Introduction
Gastric cancer(GC) is the fifth most commonly diagnosed malignant tumor and the fourth leading cause of cancer death globally [
1], causing a severe economic burden. Multidisciplinary treatment has become the mainstay for GC patients, especially with the development of immunotherapy and targeted therapy [
2]. However, the number of patients benefiting from the new treatment is limited due to the heterogeneity of the patient population. Therefore, it is critical to investigate the fundamental molecular mechanisms of GC pathogenesis and identify new diagnostic biomarkers and therapeutic targets.
Autophagy is a self-eating process that recycles wastes of cells and maintains homeostasis by clearing longevity proteins or damaged organelles. These cellular contents and organelles are sequestered in double-membrane structures called autophagosomes, which are then fused with lysosomes and degraded [
3]. Autophagy is activated when cells encounter environmental stress, such as malnutrition, hypoxia, pathogen infection, and oxidative stress, and cells either adapt or die depending on the intensity of the stimulus and the response of the host [
4]. Autophagy is reported to exhibit crucial but conflicting functions in the progression of various tumors, including pancreatic adenocarcinoma, myeloid leukemia, gastric carcinoma, squamous cell carcinoma, sarcoma, and multiple myeloma [
5‐
8]. More importantly, there have been several clinical trials targeting autophagy in malignant glioma, non-small cell lung cancer, colon cancer, melanoma, pancreatic cancer and melanoma, implicating promising prospects for targeting autophagy therapy [
9].
Arginine methylation, which is regulated by the protein arginine methyltransferase (PRMT) family, is one of the most vital epigenetic modifications in autophagy [
10]. Coactivator-associated arginine methyltransferase-1 (CARM1) plays a special role because it is the only enzyme capable of methylating arginine with proline sequences [
11]. CARM1 has two main domains, which are required for the interaction with chromatin remodeling proteins and proteins possessing RNA binding activity, and the resultant protein complex plays essential roles in chromatin remodeling, gene transcription, DNA packing, pre-mRNA splicing, and regulation of mRNA stability [
12,
13]. The role of CARM1 in cancers is paradoxical. It acts as an oncogene in lung, prostate, colorectal, ovarian, breast cancer, osteosarcoma, myeloid leukemia and multiple myeloma [
14‐
23]. However, CARM1 inhibits the progression of liver and pancreatic cancers [
19,
20]. Previous studies have mainly focused on the role of CARM1 as a coactivator in regulating tumor-related gene expression [
24,
25]. Recent studies have shown that CARM1 is an important regulator of autophagy which functions mainly through two dependent pathways. In the nucleus, AMPK is activated under glucose starvation and subsequently phosphorylates and activates FOXO3a. This leads to a decrease in SKP2 expression and resulting in the stabilization of CARM1 in the nucleus, which acts as a coactivator of TFEB to promote autophagy-related gene transcription [
26]. In the cytoplasm, C9orf72 mediates CARM1 lysosomal degradation by interacting with the PH-like domain and modulates lipid metabolism [
27]. However, the role of CARM1 in GC has not been reported previously, and it remains unclear whether CARM1 could affect the progression of GC by regulating autophagy.
In this study, we illustrate the effect of CARM1 on GC and the mechanism by which CARM1 regulates autophagy. CARM1 and autophagy markers were upregulated in GC tissues, and higher CARM1 expression predicted worse prognosis. Knockdown of CARM1 interfered with the orderly progression of autophagy and subsequent ER stress-induced apoptosis and reduced cell proliferation by promoting G1 phase cell cycle arrest, which could be partially reversed by the autophagy agonist rapamycin. Overexpression of CARM1 exhibited the opposite results. More importantly, treatment with a CARM1 inhibitor rescued the tumor-promoting effects of CARM1 both in vitro and in vivo. Furthermore, we demonstrated that CARM1 facilitated TFE3 nuclear translocation to induce autophagy through the cytoplasmic AMPK-mTOR and nuclear AMPK-CARM1-TFE3 signaling pathways. Thus, our findings suggest that CARM1, either by itself or in combination with other signaling molecules, may serve as a novel therapeutic target for the treatment of GC.
Methods
Tissue microarray and clinical samples
Tissue microarrays (G6038-3), which comprise 48-paired GC tissues and matched adjacent non-neoplastic tissues were obtained from Servicebio Biotech (Wuhan, China). They contained clinical information such as sex, age, tumor location and size, metastasis, vessel carcinoma embolus and the overall survival of patients. After the immunohistochemical staining, immunostaining intensity was scored by two experienced pathologists, who were blinded of clinical information. The percentage of positive cells and the staining intensity were evaluated semi-quantitatively, the percentage of positive cells was divided into 5 grades and scores were assigned as follows: the number of positive cells < 5% was 0, 5–25% was 1, 26–50% was 2, 51–75% was 3, 76–100% was 4 points. The staining intensity was graded in 4 grades: negative was 0, weak was 1, moderate was 2, and strong was 3 points. The total scores, ranging between 0 and 12, were calculated by multiplying the positivity score (0–4) by the intensity score (0–3). Tissues with a total score < 7 were classified as low expression, while those with a score ≥ 7 were classified as high expression. Thirty-three pairs of clinical tissues were collected from the Second Affiliated Hospital of Xi’an Jiaotong University with informed consent from patients.
Immunohistochemistry (IHC)
Tissue microarrays and paraffin sections were soaked in xylene twice for deparaffinization. The slides were rehydrated with a series of ethanol solutions (100%, 95%, 85% and 75%). Slides were evenly spaced in citrate buffer and microwaved for perform antigen retrieval. Subsequently, 3% hydrogen peroxide was added to remove endogenous peroxidase, and then the slides were blocked with goat serum for 30 min at room temperature (RT). Primary antibodies were added at appropriate concentrations and the slides were incubated at 4 °C overnight. The next day, biotinylated secondary antibodies were added and the slides were incubated at RT for 1 h. To visualize antibody binding, fresh DAB staining solution was added, and the staining process was monitored under a microscope. When the staining was complete, slides were counterstained in hematoxylin bath for 30–60 s, dehydrated in graded ethanol (75%, 85%, 95% and 100%) transferred to dimethylbenzene, and sealed in neutral resin.
RNA extraction and qRT–PCR
Total RNA was extracted by TRIzol (Invitrogen, Thermo, Waltham, MA, USA). The messenger RNA was reverse-transcribed with the Transcriptor First-Strand cDNA Synthesis Kit (Roche, Basel, Switzerland). Subsequently, FastStart Universal SYBR Green Master (Roche) was used to conduct qRT
–PCR analysis following the manufacturer’s instructions. All primer sequences are listed in Additional file
4: Table S2.
Cell culture
The human GC cell lines HGC27, BGC823, MKN45, SNU-1 and the normal gastric epithelial cell line GES1 were purchased from GeneChem (Shanghai, China), identified by STR profiling and tested free of mycoplasma contamination. All cell lines were cultured in high glucose DMEM (HyClone, Logan, Utah, USA) supplemented with 10% fetal bovine serum (Gemini, Calabasas, CA, USA) and 1% penicillin–streptomycin (Gibco, Grand Island, NY, USA) in a humidified incubator in 5% CO2 at 37 °C.
Western blotting
Total protein was extracted using cold RIPA buffer (Beyotime, Shanghai, China). Cytoplasmic and nuclear extracts were separated by NE-PER Nuclear and Cytoplasmic Extraction Reagents (Thermo Scientific™, MA, USA). The protein concentration of each cell lysate was determined using a BCA Protein Assay Kit (TIANGEN, Beijing, China). The equal amount of protein was separated by SDS-PAGE gel and transferred to PVDF membranes. The membrane was incubated overnight at 4 °C in a primary antibody solution. The antibodies used in this study are shown in Additional file
5: Table S3. The next day, the membrane was incubated with horseradish peroxidase-conjugated secondary antibody and visualized with enhanced chemiluminescence reagents.
Lentivirus and siRNA transfection
The overexpression lentivirus and control lentivirus were obtained from Hanbio Biotechnology (Shanghai, China). The overexpression lentivirus was HBLV-h-CARM1-3xflag-ZsGreen-PURO, and the control vector was HBLV-ZsGreen-PURO. Transduction was performed according to the manufacturer’s protocol. Transduced cells were cultured in medium containing 2.5 μg/mL puromycin to select for cells stably expressing the transduced construct. HGC27 and BGC823 CARM1-overexpressing cells were transfected with specific siRNAs (GenePharma, Shanghai, China) targeting CARM1 and TFE3, respectively. Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA) was used for transfection in accordance with the manufacturer’s instructions. The interference sequences used in this study are shown in Additional file
4: Table S2.
Transmission electron microscopy
Cells were harvested and fixed at 4 °C for 2–4 h with electron microscope fixation solution (Servicebio, China) and further fixed with 1% osmium in·0.1 M phosphate buffer for 2 h. Cells were dehydrated in graded ethanol (50%, 70%, 80%, 90%, 95% and 100%) and 100% acetone twice. Cells were embedded and cut into slices of 60–80 nm thickness with an ultra-thin slicing machine. Then, the sections were double-stained with 2% uranium acetate-lead citrate. Finally, photographs of eight random fields were captured for each sample under a transmission electron microscope (HITACHI, Tokyo, Japan).
Immunofluorescence staining
Cells were cultured on small cover glass. 4% paraformaldehyde was added into the hole for cell fixation for 30 min at RT, cells were permeabilized with 0.1% Triton 100 for 15 min. Subsequently, cells were blocked with 5% goat serum and incubated without washing with a sufficient amount of primary antibodies at appropriate concentration overnight in a moist container at 4 °C. The following day, the cells were incubated with fluorescence-labeled secondary antibody for 1 h and DAPI for 1 min in the dark. Finally, the coverslip was sealed with anti-fading buffer (BD Biosciences, NJ, USA). Photographs of five random fields were captured under a confocal microscope (Nikon C2, Tokyo, Japan).
CCK-8 assay
Cell suspensions were added to 96-well plates at 3 × 103 cells per well in 100 μL DMEM and incubated for 1–4 days. Ten microliters of CCK-8 reagent (7 Sea Pharmatech Co., Ltd, Shanghai, China) was added to each well and the plates were incubated in a cell incubator for 1 h. The optical absorbance was read at 450 nm on a microplate spectrophotometer (Thermo, Waltham, MA, USA).
Cells in the exponential growth phase were plated at a density of 1 × 103 cells per well in 6-well plates and cultured for 10–14 days until visible colonies were formed. Cells were fed with fresh culture medium in a timely manner according to the pH change. When the colonies were ready for analysis, the cells were fixed with 4% paraformaldehyde for 20 min and stained with crystal violet dye for 10 min. Finally, the number of colonies containing more than 50 cells was counted.
Flow cytometry
A total of 2 × 105 cells were plated in 6-well plates, grown in serum-free medium for 24 h to synchronize the cell cycle, and cultured for another 24 h. The adherent cells were digested and fixed with 70% ice-cold ethanol at 4 °C overnight. The next day, the fixed cells were washed and resuspended in 500 μL PI/RNase Staining Buffer (BD Biosciences, Franklin Lakes, NJ, USA). After incubating for 15 min in the dark at RT, cell cycle was analyzed by flow cytometry (BD Biosciences, Franklin Lakes, NJ, USA).
A total of 1.5 × 105 cells were seeded in each well of 6-well plates and cultured to a confluence of 80%-90%. Afterwards, the proportion of apoptotic cells was determined by staining with a PE Annexin V/7-amino-actinomycin (7-AAD) Detection Kit (BD Biosciences, NJ, USA) following the manufacturer’s instructions and analyzed by flow cytometry.
In vivo xenograft tumor model
A subcutaneous xenograft model was established using 5 to 6-week-old male BALB/c nude mice (Xi’an Jiaotong University Medical Laboratory Animal Center, Xi’an, China). The experiments were approved by the animal ethics committee of Xi'an Jiaotong University. One million (1.0 × 106) tumor cells were injected subcutaneously into the left groin of nude mice where blood flow was abundant. The volume of subcutaneous tumor was evaluated every 3 days. To test the efficacy of drug treatment, nude mice injected with CARM1-overexpressing cells were randomly divided into four groups on day 6 when tumor volumes reached 50 mm3. The CARM1 inhibitor EZM2302 was administered twice a day at 100 mg/kg i.p. HCQ was administered once a day at 50 mg/kg i.p. The combination group received both EZM2302 and HCQ treatments and the control group was intraperitoneally injected with PBS. When the subcutaneous tumor showed ulceration and necrosis, the mice were anesthetized with 3% sodium pentobarbital and euthanized with a high level of carbon dioxide. Then, the transplanted tumors were removed, and the weight and volume were measured.
Immunoprecipitation assay
The cells were harvested and incubated with an appropriate amount of IP lysis buffer (containing protease inhibitor) for 30 min at 4 °C. After centrifugation, 6 μg CARM1 antibody or control IgG and 40 μL Protein A/G Mix Magnetic Beads (Merck Millipore, Germany) were added to the supernatant and incubated at 4 °C overnight. After the immunoprecipitation reaction, the protein was carefully eluted from the protein A/G-beads and denatured for western blotting analysis.
Statistical analysis
We used GraphPad Prism 6 (San Diego, CA, USA) and SPSS 20.0 (Chicago, IL, USA) to perform statistical analysis. When comparing the differences in measurement data between the two groups, Student’s t test was used. When comparing the differences between more than two groups, ANOVA was applied, in which p values were adjusted for multiple comparisons. Before Student’s t test and ANOVA, a variance homogeneity test and normality analysis were carried out. Overall survival curves were analyzed by the Kaplan–Meier method and log-rank tests. The chi-squared test was utilized to estimate the difference in CARM1 levels between GC tissues and adjacent non-tumor tissues. The hazard ratio in clinical samples was determined by a Cox proportional hazards model for univariate and multivariate analyses. All values demonstrate means ± SD. All statistical tests were two-sided, and P < 0.05 was considered statistically significant.
Discussion
GC remains one of the deadliest malignancies, and there is a lack of effective early diagnosis and prognostic molecular markers. CARM1 is elevated in various tumors and exhibits a tumor-promoting role mainly as a transcriptional coactivator by methylating histones and non-histones [
25,
39]. Paradoxically, CARM1 shows a tumor-inhibiting effect on liver and pancreatic cancers [
19,
20]. However, the role of CARM1 and the associated mechanism of pathogenesis in GC have not been studied before. Here, we demonstrated that CARM1 expression was increased by analyzing data in databases and clinical samples of GC and proved for the first time that CARM1 was an independent risk factor for predicting poor survival (Fig.
1A–D). These findings indicate that CARM1 may potentially become a marker for diagnosis and therapeutic decision making for GC.
Recent studies have shown the vital role of CARM1 in autophagy [
11]. Autophagy maintains cellular homeostasis through the degradation of long-lived proteins and damaged organelles, and is considered to be a self-defense mechanism against certain stressful conditions [
40]. The role of autophagy in tumors is paradoxical depending on the specific environment [
41,
42]. On the one hand, autophagy markers are often located in cancer-related DNA regions with frequent mutations or deletions which may be protective mechanisms to inhibit tumor initiation [
43], and it has been proven that 5-FU can inhibit GC cell growth and survival by inducing autophagy-related death [
44]. On the other hand, autophagy can help tumor cells resist nutritional deficiencies and other cancer-associated conditions to promote tumor progression and chemotherapy resistance [
45]. In this study, we demonstrated that CARM1 promoted GC cell proliferation through autophagy-regulated G1-S transition (Figs.
4A–H,
5A, B).
Furthermore, our results showed that defective autophagy process caused by CARM1 inhibition contributed to ER stress-related apoptosis, which also affected tumor growth. There are three main UPR signaling pathways initiated by IRE1α, PERK, and ATF6, respectively, to reduce protein load and enhance protein-folding capacity [
46], and we found that the ER stress initiated by CARM1-mediated autophagy was regulated mainly by the PERK pathway (Fig.
3B, C). PERK undergoes dimerization and autophosphorylation, which activates its kinase domain and subsequent phosphorylation of eIF2α. p-eIF2α relieves the unfolded protein burden of ER [
47] by inhibiting most protein translation but selectively enhancing ATF4 translation, leading to an increase in downstream protein CHOP expression [
48]. CHOP is responsible for the apoptosis of cells with dysfunctional ER [
49]. In our study, increased ER stress mediated by impaired autophagy promoted CHOP expression and eventually potentiated apoptosis (Fig.
5C).
Our results demonstrated that CARM1 exerted a significant tumor-promoting role in GC, therefore, we sought to explore the therapeutic effect of targeting CARM1 by utilizing the small molecule compound EZM2302, which is a specific inhibitor of CARM1 and has shown a significant inhibitory effect on multiple myeloma both in vitro and in vivo [
21]. In mouse models of AML and diffuse large B-cell lymphoma, CARM1i also exhibited an effective inhibitory effect [
50,
51]. Hence, we wished to determine whether CARM1i could also inhibit the progression of GC, a solid tumor. As shown in Fig.
6A–D, CARM1i retarded GC cell viability, induced G1 cell cycle arrest and triggered apoptosis in vitro, and the growth of xenograft tumors in vivo were also suppressed (Fig.
6E–G). Furthermore, the autophagy inhibitor HCQ, which has been proven to possess appreciable antitumor activity in clinical trials [
52,
53], demonstrated synergistic antitumor effects with CARM1i, suggesting the combination of CARM1i and HCQ as a potential therapeutic strategy.
Finally, we aimed to clarify how CARM1 regulated autophagy. The MiT/TFE family was reported to exert an essential function in autophagy regulation [
54]. We found that TFEB and TFE3 were both increased in CARM1-overexpressing cells (Fig.
7A). As TFEB has been clearly demonstrated to interact with CARM1 to promote autophagy [
26], we focused our study on whether CARM1 could induce autophagy by regulating TFE3. TFE3 helps maintain lysosomal homeostasis and promotes autolysosome formation by binding to the CLEAR elements of lysosome- and autophagy-relevant genes [
55]. The results showed that CARM1 promoted TFE3 nuclear translocation to activate autophagy, and silencing TFE3 reversed the increased cell proliferation and decreased apoptosis induced by CARM1-mediated autophagy. Then, we further explored the mechanisms by which increased CARM1 promoted the nuclear translocation of TFE3. AMPK is a responder to starvation and low energy states to maintain intracellular energy homeostasis. Previous studies have shown that AMPK is often activated during the development of GC, demonstrating a pro-tumor effect [
56,
57]. AMPK is considered to be a key signaling pathway regulating the activity of the MiT/TFE transcription factor family [
34]. It has been reported that TFE3 nuclear translocation is closely related to the activation of the AMPK-mTOR signaling pathway in the cytoplasm. As reported, MTOR complex 1 (MTORC1) is the most important regulator of TFE3 which directly phosphorylates TFE3 serine residue 321 and causes TFE3 cytoplasmic retention [
58]. MTORC1 activity is negatively regulated by AMPK through direct and indirect phosphorylation under energy deficiency [
59,
60]. Moreover, activated AMPK could directly phosphorylate TFE3 serine residues and regulate TFE3 transcriptional activity. Therefore, we investigated the cytoplasmic AMPK-mTOR pathway and found that the activated AMPK-mTOR pathway was responsible for the nuclear translocation of TFE3 (Fig.
8A). Furthermore, AMPK also regulates the MiT/TFE family through nuclear AMPK-CARM1 signaling pathway. When activated in the nucleus, AMPK phosphorylates FOXO3a and transcriptionally inhibits SKP2, thereby reducing the ubiquitination of CARM1 and increasing the protein level of CARM1 in the nucleus [
26]. CARM1 was reported to coactivate TFEB in the nucleus, and TFEB and TFE3 always shared regulatory networks. We then explored the interaction of CARM1 and TFE3 in the nucleus and proved that CARM1 could also regulate autophagy through the nuclear AMPK-CARM1-TFE3 signaling pathway (Fig.
8B). Furthermore, it has been reported that CARM1 can regulate the activation of AMPK in skeletal muscle during denervation-induced plasticity. CARM1 interacts with AMPK and causes the methylation of AMPK, resulting in AMPK activation [
61]. This result is consistent with the increased AMPK activation in CARM1-overexpressing cells in our study. These results suggest that there may be a regulatory loop between CARM1 and AMPK which forms positive feedback to promote tumor development. And the detailed mechanism and structure–function relationship are needed to be further explored.
In conclusion, the present study first demonstrated that CARM1 was upregulated in clinical GC tissues and cell lines, and that higher CARM1 expression was related to an unfavorable prognosis. More importantly, our study proved for the first time that CARM1i treatment significantly inhibited GC tumor growth both in vivo and in vitro and synergized with autophagy inhibitors, suggesting a promising therapeutic strategy. CARM1 promoted GC cell proliferation, accelerated G1-S transition and reduced ER stress-induced apoptosis by regulating autophagy. Mechanistically, CARM1 potentiated autophagy by facilitating TFE3 nuclear translocation and enhancing its activity through the activation of cytosolic AMPK-mTOR and nuclear AMPK-CARM1-TFE3 signaling pathways.
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