BLU-554 | Ubiquitin Signal

LRTM1 promotes the differentiation of myoblast cells by negatively regulating the FGFR1 signaling pathway

Hao-ke Li a,1, Yong Zhou a,1, Jian Ding b, Lei Xiong a, Ying-xu Shi a, Yan-ji He a, Dan Yang a, Zhong-liang Deng a, Mao Nie a,*, Yan fei Gao a,**

Abstract

The proliferation and differentiation of myoblast cells are regulated by the fibroblast growth factor receptor (FGFR) signaling pathway. Although the regulation of FGFR signaling cascades has been widely investigated, the inhibitory mechanism that particularly function in skeletal muscle myogenesis remains obscure. In this study, we determined that LRTM1, an inhibitory regulator of the FGFR signaling pathway, negatively modulates the activation of ERK and promotes the differentiation of myoblast cells. LRTM1 is dynamically expressed during myoblast differentiation and skeletal muscle regeneration after injury. In mouse myoblast C2C12 cells, knockout (KO) of Lrtm1 significantly prevents the differentiation of myoblast cells; this effect is associated with the reduction of MyoD transcriptional activity and the overactivation of ERK kinase. Notably, further studies demonstrated that LRTM1 associates with p52Shc and inhibits the recruitment of p52Shc to FGFR1. Taken together, our findings identify a novel negative regulator of FGFR1, which plays an important role in regulating the differentiation of myoblast cells.

Keywords:

Myogenesis

Skeletal muscle

LRTM1

FGFR1

1. Introduction

The myofiber is the basic unit of skeletal muscle. Satellite cells are precursors to skeletal muscle cells [1–3]. Normally, satellite cells are in a quiescent state. Upon damage or disease, these quiescent stem cells become activated. After proliferating, satellite cells migrate to the injury sites and differentiate to fuse with damaged myofibers or to form new myofibers. This process is regulated by complicated mechanisms involving a variety of transcriptional and epigenetic regulators and extracellular signal molecules [4–7]. Transcriptional factors involved in myogenesis regulation are mainly from myogenic regulatory factors (MRFs), including myogenic factor 5 (Myf5), muscle-specific regulatory factor 4 (MRF4), myoblast determination protein (MyoD), and myogenin [5,8]. These transcription factors initiate and facilitate muscle cell differentiation by activating the transcription level of downstream genes.

Several extracellular signal pathways have been proved to participate in the myogenesis of myoblast cells [6]. Fibroblast growth factors (FGFs) are essential for self-renewal of satellite cells and necessary for the maintenance and regeneration of skeletal muscle [9]. As a member of the receptor tyrosine kinase (RTK) family, FGFRs regulate satellite cells mainly by activating the intracellular signaling cascades, including ERK, p38, PI3 kinase, PLCγ, and STATs [10]. Blocking the FGFRs (mainly FGFR1 and FGFR4) signaling pathway by adding an FGFR inhibitor promotes terminal differentiation of satellite cells [11]. FGF-stimulated ERK activation is insufficient for satellite cell expansion and does not repress myogenesis, but is required for the proliferation of satellite cells [12]. ERK activation promotes the G1-to-S phase transition by increasing the expression of Cyclin D and reduces p21 expression in satellite cells [12,13]. Although the biological importance of the FGFR signaling pathway has been revealed, the inhibitory mechanisms of FGFR signaling cascades have been poorly estimated. Some studies have revealed inhibitory molecules of FGFRs, including CBL [14], which is an E3 ubiquitin ligase of FGFR and FRS2, and Sprouty proteins [15–17], which inhibit the recruitment of the Grb2–Sos complex to FRS2 or Shp2. However, the inhibitory mechanisms of the FGFR signaling pathway remain to be established.

Here, we identified a novel inhibitory regulator of the FGFR signaling pathway, namely leucine-rich repeats and transmembrane domains 1 (LRTM1). We found that LRTM1 was dynamically expressed during myoblast differentiation and skeletal muscle regeneration after injury. In mouse myoblast C2C12 cells, KO of Lrtm1 significantly prevents the differentiation of myoblast cells. This effect is associated with the reduction of MyoD transcriptional activity and the overactivation of ERK kinase. Notably, further studies demonstrated that LRTM1 associates with p52Shc and inhibits the recruitment of p52Shc to FGFR1. Taken together, our findings identify a novel negative regulator of the FGFR signaling pathway, which plays an important role in regulating the differentiation of myoblast cells.

2. Results

2.1. LRTM1 is upregulated during myogenic differentiation and skeletal muscle regeneration after injury

C2C12 myoblasts, the in vitro myogenesis model, were induced to undergo myogenic differentiation by switching growth medium to low- serum medium (DM) at different time points, and RNA-seq analyses were then conducted. According to the transcriptome assembly, there were totally 22876 annotated transcripts in our mRNA-seq data. The differential expression was calculated, and the significance of each gene was analyzed between 1 day vs 3 day and 1 day vs 5 day. As shown in Fig. 1A and B, in 1 day vs 3 day group, the 5814 significantly dysregulated genes with FC more than 2 and p value less than 0.05, including 2992 upregulated and 2822 downregulated genes, were identified. In the 1 day vs 5 day group, 3833 upregulated and 3733 downregulated genes were identified. In these dysregulated genes, we intended to focus on the driving genes that might be altered at the early stage of myogenic differentiation. Among the upregulated genes in 1 day vs 3 day group (Fig. 1B and Table S1), an unknown function gene, LRTM1 was selected for further investigation. As shown in Fig. 1C and Fig. S1A, RT-PCR analysis was used to verify the RNA-seq data. The Western blot (WB) analysis was used to confirm the upregulation of LRTM1 during differentiation in terms of protein levels (Fig. 1D and Fig. S1A). Interestingly, in the well-established cardiotoxin (CTX)-induced muscle injury model [18,19], we found elevated levels of LRTM1 mRNA in the muscle tissue three days after CTX injection (Fig. 1E and Fig. S1B), which implies that LRTM1 participated in the skeletal muscle regeneration after injury.

2.2. LRTM1 is required for the myogenesis of C2C12 myoblasts

To assess the biological function of LRTM1 in myogenic differentiation, we first attempted to genetically silence LRTM1 expression in C2C12 myoblasts. The CRISPR/Cas9 system was used to generate the stable Lrtm1 KO subclones (Lrtm1 KO#1 and KO#2). The DNA sequencing data showed that Lrtm1 had been knocked out successfully (Fig. S1C). WB analysis was used to verify the Lrtm1 KO efficiency (Fig. 2A). First, the cell growth of these subclones in growth medium was evaluated. As shown in Fig. 2B, the cell growth of LRTM1 KO cells was markedly increased in growth medium. This result indicated that LRTM1 deletion enhanced the proliferation of myoblast cells. The LRTM1 KO C2C12 subclones were then induced to differentiate according to previously indicated protocols. Fig. 2C showed that myogenic differentiation in LRTM1-KO cells was substantially altered, as indicated by a decrease in the expression of the late myogenic marker, myosin heavy chain (MHC) [20]. As shown in Fig. 2D, the fusion index dramatically declined from 30% of control cells to 5% of LRTM1-KO cells (P < 0.05). The protein levels of early and late myogenic markers myogenin and MHC were detected by WB at different time points. In Fig. 2E and Fig. S2A, the protein levels of myogenin and MHC decreased dramatically at the corresponding time points in LRTM1-KO cells, whereas the protein levels of MyoD increased slightly. To confirm that the decrease in protein levels of MHC and myogenin was due to transcription repression, we determined the mRNA levels of MHC and myogenin. As shown in Fig. 2F, the mRNA levels of MHC and myogenin in LRTM1-KO cells reduced markedly as compared to that in the same stage of control cells. It has been proven that the transcriptional expression of both MHC and myogenin is regulated by MyoD [21,22]. To validate whether the transcriptional activity of MyoD was suppressed generally in LRTM1-KO cells, we checked several other MyoD downstream genes, which had MyoD direct binding sites in their promoter or enhancer region [23]. As shown in Fig. 2G, the expression of all indicated genes reduced significantly in LRTM1-KO cells, which implied that the transcriptional activity of MyoD was suppressed generally in LRTM1-KO cells.

2.3. LRTM1 deletion leads to overactivation of ERK

To investigate the underlying molecular mechanism, we checked the expression of several molecules that had been proven to regulate the transcriptional activity of MyoD [24,25]. As shown in Fig. 3A, the protein levels of Cyclin D1 and its partner CDK4, which binds directly to MyoD and inhibits its transcriptional activity, had increased in LRTM1 KO cells. Fig. 3B shows that the interaction between MyoD and CDK4 increased markedly. This result indicated that the repression of the transcriptional activity of MyoD is mainly due to the upregulation of Cyclin D1 and CDK4. Several upstream signal regulators that could modulate the expression of CDK4 and Cyclin D1 were assessed [26,27]. The activity of ERK, p38, and AKT was detected. Fig. 3C shows that the phosphorylation levels of ERK increased dramatically in LRTM1 KO (caption on next page) Fig. 3. LRTM1 deletion represses MyoD transcriptional activity by overactivating E RK MAPK. (A) WT and LRTM1− /− #1 C2C12 cells were cultured in differentiation medium (DM) for 3 and 4 days, respectively, and the levels of the indicated proteins were examined by WB. The protein levels of Cyclin D1 and CDK4 from experiment in left panel were quantified. (B) Total lysate from WT and LRTM1− /− cells induced to undergo differentiation for 3 days were immunoprecipitated using MyoD antibodies. The expression of MyoD, Cyclin D1, and CDK4 was determined by WB analysis. (C,D) WT and LRTM1− /− C2C12 cells were cultured in differentiation medium (DM) for 3 and 4 days, respectively. (C) The indicated phosphorylation levels and the total protein levels were assessed by WB (left). Relative phosphorylation levels of p38, AKT, and ERK were normalized to the levels of their total proteins (right). Data represent mean ± SD (n = 3, *p < 0.05). (D) The phosphorylation levels of cRaf, BRaf, ERK, and MEK were determined by WB. (E) WT or LRTM1− /− C2C12 cells were treated with 10 μM U0126 in DM for 3 days. Immunofluorescence analyses were performed using the antibody against MF20 (green)- and DAPI (blue)-stained nuclei. Scale bar: 200 μm. (F)The quantitative analysis result of the nuclei number in fibers as in (E). Data represent mean ± SD from at least three independent experiments (***p < 0.001). (G) Under the same condition as that in (E), and the levels of the indicated proteins were assessed by WB.

C2C12 cells. Under the same conditions, the phosphorylation levels of levels of Raf and MEK increased (Fig. 3D and Fig. S2B). These results p38 MAPK and AKT had marginal changes, according to the quantitative indicated that the ERK signaling pathway was overactivated in LRTM1 data (Fig. 3C, right panel). Further, we checked the upstream kinases of KO cells during myogenic differentiation. To verify the hypothesis that the ERK signaling pathway. In LRTM1 KO cells, the phosphorylation myogenesis repression in LRTM1-KO cells is due to ERK overactivation, we used a MEK kinase specific inhibitor, U0126, to inhibit the activity of ERK. As shown in Fig. 3E and F, the repression effect of LRTM1 deletion on myogenesis had been reversed by U0126 treatment. Following this, the protein levels of myogenin, MHC, Cyclin D1, and CDK4 were detected. As shown in Fig. 3G and Fig. S2C, in LRTM1-KO cells, after U0126 treatment, the protein levels of myogenin and MHC increased markedly, and the protein levels of Cyclin D1 and CDK4 decreased correspondingly. These results indicated that LRTM1 deletion leads to overactivation of ERK activity and, indirectly, down-regulated the transcriptional activity of MyoD.

2.4. LRTM1 inhibits FGFR1-induced ERK activation

Both the FGFR and IGFR signaling pathways contribute to the activation of ERK and regulate the myogenesis process of myoblast cells [6, 28]. To distinguish which receptor was responsible for the overactivation of ERK in LRTM1 KO C2C12 myoblasts, the FGFR-, and IGFR-specific inhibitors were used. As shown in Fig. 4A, treatment with the FGFR inhibitor significantly declined the activation of ERK in LRTM1 KO cells, whereas the IGFR inhibitors had no effect. To further investigate, the control and LRTM1-KO C2C12 cells were cultured in DM medium and treated with bFGF for 30 min, following which the activity of ERK was assessed. As shown in Fig. 4B, in LRTM1 KO cells, the activity of ERK was dramatically increased in response to bFGF stimulation. This result indicated that FGFR was responsible for the overactivation of ERK in LRTM1-KO cells. Under the same condition, the phosphorylation levels of FGFR1 signaling downstream molecules, SHP2 and p52Shc, were determined [29]. In response to bFGF stimulation, the phosphorylation level of p52Shc increased more significantly than that of SHP2. Consistent with this finding, we found that the phosphorylation level of p52Shc increased significantly in the myogenesis process in LRTM1-KO cells (Fig. 4C and Fig. S2D). To determine whether LRTM1 overexpression inhibits the activation of ERK, we established stable LRTM1-overexpressing C2C12 cells. Under the same condition as Fig. 4B, we found that LRTM1 overexpression declined the FGF-induced activation of ERK (Fig. 4D). Taken together, these results indicated that LRTM1 inhibited FGFR1-induced ERK activation.

2.5. LRTM1 inhibits the recruitment of p52Shc to FGFR1

To investigate how LRTM1 negatively regulated the FGFR signaling pathway, we first checked whether LRTM1 associates with FGFR1. As shown in Fig. 5A, we co-overexpressed FGFR1 and LRTM1 in HEK293T cells and found that LRTM1 was immunoprecipitated by FGFR1. Furthermore, the interaction between LRTM1 and several FGFR1 interaction proteins, including p52Shc, SHP2, Grab2, and FRS2, was assessed [29,30]. As shown in Fig. 5B, LRTM1 interacted with p52Shc but not with other proteins. As p52Shc is an adapter protein linking activated receptor tyrosine kinases to the Ras/ERK pathway [31], we decide to assess whether LRTM1 interferes with the interaction between FGFR1 and p52Shc. We immunoprecipitated FGFR1 to detect its binding with p52Shc, with or without the overexpression of LRTM1. As shown in Fig. 5C, the binding of p52Shc to FGFR1 reduced dramatically after LRTM1 overexpression, which implies that LRTM1 inhibited the recruitment of p52Shc to FGFR1.

3. Discussion

In this report, we found that LRTM1 was dynamically expressed during myoblast differentiation and skeletal muscle regeneration after injury. In mouse myoblast C2C12 cells, KO of Lrtm1 significantly prevented the differentiation of myoblast cells; this effect was associated with the reduction of MyoD transcriptional activity and the overactivation of ERK kinase. Notably, further studies demonstrated that LRTM1 associates with p52Shc and inhibits the recruitment of p52Shc to FGFR1 (Fig. 5D).

LRTM1 belongs to the extracellular LRRs superfamily and is located on chromosome 3p14.3 in human genome [32–35]. Like other family members, LRTM1 is also a transmembrane protein. In the extracellular region, it contains a signal peptide, LRR amino terminus, six LRRs, and an LRR carboxyl terminus, and in the intracellular region, it has a short tail with only 33 amino acids. This feature enables LRTM1 to participate in the extracellular signal transduction. In this study, we found that LRTM1 associated with another transmembrane receptor, FGFR1, and regulated its signaling transduction by inhibiting the recruitment of the adaptor protein p52Shc (Fig. 5). It has been reported that LRTM1 can be used as a cell surface marker to enrich midbrain dopaminergic (mDA) progenitors, as hiPSC-derived LRTM1-positive cells differentiated more efficiently into mDA neurons [32]. Hence, it is rational to hypothesize that the function of LRTM1 in the nervous system is related to its interaction with FGFR1. This possibility needs to be further investigated.

Previous data showed that LRTM1 mRNA was highly expressed in the brain but weakly expressed in the eye, lung, and heart of E11.5 fetal mouse [32] and was reduced in the diaphragms of Fgfrl1 homozygous null mice [35]. In our study, we found that LRTM1 was unregulated by transcriptional regulation in myoblasts and was involved in skeletal muscle regeneration after injury (Fig. 1C and E). Thus, there should be a positive feedback between LRTM1 and MyoD. According to the result that the transcription level of LRTM1 was elevated in the myogenesis process, LRTM1 may be a MyoD directly activated gene. We assessed several MyoD-Chip datasets and found a MyoD-occupied site at the promoter region of LRTM1 (data not show). The expression of LRTM1 declined FGFR1-induced ERK activation and subsequently reduced the level of CDK4, an inhibitor of MyoD transcriptional activity. This positive feedback facilitates the myogenesis of myoblast cells.

According to the published data, several inhibitory mechanisms of the FGFR signaling pathway have been revealed [10]. CBL, a E3 ubiquitin ligase, promotes the ubiquitination and degradation of FGFR and FRS214. A high expression level of GRB2 interferes with PLCγ binding to FGFR [36]. Sprouty-1 and Sprouty-2 interact with GRB2 to inhibit the recruitment of the Grb2–Sos complex to FRS2 or Shp215. In the skeletal muscle system, the biological function of Sprouty proteins has been estimated [16,17]. It has been shown that Sprouty-1 is required for maintaining the satellite cell pool during repair and that artificial overexpression of Sprouty-2 promoted the myogenesis of C2C12 cells. These findings indicated that the biological function of LRTM1 was similar to that of Sprouty proteins, which promote the myogenic differentiation of myoblast cells by inhibiting the activity of the FGFR1 signaling pathway. Because the binding proteins were different, as shown in Fig. 5B, we examined the interaction of LRTM1 with Grb2 but found no such interaction; instead, we found a strong relationship between LRTM1 and p52Shc, an adaptor protein for FGFR1 signaling transduction. Further studies are needed to understand the precise molecular mechanisms and the crosstalk between Sprouty proteins with LRTM1.

Taken together, our findings reveal a novel negative regulator of FGFR1, which plays an important role in regulating the differentiation of myoblast cells.

4. Materials and methods

4.1. Cell culture, cell lines, and treatment

C2C12 mouse myoblasts were purchased from ATCC (Manassas, VA, USA) and cultured in growth medium (GM)—DMEM containing 10% fetal bovine serum (FBS) and maintained in a humidified incubator with 5% CO2 at 37 ◦C. For myogenic differentiation, when the confluence of cells reached 80%, C2C12 cells were transferred to a differentiation medium (DM)—DMEM containing 2% horse serum (HS). 293T cells were cultured in DMEM containing 10% FBS and maintained in a humidified incubator with 5% CO2 at 37 ◦C. C2C12 cells were treated with the MEK1/2 specific inhibitor U0126 (10 μM; S1102, Selleck) or the IGF- 1 and FGFR1/2/3 specific inhibitors Linsitinib (1 μM; HY-10191, MCE) and BGJ398 (100 nM; M1840, AbMole), respectively.

4.2. Cardiotoxin injury

These assays were carried out as previously described [18]. Cardiotoxin from Naja Mossambica mossambica (SigmaAldrich, USA) was dissolved to a final concentration of 10 μM 50 μl of cardiotoxin were injected with a 27 Gauge needle into one TA muscle; the other muscle was injected with saline as control.

4.3. Transfection and plasmids

For plasmid transfection, when the cells reached 60–70% confluence, the plasmids were transfected using the transfection reagent FuGene (Promega, Spain, cat. #: E231A) according to the manufacturer’s protocol. The LRTM1-FLAG and FGFR1-FLAG plasmids were purchased from FulenGen (Guangzhou, China), and LRTM1-Myc, FRS2- Myc, SHP2-Myc, and Grb2-Myc plasmids were purchased from Origene (China).

4.4. RNA extraction and real-time quantitative PCR

Total RNA was isolated with Trizol reagent (Invitrogen, Waltham, MA, USA) according to the manufacturer’s protocol. A total of 700 ng of RNA was reverse-transcribed using the PrimeScript™ RT reagent kit (TaKaRa, Tokyo, Japan). The SYBR green (TaKaRa) method was used with the Realplex real-time PCR detection system (Eppendorf) to detect gene expression. Real-time PCR was performed in triplicate. The sequences of the primers were as follows: mGAPDH (Forward, 5′- AGTGTTTCCTCGTCCCGTAG-3′, Reverse, 5′-GCCGTGAGTGGAGTCATACT-3′); mLRTM1 (Forward, 5′-TGTTGAATGAGGGTTTGTGCT-3′, Reverse, 5′-TCCACGGAGTTTGATGATGG-3′); mMyosin (Forward, 5′- ACAAGCTGCGGGTGAAGAGC-3′, Reverse, 5′-CAGGACAGTGACAAAGAACG-3′); mMyoG (Forward, 5′-CTGACCCTACAGACGCCCAC-3′, Reverse, 5′-TGTCCACGATGGACGTAAGG-3′). GAPDH was used as the normalization control. The data were analyzed using the comparative Ct (2–△△Ct) method.

4.5. mRNA-seq and analysis

WT and LRTM1− /− C2C12 cells were cultured in differentiation medium for 0, 1, 3, and 5 days. Total RNA was extracted from C2C12 cells using TRIzol (Invitrogen). A total amount of 3 μg RNA per sample was used as the input material for RNA sample preparation. Sequencing libraries were generated using the NEBNext® Ultra™ RNA Library Prep Kit for Illumina® (NEB, USA) following the manufacturer’s recommendations, and index codes were added to attribute sequences to each sample. mRNA was purified from total RNA using poly-T oligo-attached magnetic beads. Fragmentation was performed using divalent cations under elevated temperature in NEBNext First Strand Synthesis Reaction Buffer (5X). First strand cDNA was synthesized using a random hexamer primer and M-MuLV Reverse Transcriptase (RNase H-). Second strand cDNA synthesis was subsequently performed using DNA Polymerase I and RNase H. The remaining overhangs were converted into blunt ends by exonuclease/polymerase activities. After adenylation of. 3ʹ ends of DNA fragments, an NEBNext Adaptor with hairpin loop structure was ligated to prepare for hybridization. To select cDNA fragments of preferentially 250–300 bp in length, the library fragments were purified with AMPure XP system (Beckman Coulter, Beverly, USA). Then, 3 μl USER Enzyme (NEB, USA) was used with size-selected, adaptor-ligated cDNA at 37 ◦C for 15 min followed by 5 min at 95 ◦C before PCR. The PCR products were then purified (AMPure XP system), and the library quality was assessed on the Agilent Bioanalyzer 2100 system. After cluster generation, the library preparations were sequenced on an Illumina Hiseq platform, and 125 bp/150 bp paired- end reads were generated. Gene Ontology (GO) enrichment analysis of differentially expressed genes was implemented by the clusterProfiler R package, and hierarchical clustering heatmap was made with the ggplot library.

4.6. Immunoprecipitation and immunoblotting

Cells were lysed in E1A buffer (250 mM NaCl, 0.2% NP-40, 1 mM EDTA, 50 mM Tris-HCl, pH 7.6) supplemented with protease inhibitors (Cocktails, Roche) and 1 mM DTT; the lysates were incubated with 4 μg antibody for each group overnight at 4 ◦C, followed by 4 h incubation with agarose beads. Immunocomplexes were washed five times with E1A buffer before being resolved by SDS-PAGE for WB.

For WB, the cells were lysed in RIPA buffer. Protein samples were separated by SDS-PAGE and transferred to PVDF membranes. The membranes were then blocked with 5% milk in TBST and probed with primary antibodies, including β-tubulin (sc-166729, Santa Cruz), FLAG (F1804, Sigma), Shc (12496-1-AP; Proteintech), MYC (16286-1-AP; Proteintech), HSP90 (TA500494, ORIGENE), FGFR1 (9740, Cell Signaling Technology), AKT (2920, Cell Signaling Technology), p-AKT Thr308 (4056, Cell Signaling Technology), p-Shc Tyr239/240 (2434, Cell Signaling Technology), p-ERK1/2 Thr202/Tyr204 (9101; Cell Signaling Technology), p-MEK Ser217/221 (9154, Cell Signaling Technology), p-c-Raf Ser338 (9427; Cell Signaling Technology), p-b-Raf Ser445 (2696; Cell Signaling Technology), LRTM1 (sc-139390, Santa Cruz), myogenin (sc-12732, Santa Cruz), MF20 (14-6503-80; Invitrogen), MyoD (sc-377460, Santa Cruz), CDK4 (sc-23896; Santa Cruz), ERK1/2(220003, ZEN BIO), p-p38 Thr180/Tyr182 (310091, ZEN BIO), p38 (340697; ZEN BIO), and Cyclin D1 (380999; ZEN BIO).

4.7. Immunofluorescence staining

C2C12 cells were cultured on glass coverslips and induced to differentiate for 3 days. The cells were then fixed with 4% paraformaldehyde for 15 min, permeabilized with 0.2% Triton X-100 for 10 min, blocked with 5% goat serum solution, incubated with primary antibodies (for Myosin: 1:100; Invitrogen) at 4 ◦C overnight, incubated with secondary antibodies at room temperature for 1 h, and stained with DAPI (1:1000). Immunofluorescence images were captured by microscopic examination (EVOS FL Auto Cell Imaging System, Life Technologies, USA).

4.8. Generation of LRTM1 KO C2C12 cell line by CRISPR-Cas9

pCRISPR-LvSG06 plasmids encoding sgRNAs against LRTM1 (gRNA sequence A: GCCGGGATATATGTTGAATG; sequence B: CCACCCATCATCAAACTCCG; sequence C: GTCCATGGGGGTAAATCACG) were purchased from FulenGen. Lentivirus from 293T cells containing the CRISPR-Cas9 system was used to infect C2C12 cells. Stable cell lines were selected using puromycin (2 μg/ml). Single cells were selected in 96-well plates. To validate gene editing, target regions were identified with PCR amplification and sequenced and also validated by immunoblotting.

4.9. Statistical analysis

ImageJ software was used to measure the relative intensity of each band for quantification of the WB data. GraphPad Prism 7 was used for data analysis. Statistical comparisons between the two groups were performed using Student’s t-test. Data are expressed as mean ± standard deviation (s.d.) from at least three independent experiments. p < 0.05 was considered statistically significant. *p < 0.05, **p < 0.01, ***p < 0.001.

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[27]W. Jiang, J. Zhu, X. Zhuang, X. Zhang, T. Luo, K.A. Esser, et al., Lipin1 regulates skeletal muscle differentiation through extracellular signal-regulated kinase (ERK) activation and cyclin D complex-regulated cell cycle withdrawal, J. Biol. Chem. 290 (39) (2015) 23646–23655.

[28]D.M. Ornitz, P.J. Marie, Fibroblast growth factor signaling in skeletal development and disease, Genes Dev. 29 (14) (2015) 1463–1486.

[29]A.C. Schuller, Z. Ahmed, J.A. Levitt, K.M. Suen, K. Suhling, J.E. Ladbury, Indirect recruitment of the signalling adaptor Shc to the fibroblast growth factor receptor 2 (FGFR2), Biochem. J. 416 (2) (2008) 189–199.

[30]N. Turner, R. Grose, Fibroblast growth factor signalling: from development to cancer, Nat. Rev. Canc. 10 (2) (2010) 116–129.

[31]E. Migliaccio, S. Mele, A.E. Salcini, G. Pelicci, K.M. Lai, G. Superti-Furga, et al., Opposite effects of the p52shc/p46shc and p66shc splicing isoforms on the EGF receptor-MAP kinase-fos signalling pathway, EMBO J. 16 (4) (1997) 706–716.

[32]B. Samata, D. Doi, K. Nishimura, T. Kikuchi, A. Watanabe, Y. Sakamoto, et al., Purification of functional human ES and iPSC-derived midbrain dopaminergic progenitors using LRTM1, Nat. Commun. 7 (2016) 13097.

[33]A. Herger, K. Dunser, J. Kleine-Vehn, C. Ringli, Leucine-rich repeat extensin proteins and their role in cell wall sensing, Curr. Biol. : CB 29 (17) (2019) R851–R858.

[34]J. de Wit, W. Hong, L. Luo, A. Ghosh, Role of leucine-rich repeat proteins in the development and function of neural circuits, Annu. Rev. Cell Dev. Biol. 27 (2011) 697–729.

[35]N. LopezJimenez, S. Gerber, V. Popovici, S. Mirza, K. Copren, L. Ta, et al., Examination of FGFRL1 as a candidate gene for diaphragmatic defects at chromosome 4p16.3 shows that Fgfrl1 null mice have reduced expression of Tpm3, sarcomere genes and Lrtm1 in the diaphragm, Hum. Genet. 127 (3) (2010) 325–336.

[36]C.C. Lin, F.A. Melo, R. Ghosh, K.M. Suen, L.J. Stagg, J. Kirkpatrick, et al., Inhibition of basal FGF receptor signaling by dimeric Grb2, Cell 149 (7) (2012) 1514–1524.

LRTM1 promotes the differentiation of myoblast cells by negatively regulating the FGFR1 signaling pathway

Hao-ke Li a,1, Yong Zhou a,1, Jian Ding b, Lei Xiong a, Ying-xu Shi a, Yan-ji He a, Dan Yang a, Zhong-liang Deng a, Mao Nie a,*, Yan fei Gao a,**

Abstract

Keywords:

Myogenesis

Skeletal muscle

LRTM1

FGFR1

1. Introduction

2. Results

2.1. LRTM1 is upregulated during myogenic differentiation and skeletal muscle regeneration after injury

2.2. LRTM1 is required for the myogenesis of C2C12 myoblasts

2.3. LRTM1 deletion leads to overactivation of ERK

2.4. LRTM1 inhibits FGFR1-induced ERK activation

2.5. LRTM1 inhibits the recruitment of p52Shc to FGFR1

3. Discussion

Taken together, our findings reveal a novel negative regulator of FGFR1, which plays an important role in regulating the differentiation of myoblast cells.

4. Materials and methods

4.1. Cell culture, cell lines, and treatment

4.2. Cardiotoxin injury

4.3. Transfection and plasmids

4.4. RNA extraction and real-time quantitative PCR

4.5. mRNA-seq and analysis

4.6. Immunoprecipitation and immunoblotting

4.7. Immunofluorescence staining

4.8. Generation of LRTM1 KO C2C12 cell line by CRISPR-Cas9

4.9. Statistical analysis

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[28] D.M. Ornitz, P.J. Marie, Fibroblast growth factor signaling in skeletal development and disease, Genes Dev. 29 (14) (2015) 1463–1486.

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[31] E. Migliaccio, S. Mele, A.E. Salcini, G. Pelicci, K.M. Lai, G. Superti-Furga, et al., Opposite effects of the p52shc/p46shc and p66shc splicing isoforms on the EGF receptor-MAP kinase-fos signalling pathway, EMBO J. 16 (4) (1997) 706–716.

[32] B. Samata, D. Doi, K. Nishimura, T. Kikuchi, A. Watanabe, Y. Sakamoto, et al., Purification of functional human ES and iPSC-derived midbrain dopaminergic progenitors using LRTM1, Nat. Commun. 7 (2016) 13097.

[33] A. Herger, K. Dunser, J. Kleine-Vehn, C. Ringli, Leucine-rich repeat extensin proteins and their role in cell wall sensing, Curr. Biol. : CB 29 (17) (2019) R851–R858.

[34] J. de Wit, W. Hong, L. Luo, A. Ghosh, Role of leucine-rich repeat proteins in the development and function of neural circuits, Annu. Rev. Cell Dev. Biol. 27 (2011) 697–729.

[35] N. LopezJimenez, S. Gerber, V. Popovici, S. Mirza, K. Copren, L. Ta, et al., Examination of FGFRL1 as a candidate gene for diaphragmatic defects at chromosome 4p16.3 shows that Fgfrl1 null mice have reduced expression of Tpm3, sarcomere genes and Lrtm1 in the diaphragm, Hum. Genet. 127 (3) (2010) 325–336.

[36] C.C. Lin, F.A. Melo, R. Ghosh, K.M. Suen, L.J. Stagg, J. Kirkpatrick, et al., Inhibition of basal FGF receptor signaling by dimeric Grb2, Cell 149 (7) (2012) 1514–1524.