Epidermal growth factor receptor contributes to indirect regulation of skeletal muscle mass by androgen

Abstract

Androgen is widely acknowledged to regulate skeletal muscle mass. However, the specific mechanism driving muscle atrophy resulting from androgen deficiency remains elusive. Systemic androgen receptor knockout (ARKO) mice exhibit reduction in both muscle strength and muscle mass while skeletal muscle fiber specific ARKO mice have decreased muscle strength without affecting skeletal muscle mass in the limbs. Therefore, androgens may indirectly regulate skeletal muscle mass through effects on non-myofibers. Considering this, our investigation focused on blood fluid factors that might play a role in the regulation of skeletal muscle mass under the influence of androgens. Using a male mouse model of sham, orchidectomy and DHT replacement, mass spectrometry for serum samples of each group identified epidermal growth factor receptor (EGFR) as a candidate protein involving the regulation of skeletal muscle mass affected by androgens. Egfr expression in both liver and epididymal white adipose tissue correlated with androgen levels. Furthermore, Egfr expression in these tissues was predominantly elevated in male compared to female mice. Interestingly, male mice exhibited significantly elevated serum EGFR concentrations compared to their female counterparts, suggesting a connection with androgen levels. Treatment of EGFR to C2C12 cells promoted phosphorylation of AKT and its downstream S6K, and enhanced the protein synthesis in vitro. Furthermore, the administration of EGFR to female mice revealed a potential role in promoting an increase in skeletal muscle mass. These findings collectively enhance our understanding of the complex interplay among androgens, EGFR, and the regulation of skeletal muscle mass.

Introduction

In a super-aging population, extending healthy life expectancy is an urgent issue. Primary factors compromising healthy life expectancy are musculoskeletal diseases including sarcopenia, which refers to the age-related atrophy of skeletal muscle [1]. Sarcopenia serves as a crucial pathophysiological contributor to frailty in the elderly, characterized by diminished muscle strength, reduced muscle mass, and impaired physical function [2]. Several studies have reported that sarcopenia increases both mortality and risk of long-term care [3-5]. Sarcopenia is categorized into two types: primary sarcopenia, attributed to age-related muscle degradation, and secondary sarcopenia, arising from non-age-related factors [6]. Secondary sarcopenia has multifaceted etiologies, one of which includes the decline in androgen levels in men [6]. Androgens such as testosterone and its active form, dihydrotestosterone (DHT), bind to the androgen receptor (AR) to form a dimer that translocates into the nucleus, and then the dimer binds to the androgen response elements (AREs) to activate target genes, thereby manifesting androgenic effects [7, 8]. Androgens play a crucial role in skeletal muscle mass and strength, and the reduction in androgen levels is considered a cause of sarcopenia in elderly males [9]. Because prostate growth is androgen-dependent, androgen deprivation therapy (ADT) is a common treatment for prostate cancer [10]. Long-term ADT has been reported to decrease skeletal muscle mass [10, 11]. Androgen replacement increases skeletal muscle mass and strength, but has serious problems, including cardiovascular adverse events [12]. While the relationship between androgens and skeletal muscle is well recognized, the detailed molecular mechanism remains unknown.

Male systemic AR knockout (KO) mice exhibit reductions in both skeletal muscle mass and muscle strength [13]. On the other hand, skeletal muscle fiber-specific ARKO mice demonstrate reduced muscle strength without a significant impact on skeletal muscle mass [14]. Furthermore, DHT administration to female skeletal muscle fiber specific ARKO mice has been shown to increase skeletal muscle mass [15]. These findings indicate that muscle strength is directly regulated by androgen action on skeletal muscle fibers, while the regulation of skeletal muscle mass may occur indirectly through AR activity in cells or tissues other than skeletal muscle fibers.

We hypothesized that androgens could affect skeletal muscles via blood factors through the AR outside of muscle fibers. To test this hypothesis, we performed liquid chromatograph–tandem mass spectrometry (LC-MS/MS) of serum from castrated and androgen treated mice. We found that epidermal growth factor receptor (EGFR) is a candidate factor in serum affecting skeletal muscle. EGFR belongs to the HER family of membrane-bound receptors, has tyrosine kinase activity, and is involved in various intracellular signaling [16]. We investigated in detail the relationship between soluble EGFR (sEGFR) and androgen-mediated skeletal muscle regulation.

Materials and Methods

Animals

C57BL/6J male mice (CLEA Japan and Jackson Laboratory) were divided into three groups of 4–5 mice: sham surgery, androgen deprivation through orchidectomy (ORX), and dihydrotestosterone (DHT) replacement after ORX. All operations were performed under anesthesia. They received either vehicle (Veh) or DHT (Stanolone, TCI, Cat# A0462, 2.5 mg/mouse/week) subcutaneously every week starting at 10 weeks of age and were sampled for skeletal muscle and serum at 12 weeks of age. All three groups of mice were treated for a total of 4 and 8 weeks, and serum was collected at 14 and 18 weeks old. In the soluble EGFR (sEGFR) administration experiment, the DHT-treated group was used as a positive control. Female C57BL/6J mice were subcutaneously administered Veh, DHT (2.5 mg/mouse/week), or intraperitoneally administered sEGFR (SIN Sino Biological Inc, Cat# 51091-M08H, 20 μg/mouse/week) every week starting at 8 weeks old. Five mice were used per group. All mice were housed in a specific pathogen-free environment under controlled climatic conditions at room temperature, subject to a 12-hour light/dark cycle. They were given unrestricted access to water and a standard diet. Animal experiments received approval from the Animal Experiment Committee of Ehime University (Approval No. 37A1-1·16) and were conducted in strict accordance with the Guidelines for Animal Experiments at Ehime University.

Orchidectomy and sham surgery

Surgical procedures were carried out on 10-week-old male mice utilizing isoflurane as the anesthetic agent. Following skin disinfection, an incision was made to access the testicular region. For ORX, the testes were bilaterally excised, with blood vessels cauterized to mitigate bleeding. Conversely, for the control group, the testes were left intact and repositioned within the body cavity. The wound was sutured twice with nylon thread.

Adjustment of DHT

DHT (Stanolone, TCI, Cat# A0462) was dissolved in 99.5% ethanol (Wako, Cat# 057-00456) to prepare a 50 mg/mL DHT solution, then stored at –80°C. Immediately prior to use, the solution was further diluted ten times using 0.3% Hydroxypropyl Cellulose (HPC) (Wako, Cat# 085-07932). As a vehicle control, 99.5% ethanol was also diluted tenfold in 0.3% HPC.

Evaluation of grip strength

Forelimb grip strength in mice was assessed using a strain gauge (Melquest, Cat# GPM-100B). Each mouse underwent ten repeated measurements, and the maximum force was recorded. The mean value of the two highest records was utilized to determine the grip strength for each individual mouse.

Determination of serum EGFR

The concentration of serum EGFR was determined by ELISA with Quantikine kit for mice (RSD, Cat# MEGFR0) according to the manufacturer’s instructions.

Liquid chromatograph–tandem mass spectrometry (LC–MS/MS)

LC-MS/MS analyses were conducted by SHIMADZU. We provided three serum samples from each group of mice: sham surgery, androgen deprivation through ORX, and androgen replacement after ORX. Proteins recognized in the same set of identified peptides were grouped together as one group. Peptides with a false discovery rate (FDR) <0.01 were considered significant. Proteins with a 1.5-fold or greater increase or decrease in the sample-to-sample abundance ratio were selected.

RNA preparation, cDNA synthesis, and quantitative real time PCR

Total RNA from skeletal muscle and other non-adipose tissues was isolated using ISOGEN (NIPPON GENE, Cat# 319-90211). For adipose tissue and adipocytes, total RNA was extracted using QIAzol Lysis Reagent (QIAGEN, Cat# 79306). RNA samples were reverse-transcribed using PrimeScript RT Master Mix (Takara, Cat# RR036A), adhering to the manufacturer’s guidelines, with each reverse transcription employing 500 ng of total RNA. Quantitative reverse transcription PCR (RT-PCR) was conducted using SYBR Premix Ex Taq II (Takara, Cat# RR820L) in accordance with the manufacturer’s protocols. The PCR conditions consisted of 40 cycles at 95°C for 5 seconds, followed by 60°C for 30 seconds, using the Thermal Cycler Dice Real Time System III (Takara, Cat# TP950). Gene expression was normalized to the expression levels of the housekeeping gene, Rpl13a. All primer sequences are provided in Supplementary Table 1.

Adjustment of EGFR for the in vivo experiment

EGFR recombinant protein, consisting of the extracellular domain of mouse EGFR (Met 1-Ser 647) fused to a C-terminal polyhistidine tag (SIN Sino Biological Inc, Cat# 51091-M08H), or soluble form of EGFR, was used for in vivo treatment. The recombinant sEGFR protein was diluted with sterile water to 0.1 mg/mL. Sterile water was administered as a vehicle control.

Reanalysis of ChIP-seq and ChIP-qPCR

Previous ChIP-seq data for AR were searched by Cistrome Data Browser [17, 18]. Bigwig files from GSM 1907200 [19], GSM 1328954 [20] were displayed by Integrative Genomics Viewer (IGV) [21]. The GSM 1276800 [22] dataset was displayed by Cistrome Data Browser [17, 18]. LNCaP and 3T3-L1 cell lines were used for ChIP-qPCR analysis. LNCaP cells were incubated at 37°C in RPMI-1640 (Wako, Cat# 189-02025) with 10% FBS, 1% antibiotic mixture of Penicillin Streptomycin (Gibco, Cat# 15140-122), and Amphotericin B (Gibco, Cat# 15290-018) at 10:1. The medium was replaced with phenol red-free RPMI-1640 (Wako, Cat# 186-02155) with 10% charcoal-stripped FBS (Sigma-Aldrich, Cat# 173012) and 1% antibiotic mixture, allowed to stand overnight, then DHT of 10^–7 M was added and incubated for 12 hours. 3T3-L1 cells were incubated at 37°C in DMEM (Wako, Cat# 043-30085) with 10% FBS, 1% antibiotic mixture, and Amphotericin B (Gibco, Cat# 15290-018) at 10:1. The medium was replaced with phenol red-free DMEM (Gibco, Cat# 31053-028) with 10% charcoal-stripped FBS and 1% antibiotic mixture, allowed to stand overnight, then 10^–7 M DHT was added and incubated for 12 hours. Cell fixation and subsequent ChIP-qPCR were performed according to the protocol of ChIP-IT High Sensitivity kit (Active Motif, Cat# 53040).

Western blotting for Akt signaling

Differentiated C2C12 cells were cultured overnight in DMEM medium without FBS, and sEGFR was added at concentrations of 0, 0.5, 2, and 8 nM and incubated at 37°C for 5 minutes. Collected cells were sonicated for 15 min and centrifuged at 20,000 g for 20 min at 4°C. The upper clearings were collected, and lysed in RIPA buffer (Wako, Cat# 188-02453) containing a cocktail of proteases (Nacalai Tesque, Cat# 25955-11) and a cocktail of phosphatase inhibitor (Nacalai Tesque, Cat# 07574-61). The gastrocnemius muscles at 10 weeks of age treated with Veh, sEGFR or DHT were stirred in a tube with an iron ball in RIPA buffer with an added cocktail of proteases and a cocktail of phosphatase inhibitor. The solution was centrifuged at 16,000 g for 20 minutes at 4°C and the supernatant was collected. The concentration of the extracted proteins was measured by NanoDrop (ThermoFisher, Cat# ND-2000). The proteins were mixed with sodium dodecyl sulfate (SDS) sample buffer and boiled at 95°C for 5 min. Samples were applied to 12.5 % SuperSep Ace polyacrylamide gels (Wako, Cat# 196-14981) and conducted to SDS-polyacrylamide gel electrophoresis for 60 minutes followed by transferring to PVDF membranes (Bio-Rad Laboratories, Cat# 1620177) with a mini transblot cell (Bio-Rad Laboratories, Cat# 1703930JA) for 90 minutes. Membranes were blocked with 5% skim milk for 60 minutes at room temperature. Primary antibodies including anti-Akt1/2/3 (Santa Cruz Biotechnology, Cat# sc-81434; 1:200 dilution), anti-phospho-Akt (Cell Signaling, Cat# 4060; 1:2,000 dilution), anti-p70 S6 Kinase (Cell Signaling, Cat# 2708; 1:1,000 dilution), anti-phospho-p70 S6 Kinase (Cell Signaling, Cat# 9234; 1:1,000 dilution), anti-β-actin (MBL, Cat# M177-3; 1:5,000 dilution) for proteins from C2C12, and anti-GAPDH (Cell Signaling, Cat# 5174; 1:1,000 dilution) for gastrocnemius muscles were added and incubated overnight at 4°C. Following washing with TBS containing 0.05% Tween 20, the membranes were treated with secondary antibodies including horseradish peroxidase (HRP)-conjugated anti-rabbit IgG (Dako, Cat# P0448; 1:2,500 (pAkt, S6k, pS6k, GAPDH) dilution) and HRP-conjugated anti-mouse IgG (Promega, Cat# W402B; 1:2,500 (Akt) and 1:10,000 (β-actin) dilution), for 60 minutes at room temperature. The signals were detected by ECL Prime Western Blotting Detection Reagent (cytiva, Cat# RPN2232). The membranes were then visualized using an Image Quant LAS4010 (GE HealthCare, Cat# LAS4010). Quantification of the band intensity was determined using the Image J wand tool.

Quantitative evaluation of protein synthesis by SUnSET analysis

Differentiated C2C12 cells were cultured overnight in DMEM medium without FBS, and sEGFR was added at concentrations of 0, 0.5, 2 nM and incubated at 37°C for 24 hours. The cells were treated with 1 μM puromycin for one hour at 37°C and collected. The subsequent protein extraction and Western blotting were performed as described above, except for the antibodies. Primary antibodies were anti-puromycin (EMD Millipore Corp, Cat# MABE343; 1:1,000 dilution) and anti-β-actin (MBL, Cat# M177-3; 1:5,000 dilution). The secondary antibody was HRP-conjugated anti-mouse IgG (Promega, Cat# W402B). The antibodies were used at concentrations of 1:2,500 dilution for anti-puromycin and 1:10,000 dilution for anti-β-actin. All bands in each lane were quantified, and quantification of the band intensity was determined using the Image J wand tool.

Statistical analyses

All statistical analyses were performed by GraphPad Prism, version 10. Significant differences between mean values were evaluated using unpaired t-tests with Welch’s correction (when two groups were analyzed) or Brown-Forsythe and Welch ANOVA tests followed by Dunett’s T3 multiple comparisons tests (when three groups were analyzed).

Results

The concentration of EGFR in serum positively correlates with androgen levels

To investigate the liquid factors in blood that vary with androgen levels, we generated three mouse groups: the sham surgery group, the androgen deprivation through orchidectomy (ORX) group, and the dihydrotestosterone (DHT) replacement group after ORX (Fig. 1A). The body weight of mice receiving DHT replacement significantly increased when compared to the other two groups (Fig. 1B). As we confirmed that skeletal muscle mass, especially of the levator ani/bulbocavernosus (LA/BC) muscles known for its high androgen sensitivity [23], decreased with androgen depletion due to ORX, and increased with DHT replacement (Fig. 1C), a proteomic analysis was performed on blood serum samples collected from the three groups after a 2-week treatment period, utilizing mass spectrometry. We identified approximately 450 proteins, among which a subset showed a positive correlation with androgen levels—specifically, those that decreased in the ORX group and increased with androgen replacement (Table 1). Epidermal growth factor receptor (EGFR) was identified as the protein most strongly correlated with serum androgen levels, characterized by the highest count of unique peptides. These observations were consistent after both 4 and 8 weeks of Veh or DHT administration, as assessed by ELISA (Fig. 1D). Considering that males inherently have higher androgen levels than females [24], we examined the concentration of EGFR between sexes. EGFR concentration was significantly higher in male than female mice (Fig. 1E), emphasizing a higher androgen dependency. Furthermore, the Human Genetic Evidence (HuGE) score [25], which quantifies genetic support for the involvement of genes in a disease or trait based on several human genetic outcomes, showed that appendicular lean body mass was predominantly associated with EGFR (Fig. 1F). However, there is no evidence that there is a sex difference in appendicular lean body mass or that it is significant in relation to EGFR. Furthermore, it has not been reported that SNPs in the AR gene are mainly correlated with appendicular lean body mass. Taken together, our findings show a positive correlation between androgen levels and serum EGFR concentrations, indicating that EGFR varies in an androgen-dependent manner.

Fig. 1  The concentration of EGFR in blood serum correlates with androgen levels. (A) Sham or ORX surgery was performed on 10-week-old male mice, and Veh or DHT (2.5 mg/mice/week) was administered every week; n = 5 mice for each group. At 12 weeks of age, the tibialis anterior (TA), gastrocnemius, soleus, levator ani/bulbocavernosus (LA/BC) muscles and blood were sampled. (B) Change of body weight (left) and the body weight at 12 weeks old (right). (C) Comparison of skeletal muscle mass at 12 weeks of age. (D) Concentration of EGFR using ELISA in serum for 4 and 8 weeks of DHT administration; n = 4 or 5/group. Brown-Forsythe and Welch ANOVA tests. (E) Concentration of EGFR in serum of male and female C57BL/6J mice using ELISA; n = 5/group. Welch’s t-test. (F) Human Genetic Evidence (HuGE) scores showing the association between appendicular lean body mass and EGFR locus. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

Table 1 The list of proteins identified to exhibit a correlation with androgen concentrations in the blood serum.

Gene name	Description	(ORX + Veh)/(sham + Veh)	(ORX + DHT)/(ORX + Veh)	Unique Peptide
Egfr	epidermal growth factor receptor	0.519	1.685	31
Tln1	talin 1	0.826	1.645	3
Hbb-b1	hemoglobin, beta adult major chain	0.743	1.603	11
Hba	hemoglobin alpha chain complex	0.792	1.473	9
Mb	myoglobin	0.794	1.453	4
Cap1	CAP, adenylate cyclase-associated protein 1 (yeast)	0.746	1.403	2
Ces2e	carboxylesterase 2E	0.729	1.401	6
Ttr	transthyretin	0.653	1.335	8
Cycs	cytochrome c, somatic	0.723	1.241	2
Blvrb	biliverdin reductase B (flavin reductase (NADPH))	0.772	1.238	4

EGFR in blood serum could be supplied from non-myofibers by AR

EGFR, a membrane-bound receptor, is released into the blood as a soluble receptor through ectodomain shedding of receptors on the membrane [26] or alternate splicing of mRNA transcripts [27]. To identify the cellular or tissue origin of soluble EGFR (sEGFR), we investigated Egfr expression across various tissues in mice with modulated androgen levels, including both male and female mice using quantitative RT-PCR (Fig.2). A positive correlation was observed between androgen concentrations and Egfr expression in liver and epididymal white adipose tissue (eWAT), commonly known as visceral fat (Fig. 2A). Furthermore, the expression of Egfr was significantly higher in male than female mice in liver and eWAT (Fig. 2B). These results indicate that the expression of Egfr in liver and eWAT, which are candidates for the origin of sEGFR, could be regulated by androgen.

Fig. 2  The presence of EGFR in blood serum could be attributed to its supply from non-myofiber tissues mediated by AR. (A) Egfr transcript in liver, kidney, tibialis anterior (TA), subcutaneous white adipose tissue (sWAT), and epididymal WAT (eWAT) of mice subjected to androgen deprivation and subsequent replacement; n = 5/group. (B) The relative mRNA expression of Egfr in the tissues from male and female C57BL/6J mice; n = 5/group. Welch’s t-test. **p < 0.01, ***p < 0.001.

The regulation of EGFR transcription is controlled by AR

To investigate the regulation of EGFR by androgens and AR, we examined the involvement of AR in the transcriptional regulation of EGFR expression by DHT administration. Previous ChIP-seq data demonstrated that AR binding peaks were found both upstream and downstream of the promoter region of EGFR in LNCaP, a human prostate cancer cell line with lymph node metastasis, and VCaP, a human prostate cancer cell line derived from spinal metastasis (Fig. 3A). To confirm the binding of AR at the EGFR locus, we treated LNCaP with DHT for 12 hours for ChIP-qPCR (Fig. 3B). Indeed, AR binding in the promoter region of EGFR was induced by DHT treatment (Fig. 3C), alongside the AR binding to kallikrein-related peptidase 3 (KLK3), whose expression is highly androgen-responsive [28, 29]. Furthermore, previous ChIP-seq data also demonstrated that AR binding peaks were localized in the transcription start site (TSS) of Egfr in epithelium of prostates of mice (Fig. 3D). We identified the Androgen response element (ARE) sequence and verified that it is highly conserved (Fig. 3E). To confirm the binding of AR at the Egfr locus in mouse adipocytes, we treated 3T3-L1, which is a mouse fibroblast cell line used to study adipogenesis, with DHT (Fig. 3F). In the cells differentiated into adipocyte-like cells, ChIP-qPCR confirmed that the AR binding in TSS of Egfr was induced by DHT treatment (Fig. 3G). These results suggest that Egfr expression, at least adipocyte cell lines, is directly regulated by androgens via the AR.

Fig. 3  Transcriptional regulation of EGFR is mediated by AR. (A) AR peaks by ChIP-seq around the promoter region of the EGFR locus in the human prostate cell line illustrated by IGV. Primer A was designed for the transcription start site of EGFR and primer B for the peak near the center of EGFR. (B) ChIP-qPCR preparation protocol for LNCaP. (C) ChIP relative quantity showing the induction of AR binding, to the EGFR locus, alongside the KLK3 locus, in LNCaP treated with DHT. (D) AR peaks by ChIP-seq around the promoter region of the Egfr locus in mouse epithelium of prostates. The primer was designed for near the transcription start site of Egfr. (E) Conservation of the ARE sequence, located within the primer target region shown in (D). (F) ChIP-qPCR preparation protocol of 3T3-L1. (G) ChIP relative quantity showing the induction of AR binding to Egfr locus in 3T3-L1 treated with DHT; n = 3/group. Welch’s t-test. *p < 0.05, **p < 0.01, ***p < 0.001.

sEGFR promotes Akt signaling and protein synthesis in vitro

The Akt signaling pathway plays an important role in skeletal muscle growth, survival, and metabolism [30, 31]. To examine whether EGFR is involved in Akt signaling in skeletal muscle cells, differentiated C2C12 cells were treated with sEGFR for 5 minutes in serum-deprived medium. We found that phosphorylation of Akt and its downstream p70 S6 Kinase (S6k) was significantly increased by the treatment with sEGFR (Fig. 4A, B). The degree of phosphorylation is dependent on the concentration of sEGFR (Fig. 4B). Furthermore, to investigate whether protein synthesis is promoted by sEGFR, SUnSET analysis [32] was performed in differentiated C2C12 treated with sEGFR for 24 hours. EGFR tended to promote protein synthesis compared to controls (Fig. 4C, D). These results suggest that sEGFR treatment in skeletal muscle cell lines promotes Akt signaling and protein synthesis in vitro.

Fig. 4  Administration of sEGFR promotes phosphorylation of Akt signaling in vitro. (A) Evaluation of Akt and S6K phosphorylation by western blotting (WB) using C2C12 after treatment with sEGFR. (B) Quantification of WB images. (C) Evaluation of protein synthesis by SUnSET analysis. The leftmost lane is a sample taken without puromycin added. All other lanes have puromycin added. (D) Quantification of the proteins shown in (C). Brown-Forsythe and Welch ANOVA tests. *p < 0.05, **p < 0.01, ***p < 0.001.

Administration of EGFR contributed to an increase in skeletal muscle mass of female mice

To explore the influence of EGFR on skeletal muscle mass in vivo, we administered recombinant sEGFR protein via intraperitoneal (i.p.) injections to female mice which have naturally low androgen levels [24], thus allowing a clearer assessment of androgen’s role. Eight-week-old female mice received weekly i.p. injections of either a control vehicle (Veh), recombinant sEGFR protein, or DHT for two weeks (Fig. 5A). No significant weight difference was observed between the Veh and sEGFR-administered groups, whereas the DHT-treated group exhibited a significant increase in body weight compared to the other groups after two weeks of treatment (Fig. 5B). Importantly, the sEGFR-administered group showed a significant increase of gastrocnemius muscle mass relative to the control group as well as in the DHT-treated group (Fig. 5C). We also found that the weight of eWAT was increased predominantly by DHT, but did not show a consistent trend with sWAT (Fig. 5D). Given the established cardiovascular event risks associated with androgen replacement, we also measured cardiac weight. In the DHT-treated group, the weight of the heart was significantly greater compared to the control, whereas the sEGFR-treated group did not display this effect (Fig. 5E). Grip strength tests also revealed that sEGFR treatment increased muscle strength (Fig. 5F). Furthermore, Akt signaling in the gastrocnemius muscle at 10 weeks of age treated with Veh, sEGFR or DHT was examined in WB. We found that Akt phosphorylation was predominantly enhanced by sEGFR administration in vivo (Fig. 5G, H) as well as in vitro. While the effects induced by DHT are more pronounced, our data indicate that administering EGFR to female mice could potentially contribute to the increase of skeletal muscle mass.

Fig. 5  Administration of sEGFR contributes to the increase in skeletal muscle mass of female mice. (A) Experimental design for sEGFR treatment. Veh and DHT (2.5 mg/mice/week) were intraperitoneally administered and sEGFR (20 μg/mice/week) was subcutaneously administered to female mice every week starting at 8 weeks of age; n = 5 /group. At 10 weeks of age, skeletal muscles were sampled. (B) Alteration in body weight (left) and body weight at 10 weeks of age (right). (C–F) Comparison of skeletal muscles mass (C), eWAT and sWAT (D), heart (E), and grip test (F) at 10 weeks of age. (G) Evaluation of Akt phosphorylation by WB using gastrocnemius muscle at 10 weeks of age; n = 5 /group. (H) Quantification of the proteins shown in (G). Brown-Forsythe and Welch ANOVA tests. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

Discussion

The aim of this study was to identify androgens’ indirect mechanism governing skeletal muscle mass via non-myofibers. We identified proteins in serum that vary in concentration in response to androgen levels through ORX-induced androgen depletion and androgen replacement in male mice by LC-MS/MS. Especially, the level of EGFR exhibited changes consistent with androgen level. Previous studies have primarily examined sEGFR in the context of cancer. Plasma sEGFR has been found to be valuable in predicting survival in advanced non-small cell lung cancer [33]. Additionally, sEGFR serves as a useful marker for predicting prognosis or therapeutic response in cervical, colorectal, ovarian, and breast cancers [34]. In contrast, we showed for the first time that the EGFR level is lower in mice with low androgen, including female mice, and higher in androgen treated mice. On the other hand, the protein profiles identified in our study did not perfectly align with previous analyses of blood protein changes in response to androgen levels in humans [35]. Giwercman et al. [35] reported that insulin-like growth factor-binding protein 6 (IGFBP6) level changed in the same direction as androgen. The concentration of EGFR was constant in human samples regardless of androgen concentration in blood. Using a chemical castration human model could explain these discrepancies. The serum EGFR concentrations identified by ELISA in this study were much higher compared to the previous report in humans [36]. Although there are variations in EGFR blood levels between species, our data demonstrate a relative relationship between androgens and EGFR in mice. The causes of species-specific differences in androgen blood levels and blood proteins require further study.

We found that liver and eWAT were potential organs for the origin of serum EGFR regulated by androgen. Because alternative splicing of EGFR is found in liver [27], sEGFR could be derived from these transcripts’ variants. On the other hand, ectodomain shedding is another mechanism to produce sEGFR [26]. While the presence of shedding-derived sEGFR is demonstrated in conditioned medium of both an immortalized keratinocyte cell line [37] and breast cancer cells [36], there is no conclusive evidence pointing to a specific normal tissue as the exclusive source, and it remains plausible that multiple tissues contribute to its production. The specific sequence of EGFR released from the cell membrane, if known, would further contribute to the identification of its origin. We also confirmed previous reports [19, 20] by ChIP-qPCR that adding androgen increased AR binding to the promoter region of EGFR in human prostate cancer cell lines, as well as the peaks of AR by ChIP-seq. The EGFR signaling pathway could potentially serve as a mechanism through which AR drives the aggressive biology of epithelial ovarian cancer [38, 39]. Androgens have been demonstrated to up-regulate EGFR expression in prostate carcinoma cells, leading to enhanced cellular proliferation [40]. The actual binding of AR to the EGFR locus in candidate organs including liver and adipose tissue should be investigated in the future.

The Akt signaling pathway is crucial for the development, survival, and metabolic processes of skeletal muscle, significantly contributing to the maintenance of muscle function and integrity [30, 31]. This pathway oversees various cellular activities including growth, division, and survival, highlighting its fundamental role in muscle physiology. Activation of Akt promotes protein synthesis via the mTOR (mammalian target of rapamycin) pathway, resulting in muscle hypertrophy [30, 31]. This is important in muscle hypertrophy following training and exercise [41, 42]. We showed that the presence of sEGFR promoted phosphorylation of Akt and S6k downstream of the signal, regardless of its concentration. Under the same conditions, it tended to promote protein synthesis. In general, EGFR ligands enhance Akt signaling [43]. However, there are no reports that sEGFR is involved in Akt signaling. Our finding that the presence of sEGFR promotes Akt signaling and protein synthesis suggests the existence of an entirely new signaling system, either by binding to EGF ligands on the membrane and generating reverse-signals into the cell, or by the presence of an unknown receptor for sEGFR. The results that sEGFR enhanced Akt signaling may rule out the function of sEGFR as a decoy receptor for EGF ligands. We observed that DHT administration resulted in a greater increase in skeletal muscle mass compared to sEGFR administration. On the other hand, our in vitro experiments suggest that EGFR can directly influence skeletal muscle by rapidly activating AKT phosphorylation. Given that DHT has been shown to elevate sEGFR levels, it is possible that DHT promotes skeletal muscle mass not only through direct AR signaling but also through the indirect activation of AKT signaling via EGFR. This dual mechanism may explain why DHT administration leads to a more pronounced increase in skeletal muscle mass compared to sEGFR administration alone. The mechanism by which sEGFR enhances Akt signaling is still unknown and requires further study.

We found that LA/BC muscle mass was greater than the control without the complete recovery of sEGFR concentration after DHT treatment. These findings imply the involvement of regulatory mechanisms beyond the sEGFR-mediated pathway in the indirect regulation of skeletal muscle mass by DHT. Actually, DHT treatment can increase muscle mass even in the absence of AR in muscle fibers [15]. We recently demonstrated that skeletal muscle mass may be regulated through AR located on non-muscle fibers [44]. Additionally, glucocorticoids synthesized in the zone fasciculata of the adrenal cortex under androgen control are involved in regulating skeletal muscle size [45]. These results indicate that skeletal muscle mass is regulated by non-muscle cells expressing AR, either within or outside of skeletal muscle tissue. Further research is needed to clarify how these various indirect mechanisms of androgens work together to regulate skeletal muscle mass.

Upon administering recombinant sEGFR protein to female mice, hypertrophic tendencies were observed in the gastrocnemius muscle. Muscle strength was also enhanced by the administration of sEGFR, which may be due to the effect of increased skeletal muscle mass. It has been reported that downregulation of EGFR promotes normal differentiation in early differentiation of human myoblasts [46], indicating that EGFR is an important factor in the regulation of muscle differentiation. Blocking EGFR signaling in slow-twitch muscle promotes muscle development and maintenance [47]. Because EGFR is expressed in both muscle fibers and skeletal muscle stem cells [48], skeletal muscle can be a target of EGFR ligand including EGF. Actually, the administration of EGF decreases body weight induced by energy metabolism alteration, as EGF administration increases the mRNA level of UCP3 in skeletal muscles [49]. Because sEGFR contains only the extracellular domain [36, 50], and as a decoy receptor has high affinity for ligands that inhibit EGF signaling activity [51], EGFR treatment could suppress EGF signaling leading to the increase of muscle mass. On the other hand, there are reports to the contrary that EGF promotes satellite cell proliferation [52]. Upregulation of heparin-binding epidermal growth factor-like growth factor (HB-EGF), which is another ligand to EGFR, in exercise-contracted skeletal muscle has been suggested to act as an insulin sensitizer that promotes glucose consumption, but overexpression of HB-EGF did not induce hypertrophy [53]. One potential explanation for this inconsistency is the age and sex of mice, which drastically affect the muscle physiology [54-56]. Another possible mechanism is reverse signaling, which is induced by binding of sEGFR to membrane-bound EGF ligands [57]. Furthermore, when the ectodomain of a membrane-type EGF ligand, such as HB-EGF, undergoes shedding, it produces soluble fragments (HB-EGF) and carboxy-terminal fragments (HB-EGF-CTF) [57]. Electron microscopy has revealed the presence of HB-EGF-CTF in the inner nuclear envelope, suggesting its potential involvement in transcriptional activity [57]. Although EGFR knockout mice are embryonic lethal or have incomplete organogenesis [58, 59], further analysis in models of EGFR deficiency or inhibition will be required for clarifying mechanisms of the increase of skeletal muscle mass by sEGFR. The stability and biological activity of the recombinant proteins used in this study are unknown and are limitations of this experiment.

In humans, administration of androgen has long been reported to cause serious cardiovascular adverse events [12]. On the other hand, androgen replacement therapy for hypogonadal patients has also been reported to be non-inferior to placebo in the incidence of cardiovascular events [60], although the cardiovascular impact is noteworthy. Because sEGFR administration increased skeletal muscle mass but did not alter heart weight, EGFR could be considered as a potential candidate for treating muscle atrophy including sarcopenia without causing cardiovascular side effects.

We propose that skeletal muscle mass is indirectly regulated by androgens while being modulated by EGFR via AR outside skeletal muscle tissue. Alternatively, sEGFR acts on other tissues and affects skeletal muscle via secondary factors (Graphical Abstract). Given that the maintenance of skeletal muscle is an essential component of healthy life expectancy in the elderly, our study provides the mechanisms by which androgens regulate skeletal muscle through non-myofibers and the molecular basis for the treatment of various muscle atrophy conditions associated with androgen deficiency.

Graphical Abstract The proposed model for indirect regulation by androgen through EGFR via AR in non-myofibers

Acknowledgments

We thank the Division of the Advanced Research Support Center (ADRES), Laboratory Animal Research, Ms. Yuka Sato and members of the Division of Pathophysiology, Proteo-Science Center (PROS), Ms. Izumi Tanimoto of the Division of Urology of Ehime University. The illustrations in the figure were created with BioRender.com.

Author Contributions

Tomoya Onishi (Conceptualization, Methodology, Formal analysis, Investigation, Data curation, Writing—original draft, visualization), Hiroshi Sakai (Methodology, Formal analysis, Investigation, Funding acquisition, Writing—review and editing), Hideaki Uno (Methodology), Iori Sakakibara (Methodology, Formal analysis, Investigation), Akiyoshi Uezumi (Methodology), Mamoru Honda (Resources), Tsutomu Kai (Resources), Shigeki Higashiyama (Resources), Noriyoshi Miura (Conceptualization, Supervision, Funding acquisition, Writing—review and editing), Tadahiko Kikugawa (Conceptualization, Supervision, Funding acquisition, Writing—review and editing), Takashi Saika (Conceptualization, Supervision, Funding acquisition, Writing—review and editing), and Yuuki Imai (Conceptualization, Methodology, Resources, Funding acquisition, Project administration, Supervision, Writing—review and editing)

Data Availability

The parts of results and analytical data obtained in this study are not disclosed in the paper. We will disclose raw data of the LC–MS/MS, etc., upon reasonable request.

Funding

This work was supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI (Grant Number JP21K09373).

Disclosure

None of the authors have any potential conflicts of interest associated with this research. Yuuki Imai is a member of Endocrine Journal’s Editorial Board.

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