MICAL1 stability by PlexinA1 promotes gastric cancer cell migration

doi:10.21203/rs.3.rs-3306835/v1

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MICAL1 stability by PlexinA1 promotes gastric cancer cell migration

https://doi.org/10.21203/rs.3.rs-3306835/v1

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Background

For metastasis to proceed, tumor cells must become mobile by modulating their cytoskeleton. MICAL1 is known as an actin cytoskeleton regulator, but the mechanisms by which it drives cancer cell migration are still unclear.

Methods

Immunohistochemistry assays and western blotting were used to detect the expression levels of MICAL1 in gastric cancer tissues and cells. Immunoprecipitation and immunofluorescence staining were used to detect the interactions of proteins. Wound-healing and transwell assays were performed to demonstrate the migratory function of MICAL1 in gastric cancer cells. In addition, qPCR, bioinformatics analysis, pulldown assay, ROS detection as well as western blotting were conducted to verify the mechanism of MICAL1 in gastric cancer cell migration.

Results

Analysis of gastric cancer tissues revealed that MICAL1 are elevated in gastric cancer tissues compared with non-tumor tissues and that its high expression is predictive of poor survival. PlexinA1 and MICAL1 were directly interact with each other. Specific inhibition of PlexinA1 accelerated MICAL1 ubiquitination and proteasome-based degradation. Furthermore, PlexinA1 positively regulates MICAL1 expression via Rac1 activation and following ROS production. Functional studies confirmed that PlexinA1 and MICAL1 facilitated gastric cancer cell migration via promoting vimentin expression.

Conclusions

These results indicate that PlexinA1 is a key regulator of MICAL1 stability via a Rac1/ROS dependent manner, and MICAL1 stability may be involved in promoting vimentin expression and gastric cancer cell migration.

gastric cancer

MICAL1

PlexinA1

migration

Gastric cancer is one of the most common solid malignancy worldwide, with more than 1,000,000 new cases diagnosed and approximately 760,000 deaths in 2020(Sung et al., 2021). Metastasis promotes the development of tumors and is a significant cause of gastric cancer death. Many studies have shown that cytoskeletal rearrangement is a key event in cancer metastasis and that the members of molecules interacting with the CasL (MICALs) can promote tumor migration by regulating the cytoskeleton(Rajan et al., 2023). However, the underlying mechanisms that MICALs use to regulate gastric cancer cell migration are still unclear.

MICALs are a class of oxidation–reduction enzymes that can bind to actin filaments and promote their disassembly, contributing to cytoskeletal dynamics and intracellular trafficking(Ortegon Salas et al., 2020; Yoon et al., 2021). In mammals, the MICAL family consists of MICAL1–3, MICAL-L1 and MICAL-L2. MICALs have been shown to facilitate tumor metastasis in multiple human malignancies. For example, high cytoplasmic MICAL2 was positively associated with lymphatic metastasis and mortality rate in patients with lung adenocarcinoma(Zhou et al., 2020). MICAL-L2 potentiates gastric cancer cell migration by inhibiting EGFR degradation in lysosome(Min et al., 2019). In particular, MICALs such as MICAL-L2 were found to regulate the epithelial–mesenchymal transition (EMT) process and hence promote ovarian cancer metastasis(Zhu et al., 2015). Recently, MICAL1 silencing has been reported to be involved in reduced E-cadherin and vimentin levels, but upregulated N-cadherin, in oral squamous cell carcinoma(Grauzam et al., 2018). The potential role of MICAL1 in tumor progression, particularly in gastric cancer cell migration, needs to be elucidated.

Plexins are single transmembrane proteins that function as receptors for semaphorins. Plexins are composed of four members: PlexinA (1–4), PlexinB (1–3), PlexinC1, and PlexinD1. Unlike other single transmembrane receptors, the cytoplasmic region of plexins has a GTPase activating protein (GAP) domain that mediates intracellular signaling, which can cause reorganization of the cytoskeleton and affect some physiological processes related to cell migration(Fard and Tamagnone, 2021; Neufeld et al., 2016). Among the plexin members, high expression of PlexinA1 was found in gastric carcinoma tissues(Liu et al., 2022). Silencing of PlexinA1 interrupted JAK-STAT3 signaling and inhibited gastric cancer cell EMT induced by β2-AR(Liu et al., 2022). In diabetic mice, PlexinA1 was shown to interact with MICAL1 to mediate Sema3A-induced F-actin collapse(Aggarwal et al., 2015). The binding of PlexinA1 to MICAL1 has been proposed to relieve MICAL1’s autoinhibited conformation(Loria et al., 2015). Consequently, we hypothesized that PlexinA1 and MICAL1 might act cooperatively to regulate migration of gastric cancer cells.

The exact biological function of MICAL1 varies in different cancer types. For example, high expression of MICAL1 was found in invasive breast cancer samples which contributes to directional breast cancer cell migration and proliferation(Deng et al., 2016; Deng et al., 2018; McGarry et al., 2021); however in colorectal cancer, MICAL1 has recently been proposed as a cancer cell migration and proliferation suppressor(Gu et al., 2022). In the present study, we investigate the expression of MICAL1 in gastric cancer tissue samples by bioinformatics analysis and immunohistochemical assays. Then, we investigated the association of PlexinA1 with MICAL1 in gastric cancer cell migration and explored their underlying mechanisms of action. As far as we know, this is the first study focused on the relationship between PlexinA1 and MICAL1 in gastric cancer cell migration. These results enhance our understanding of gastric cancer cell migration and offer potential therapeutic targets for PlexinA1- or MICAL1-directed diagnostics and treatments for gastric cancer.

Ethics statement

All immunohistochemical assays involving human cancer specimens were approved by Nanjing Medical University ethics committee.

Cell culture

The normal gastric epithelial cell line GES-1, and gastric cancer cell lines SGC-7901, MGC-803, BGC-823 were purchased from the Cell Biology Institute of the Chinese Academy of Science (Shanghai, China). The cells were grown in DMEM (Hyclone, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% FBS (Gibco, Carlsbad, CA, USA) and kept at 37°C with 5% CO₂ in a humidified incubator.

Plasmids and siRNAs

The human MICAL1 cDNA were obtained from YouBio (Changsha, China). The human PlexinA1 cDNA was constructed in our lab. The siRNAs used in this study were obtained from GenePharma (Shanghai, China) as previously described(Min et al., 2020). The sequences of the siRNAs were as follows: siMICAL1 #1, 5′-AUGUCUG UGACAGAAGCUGTT-3′; siMICAL1 #2, 5′-UAAUACAGCACUGGAGGUCTT-3′; siMICAL1 #3, 5′-UAGUUAUUCAUGGUGCCUGTT-3′; siPLXNA1 #1, 5′-UUGA CGUUGUCAGUACUGCTT-3′; siPLXNA1 #2, 5′-AUGACAAAGUUGCCGUUC CTT-3′; siPLXNA1 #3, 5′-AGUUGUACUUCACCUCAGGTT-3′. Cells were transfected with plasmids and siRNA using Lipofectamine 2000 according to the manufacturer’s protocol. After 48 h, the cells were lysed and analyzed by western blotting or other assays as specified in each experiment.

Cell migration assay

Cell migration ability was examined by either wound-healing or transwell assays. For wound-healing assays, cells were seeded on 6-well plates and incubated to confluence. Scratching was made with a 10-µL plastic pipette tip. After rinsing with phosphate-buffered saline (PBS), cells were treated with the indicated stimulator for the indicated time. Images of initial gap width and the residual gap width were observed under the inverted microscope (Carl Zeiss Meditec, Jena, Germany).

To further determine the migratory capability of the cells, 24-well transwell insert with 8-µm pores were used. 1×10⁶ cells were added to the insert in serum-free medium whereas the lower well contained 10% FBS culture medium. After 24 h of incubation, the remaining cells on the upper side of the insert were removed. The chambers were fixed and stained with 0.1% crystal violet and then counted in 5 randomly selected fields under the inverted microscope.

Western blotting

Western blot analysis was performed as previously described(Wang et al., 2022). Primary antibodies used were MICAL1 (14818-1-AP, Proteintech, Wuhan, China), PLXNA1 (CBAQ0320121, R&D Systems, Minnesota, USA), GAPDH (BS72410, Bioworld, Nanjing, China), E-cadherin (20874-1-AP, Proteintech, Wuhan, China), Myc-tag (60003-2-IG, Proteintech, Wuhan, China), vimentin (60330-1-IG, Proteintech, Wuhan, China), Rac1 (24072-1-AP, Proteintech, Wuhan, China), and HA-tag (S1064-2-APand66006-2-IG, Proteintech, Wuhan, China). Secondary antibodies used were HRP-conjugated normal rabbit IgG (Jackson, Lancaster, PA, USA), HRP-conjugated normal mouse IgG (Jackson, Lancaster, PA, USA), and rabbit anti-goat IgG (Zhifan, Nanjing, China). Proteins were visualized by the enhanced chemiluminescence reagent (FuDeBio, Hang Zhou, China).

For protein stability assays, Cells were treated with cycloheximide (CHX) for 0, 4, 8, and 12 h and harvested the total proteins. Then, the proteins were analyzed by western blotting assays.

Real-time qPCR

Real-time qPCR was conducted as previously described(Min et al., 2020). The sequences of the primers used were: MICAL1: 5′-GGCAC TCGGTGCTAAGAAGTT-3′ (sense) and 5′-CCCCAGTGAATTTCCACCCC-3′ (antisense); PLXNA1: 5′-ACGAGGA GGATGCCGACAT-3′ (sense) and 5′-ACAGAGCCGCACGATCTTG-3′ (antisense); E-cadherin: 5′-ATTTTTCCCTCGACACCCGAT-3′ (sense) and 5′-TCCCAGGCGTAGACCAAGA-3′ (antisense); vimentin: 5′-AGTCCACTGAGT ACCGGAGAC-3′ (sense) and 5′-CATTTCACGCATCTGGCGTTC-3′ (antisense); GAPDH: 5′-TCGGATCAACGGATTTGGT-3′ (sense) and 5′-TTCCCGTTC TCAGCCTTGAC-3′ (antisense).

Measurement of reactive oxygen species (ROS)

Briefly, cells were plated in 6-well plates and pretreated with the indicated interfering strands, plasmids, or drugs including NAC (Beyotime, Shanghai, China), ML-099 (MCE, New Jersey, USA), IA-116 (MCE, New Jersey, USA), or NSC23766 (MCE, New Jersey, USA) for 4 h. Then, 10 µM of 2’-7’dichlorofluorescin diacetate (DCFH-DA) (Beyotime) was added to the wells and incubated for 30 min at 37°C. Then the dye was washed away and fluorescence was measured with an Olympus BX51 microscope (Olympus, Tokyo, Japan).

Immunohistochemistry

Gastric cancer surgical specimens were obtained from Outdo Biotech (Shanghai, China). Immunohistochemical analysis was described previously(Qi et al., 2021). Tissue sections were incubated with primary antibodies overnight: MICAL1 (14818-1-AP, Proteintech, Wuhan, China) and vimentin (60330-1-IG, Proteintech, Wuhan, China). After washing with PBS, the sections were incubated with the HRP-conjugated secondary antibody (Jackson, Lancaster, PA, USA) for 1 h. After counterstained with hematoxylin, sections were placed under the microscope and images were taken.

Pulldown assay

Pulldown assays of Rho GTPase were performed as described previously (Min et al., 2019). Briefly, cells were washed and lysed with lysis buffer. After centrifugation, supernatants were mixed with PAK-CRIB beads. After incubated for 30 min at 4°C, the beads were washed and the bound proteins (GTP-Rac1) were resolved by 10% SDS-PAGE and visualized by western blot analysis using the corresponding antibody. Total Rac1 was also analyzed by western blotting.

Ubiquitination assays

Ubiquitination assays were conducted as previously described(Wang et al., 2022). Briefly, after transfected with HA-ubiquitin plasmids for 48 h, cells were harvested and lysates were incubated with MICAL1 antibody at 4°C overnight, then antibody-bound affinity beads (Beyotime) were added to the mixture. After washing steps, the bound proteins were subjected to SDS-PAGE and western blotting.

Co-IP assays

In brief, the lysate was incubated with indicated antibody or IgG control antibody and gently agitated at 4°C overnight. Antibody–protein complexes were collected by adding protein A + G agarose beads (Beyotime). The bound proteins were separated by SDS-PAGE and subjected to western blotting.

Clinicopathological analysis of MICAL1 and PlexinA1

Gastric cancer data from TCGA and Kaplan–Meier plotter were downloaded to analyze MICAL1 and plexinA1 gene expressions, as well as survival data of gastric cancer patients(Lanczky and Gyorffy, 2021). Pathway analysis was performed using GO(Ashburner et al., 2000; Gene Ontology, 2021), KEGG(Kanehisa, 2019; Kanehisa et al., 2021; Kanehisa and Goto, 2000), and GSEA(Mootha et al., 2003; Subramanian et al., 2005).

Statistical analysis

The statistical analysis was performed using SPSS and data were presented as means ± standard error. The Student’s t-test was used to test for significance between two groups. One-way ANOVA was used to test for significance among more groups. Differences were considered significant when the level of P < 0.05.

Aberrant MICAL1 expression in gastric cancer

We investigated the difference in the expression of MICAL1 between normal tissues and tumor by applying gastric cancer data from the database of TCGA. As we can see from Fig. 1A, MICAL1 mRNA expression was higher in gastric cancer samples (335 cases) than that in normal samples (35 cases) from TCGA (P < 0.05). Figure 1.B showed that MICAL1 expression was greater than the matching adjacent tissue samples (n = 27). Similarly, MICAL1 expression in lymph node stage N3 samples was markedly higher than lymph node stage N0, indicating that high MICAL1 may contribute to cancer cell metastasis. Kaplan–Meier survival curve analysis indicated that patients with relatively high MICAL1 expression predicted poor survival (Fig. 1D–1F). We used an immunohistochemical method to determine MICAL1 expression in gastric cancer tissues (Fig. 1G). MICAL1 expression was found elevated in gastric cancer specimens than in paracancerous tissues. The expression level of MICAL1 was also correspondingly higher in gastric cancer cells than in normal gastric epithelium cells (Fig. 1H). These findings demonstrate that MICAL1 is highly expressed in patients of gastric cancer, indicating that it may be a potential diagnostic biomarker.

Function enrichment of MICAL1 in gastric cancer

Next, we sought to explore the putative cellular mechanisms underlying the effects of MICAL1 in gastric cancer through bioinformatic pathway analysis. As shown in Fig. 2A, the following biological processes were significantly affected: adaptive immune response, protein folding, regulation of small GTPase-mediated signal transduction, etc. Receptor complex, side of membrane were high level GO cellular component terms. GO terms under molecular function were enriched in SH3 domain binding, peptide receptor activity, etc. The KEGG terms were mainly involved in cytokine signaling pathway, cell adhesion molecules, cytokine–cytokine receptor, etc. (Fig. 2B). The results of GSEA showed that the significant association between MICAL1 and extracellular matrix function. Passive transmembrane transporter activity, immune response-regulating signaling pathway, cytokine–cytokine receptor interaction, actin cytoskeleton, cell adhesion molecules, ubiquitin ligase complex, etc. (Fig. 2C–2H). Based on these data, in order to gain a better understanding the function of MICAL1 in gastric cancer, we chose to focus on the cytokine receptor, cell adhesion, and migration pathways.

MICAL1 promotes gastric cancer cell migration

To test the role of MICAL1 in gastric cancer cell migration, we carried out MICAL1 loss-of-function assays. First, by using siMICAL1, we interfered with MICAL1 expression in MGC-803 and BGC-823 gastric cancer cells. To detect the knockdown efficiency, qPCR and western blotting were performed. As shown in Fig. 3A and 3B, the expressions of MICAL1 in the knockdown group (siMICAL1 #1 and #2) were lower than that of the control group in both cell lines. Then the effect of MICAL1 on cell migration was checked by wound healing and transwell assays. The results indicated that cells migration abilities of MGC-803 and BGC-823 cells were inhibited effectively by the knockdown of MICAL1 (Fig. 3C–3F). Contrarily, the overexpression of MICAL1 in SGC-7901 cells (Fig. 3G and 3H) led to increased protein levels of vimentin but decreased E-cadherin in gastric cancer cells; however, vimentin and E-cadherin mRNA levels were not altered (Fig. 3I and 3J). Meanwhile, MICAL1 overexpression markedly increased SGC-7901 cell migration (Fig. 3K and 3L).

The formation of the PlexinA1/MICAL1 complex

PlexinA receptor, the receptor required for repulsive axon guidance, was identified as MICAL1 interacting protein previously (Terman et al., 2002). In order to elucidate mechanisms of MICAL1 accumulation in gastric cancer cells, the correlation analysis between PlexinA1 and MICAL1 was investigated firstly. It revealed that PlexinA1 correlated positively with MICAL1 (Fig. 4A). We then tested whether PlexinA1 modulates gastric cancer progression. We noticed that PlexinA1 expression was greatly higher in tumor tissues versus normal or matched-adjacent samples (Fig. 4B and 4C, Fig. S1A). Kaplan–Meier survival curve analysis showed that patients with high PlexinA1 levels had significantly poorer prognosis, when compared with patients with low PlexinA1 accumulation (Fig. 4D–4F, Fig. S1B). Bioinformatics analysis showed that PlexinA1 had good discriminative power for stomach adenocarcinoma (STAD) samples (Fig. S1C) and it was related to EMT, cell adhesion, and cell migration processes (Fig. 4G and 4H). The expression level of PlexinA1 was also increased in gastric cancer cells than in normal gastric epithelium cells (Fig. S1D).

Further to this, we explored whether PlexinA1 co-localized with MICAL1 in gastric cancer cells. Immunofluorescence staining results indicated that they occurred at the same cell position (Fig. 4I and 4J). The direct interaction between PlexinA1 and MICAL1 was confirmed by Co-IP assays (Fig. 4K). Based on their known domains, we subsequently divided MICAL1 into fragments and individually expressed them in Cos-7 cells. The results showed that the FAD + CH + LIM domain region of MICAL1 was required for its interaction with PlexinA1 (Fig. S2).

PlexinA1 regulates MICAL1 stability by attenuating MICAL1 degradation

Gastric cancer cells were transfected with siPlexinA1, with the knockdown efficiency determined using qPCR and western blotting (Fig. 5A and 5B). As measured by transwell and wound-healing assays, we noticed that transfection with siPlexinA1 effectively impaired the migration ability of those cancer cells (Fig. 5C and 5D). As expected, transfection with siPlexinA1 in BGC-823 and MGC-803 cells reduced MICAL1 and vimentin expression of protein levels but did not affect their mRNA levels significantly (Fig. 5E–5H). We also found that after ectopic expression of PlexinA1 in SGC-7901 cells, both MICAL1 and vimentin protein expression, but not their mRNA expression (Fig. 5I and 5J), as well as cell migratory potential were increased (Fig. 5K and 5L). PlexinA1 overexpression-increased migration of gastric cancer cells are partially dampened by MICAL1 knockdown (Fig. 5M), indicating that PlexinA1 positively regulates the migratory potential of gastric cancer cells though MICAL1.

Since transfection with siPlexinA1 had no effect on MICAL1 mRNA levels, to further explore the relationship between PlexinA1 and MICAL1, we knocked down PlexinA1 and examined MICAL1 protein expression after pretreatment of those cells with CHX, an inhibitor of protein synthesis. The results showed that knockdown of PlexinA1 enhanced MICAL1 degradation when compared to control cells (Fig. 6A). To determine which signaling pathway is involved in the process of MICAL1 degradation, cells were treated with various inhibitors, including AICAR (macroautophagy inhibitor), chloroquine (lysosomal proteolysis inhibitor), bortezomib (proteasome inhibitor), MG-132 (proteasome inhibitor). The results indicated that only the proteasome inhibitors bortezomib and MG-132 blocked PlexinA1 silencing-enhanced MICAL1 degradation (Fig. 6B–6E), suggesting PlexinA1 may inhibit MICAL1 degradation by suppressing a proteasome-related machanism.

Next, we examined MICAL1 ubiquitination levels in PlexinA1-depleted cells. As shown in Fig. 6F and 6G, although total cellular protein ubiquitination levels had no significant change, the ubiquitination intensity of MICAL1 was increased in siPlexinA1-transfected BGC-823 cells, implying that PlexinA1 promotes the ubiquitination, and consequently proteasome-based degradation.

PlexinA1 regulates MICAL1 stability by activating Rac1/ROS

To investigate the mechanism through which PlexinA1 regulates MICAL1 stability, we checked Rac1 activity using pulldown assays. When transfected with siPlexinA1in BGC-823 cells or transfected with PlexinA1 plasmids in SGC-7901 cells, we found that Rac1 activity was significantly decreased by PlexinA1 knockdown and enhanced by overexpression of PlexinA1 (Fig. 7A and 7B). MICAL1 expression also reduced when BGC-823 cells were over-expressing Rac1-T17N, a constitutively inactive mutant of Rac1 (Fig. 7C). Pretreatment with ML-99, a Rac1 GTPase activator, reversed PlexinA1 silencing-induced MICAL1 downregulation (Fig. 7D), while pretreatment with NSC23766, a Rac1 GTPase inhibitor, reversed PlexinA1 overexpression-induced MICAL1 upregulation (Fig. 7E). Co-IP assays revealed that PlexinA1 bind to less MICAL1 when BGC-823 cells were pretreated with IA-116, another inhibitor of Rac1 (Fig. 7F). Together, these evidences indicate that PlexinA1 positively regulates MICAL1 protein contents though Rac1 activation.

Rac1 was reported to play an key role in ROS production(Acevedo and Gonzalez-Billault, 2018). To ascertain whether PlexinA1 had similar promoting effects on ROS production, we assessed the effects of both PlexinA1 knockdown and its overexpression on ROS production. As expected, we noticed that ROS production was significantly impaired by PlexinA1 knockdown and increased by PlexinA1 overexpression (Fig. 7G and 7H). To ascertain whether PlexinA1 promote ROS production though Rac1, PlexinA1-knockdown BGC-823 were pretreated with ML-99, and PlexinA1-overexpressed SGC-7901 were pretreated with IA-116. As expect, pretreatment with ML-99 increased ROS production in PlexinA1-depleted BGC-823 cells (Fig. 7I), while pretreatment with IA-116 decreased ROS production in PlexinA1-overexpressed SGC-7901 cells (Fig. 7J). These data imply that ROS production by PlexinA1 occurs at least in part via Rac1 activation. Furthermore, pretreatment with ROS scavenger NAC decreased MICAL1 protein contents in PlexinA1-overexpressed SGC-7901 cells (Fig. 7K), even in PlexinA1-depleted BGC-823 cells (Fig. 7L). Together, these findings demonstrate that PlexinA1 promotes MICAL1 stability though Rac1/ROS-dependent manner.

The effect of PlexinA1/MICAL1 on vimentin expression

When MGC-803 and BGC-823 cells underwent a significant reduction of PlexinA1 or MICAL1, an obvious decrease in the contents of vimentin protein, but not contents of its mRNA, was noticed (Fig. 8A–8D). Immunofluorescence staining analysis also confirmed that the membrane /cytoplasmic protein levels of vimentin was decreased in MICAL1-silenced BGC-823 cells compared with the control group (Fig. 8E). Histological examination also revealed that vimentin staining in tumor samples was increased appreciably compared to paracancerous samples (Fig. 8F). In addition, immunostaining of MICAL1 and vimentin in the clinic samples displayed a positive correlation in terms of expression (R² = 0.28, P < 0.0001) (Fig. 8G). Together, the results above suggest that PlexinA1 and MICAL1 promotes gastric cancer cell migration, at least in part, by inducing expression of vimentin protein.

In summary, we report that MICAL1 functions as an oncogene in migration of gastric cancer cells. Mechanistically, PlexinA1 is a key regulator of MICAL1 stability via a Rac1/ROS dependent manner, and MICAL1 stability may be involved in promoting vimentin expression and gastric cancer cell migration (Fig. 8H).

MICAL1 is an important monooxygenase that participates in regulating cytoskeletal rearrangement(Vitali et al., 2016; Wu et al., 2018; Zucchini et al., 2011). Recently, the potential widespread function of MICAL1 on cancer pathophysiological processes has attracted increasing attention. Studies have indicated that in experimental models, for example, MICAL1 promotes the malignant development of breast tumors(McGarry et al., 2021), MICAL1 has been perceived as being markers for renal clear cell adenocarcinoma(Yang et al., 2022). Our previous study showed that hypoxia induced MICAL1 expression in gastric cancer(Zhao et al., 2019). In this study, we revealed a new link between PlexinA1 and MICAL1 in positive regulation of gastric cancer cell migration, that is PlexinA1 maintains MICAL1 expression by Rac1/ROS dependent manner, thereby enhancing vimentin expression and cancer cell migratory ability.

The results in this study showed high MICAL1 expression in gastric cancer tissues, and increased expression of MICAL1 was correlated with poor prognosis, indicating a close relation between MICAL1 and gastric cancer progression. The most novel finding was that the regulation of MICAL1 expression was strongly dependent on PlexinA1. MICAL1 was identified as an effector that took part in PlexinA1 signaling pathway for axon guidance(Jahan et al., 2022). MICAL1 can also mediate sema3a-induced F-actin collapse, thereby regulating shape changes of podocyte in diabetic mice(Aggarwal et al., 2015). Recently, the role of PlexinA1 in tumor progression has gained increasing interest from researchers. Pro-oncogenic effects of PlexinA1 were noted in certain tumors such as brain and gastric (Jacob et al., 2016; Lu et al., 2017). In the present study, we found that gastric cancer cell lines or patient samples expressed higher PlexinA1 than normal samples, and that tumors with high PlexinA1-expression are correlated with worse-survival rates. This oncogenic effect of PlexinA1 in gastric cancer was consistent with previous results from Yamada et al., who reported that PlexinA1 positively related with mRNA expression levels of proliferation-marker genes and lung cancer cell proliferation(Yamada et al., 2016). Consistent with the report that PlexinA1 is required for neural crest migration(Wagner et al., 2010), our findings revealed that PlexinA1 accelerates gastric cancer cell migration by stabilizing MICAL1 expression. Interestingly, no obvious alteration in MICAL1 mRNA levels were found after silencing of PlexinA1, suggesting that PlexinA1 might not change MICAL1 expression at transcriptional level. Then, we focused on revealing the mechanisms through which PlexinA1-induced stabilization of MICAL1.

MICAL1 consists of N-terminal FAD domain, followed by CH, LIM, and C-terminal CC domains. Normally, MICAL1 is autoinhibited by its intramolecular interactions between the CC domain and the CH and LIM domains. Physical interactions between MICAL1 and proteins such as plexins and PAK1 may spatiotemporally relieve MICAL1 autoinhibition(Schmidt et al., 2008). Here, PlexinA1 overexpression was seen in gastric cancers. Silencing of PlexinA1 prevents MICAL1 expression and gastric cancer cell migration. Moreover, we observed that PlexinA1 interacted with MICAL1 in gastric cancer cells. Differently from the suggestion that the cytoplasmic portion of plexins bind to the CC domain of MICALs(Rajan et al., 2023), we found that PlexinA1 binds to the FAD + CH + LIM domains, but not the CC domain of MICAL1. Previous studies have shown that protein degradation can be regulated by two possible mechanisms: the proteasome-based or lysosomal-based proteolytic pathway. Our results demonstrate that only proteasome inhibitors could prevent the reduction in MICAL1 degradation by PlexinA1 knockdown. In addition, MICAL1 ubiquitination levels were increased in gastric cancer cells transfected with siPlexinA1. Together, these results indicate that the knockdown of PlexinA1 promotes the ubiquitination, and consequently proteasome-based degradation of MICAL1.

It has been found that the intracellular domain of plexins binds to the small GTPase Rac1(Zhang and Buck, 2017). This study was designed to explore whether PlexinA1 could also regulate Rac1 activation. We noticed that knockdown of PlexinA1 apparently suppressed Rac1 activation. Conversely, the effects were reversed by PlexinA1 overexpression, implying that PlexinA1 maintains Rac1 activation. When we further analyzed the mechanism about how PlexinA1 maintain of MICAL1 stability, we observed that activating Rac1 significantly abrogated the downregulation of MICAL1 in PlexinA1-depleted cells. Meanwhile, inhibiting Rac1 exerted the opposite effect, suggesting that PlexinA1 maintains MICAL1 stability through Rac1 activation. A recent study showed that Rac1 directly activates NADPH oxidases to produce ROS, which have been implicated in modulating breast cancer cell migratory properties(Tobar et al., 2008). ROS also account for cancer cell genomic instability, proliferation, angiogenesis, and resistance to apoptosis(Morry et al., 2017). Here, silencing of PlexinA1 significantly decreased ROS production, and ROS scavenger NAC suppressed the high levels of MICAL1 induced by the overexpression of PlexinA1 or increased activation of Rac1. Thus, it is likely that PlexinA1 maintains MICAL1 stability in a Rac1/ROS-dependent manner. Combined with the role of ROS in activating PAK1 phosphorylation(Yang et al., 2011), and phosphorylated PAK1 involves in relieving MICAL autoinhibition by phosphorylating serine 817 and 960 residues within the C-terminus of MICALs(McGarry et al., 2022), it can be suggested that Rac1 acts downstream of PlexinA1 to regulate MICAL1 stability and normal functions via ROS-dependent PAK1 activation. The mechanisms by which PlexinA1 regulates MICAL1 stability remain to be elucidated in detail.

It is widely known that vimentin expressions are associated with EMT, which is known to enhance cell migration and invasion(Nowak and Bednarek, 2021; Usman et al., 2021). Here, we found that knockdown of PlexinA1 and MICAL1 induced a significant reduction in vimentin expression as well as a impaired cell migratory ability; in contrast, both PlexinA1 and MICAL1 overexpression activated vimentin expression, consequently promoting cell migratory ability. MICAL1 has been reported to associate with vimentin through its C-terminal region(Suzuki et al., 2002), which implies an intimate connection between MICAL1 and vimentin. As an intermediate filament protein, vimentin is well-known to play a pivotal role during cell migration, including forming filaments to modulate focal adhesion, generating directional migratory force, and modulating genes for EMT inducers(Usman et al., 2021). Our study is the first, to our knowledge, demonstrated that PlexinA1 and MICAL1 contributes to gastric cancer cell migration, at least partially through maintaining vimentin expression.

In conclusion, PlexinA1 and MICAL1 regulates gastric cancer cell migration by inducing vimentin expression. PlexinA1 binds to and stabilizes MICAL1 expression in a Rac1/ROS-dependent manner, avoiding ubiquitination and following degradation of MICAL1. Although our results indicate that PlexinA1 and MICAL1 may serve as new therapeutic targets in gastric cancer field due to its potential to restrain tumor migration, their clinical evaluation needs to be investigated in the future.

Ethics approval and consent to participate

All methods were carried out in accordance with the Declaration of Helsinki. No ethics approval was required for this work. All utilized public data sets were generated by others who obtained ethical approval.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Funding

This work was supported by the National Natural Science Foundation of China (82073226, 81773107) to Jun Du.

Author contributions

JD conceived and designed the study. FY and TX performed the experiments and acquired the data. YY, QW, WZ, PM, YZ, and YW performed the experiments and acquired the data. JD performed the bioinformatics analysis. FY and JD interpreted and analyzed the data. FY and JD wrote the manuscript.

Acknowledgements

Not applicable.

Availability of data and materials

The datasets supporting the conclusions of this article are included within the article.

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No competing interests reported.

S1.jpg
Figure S1. The expression of PLXNA family members in STAD samples. (A) The mRNA expression of PLXNA family members in STAD samples. (B) OS for PLXNA members in STAD patients by Kaplan–Meier analysis. (C) The ROC curve for PLXNA1. (D) Expressions of PLXNA1 protein in GES-1 and different types of gastric cancer cell lines. * P < 0.05, ** P < 0.01.
S2.jpg
Figure S2. PLXNA1 interacts with MICAL1 and its truncation mutants. A schematic representation of human MICAL1 domains. COS-7 cells were co-transfected with Myc-PLXNA1 and MICAL1 domain mutants, then the cell extracts were examined.

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MICAL1 stability by PlexinA1 promotes gastric cancer cell migration

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Abstract

Background

Methods

Results

Conclusions

Figures

Introduction

Materials and methods

Ethics statement

Cell culture

Plasmids and siRNAs

Cell migration assay

Western blotting

Real-time qPCR

Measurement of reactive oxygen species (ROS)

Immunohistochemistry

Pulldown assay

Ubiquitination assays

Co-IP assays

Clinicopathological analysis of MICAL1 and PlexinA1

Statistical analysis

Results

Aberrant MICAL1 expression in gastric cancer

Function enrichment of MICAL1 in gastric cancer

MICAL1 promotes gastric cancer cell migration

The formation of the PlexinA1/MICAL1 complex

PlexinA1 regulates MICAL1 stability by attenuating MICAL1 degradation

PlexinA1 regulates MICAL1 stability by activating Rac1/ROS

The effect of PlexinA1/MICAL1 on vimentin expression

Discussion

Declarations

References

Additional Declarations

Supplementary Files

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