator in the “self-renewal” functions of satellite cells, namely in the re-establishment of quiescent status after activation and proliferation. Our preliminary experiments in mouse single-myofiber cultures prepared from A2KO mice demonstrated a decrease in the population of Pax7(+)MyoD(−) quiescent satellite cells along with an complementary increase in the numbers of Pax7(−)MyoD(+) early-differentiated myoblasts (under investigation in this study) and Pax7(−)MyoD(−) cells (see Fig. 1 in Ohtsubo et al., submitted to Data in Brief). The particular idea of a role for APOBEC2 in the self-renewal functions would extend our understanding of APOBEC2-driven negative-regulation of myoblast differentiation and fusion; it is possible that APOBEC2 regulates a competitive balance between two trajectories of proliferated myoblasts during muscle regeneration: a return to cell quiescence which would reestablish the satellite cell pool and their differentiation and fusion which results in myotube formation. Further study is necessary to clarify the molecular basis of APOBEC2 functions and the impact on muscle regeneration and ultimately the physiology of regenerated muscle.

Acknowledgements The authors are grateful to the late Dr. Michael S. Neuberger (Medical Research Council Laboratory of Molecular Biology, United Kingdom) for the generous gift of APOBEC2-KO mice and to Dr. Koichi Ojima (NARO Institute of Livestock and Grassland Science, Japan) for the generous gift of mouse satellite cell-derived myoblasts (used for Fig. 3). The authors also acknowledge to Dr. Judy E. Anderson (University of Manitoba, Canada) for critical review of this manuscript. Special thanks to Ms. Akiko Sato, Mr. Shuichi Kitaura, and Mr. Junpei Goto (Kyushu University) for animal care. The mouse anti-Pax7 monoclonal antibody developed by D. A. Kawakami was obtained from the Developmental Studies Hybridoma Bank, created by the NICHD of the NIH and maintained at The University of Iowa, Department of Biology, Iowa City, IA 52242. This work was funded by Grant-in-Aid for Scientific Research (B) 23380159 from the Japan Society for the Promotion of Science (JSPS) (to Y. Ikeuchi). Research was also supported, in part, by Grants-in-Aid for Scientific Research (A) 16H02585 and (B) 22380145 and 25292164, by the Invitation Fellowship Program for Research in Japan (JSPS), and by grant funds from the Ito Foundation and Graduate School of Agriculture, Kyushu University (all to R. Tatsumi). H. Ohtsubo received a scholarship from Kyushu University during the course of this research.

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