The Ubiquitin-like Protein FAT10 Stabilizes eEF1A1 Expression to Promote Tumor Proliferation in a Complex Manner

Human HLA-F adjacent transcript 10 (FAT10) belongs to a class of ubiquitin-like proteins (UBL) that have a protein sequence and three-dimensional core structure similar to that of ubiquitin (Ub; refs. 1, 2). The researches have indicated that FAT10 is involved in various essential cellular development processes, including immune-mediated inflammation, apoptosis, the cell cycle, and the regulation of cell proliferation (3–6). In recent years, the relationship between FAT10 and tumors has attracted wide attention, and FAT10 has been found to be closely involved in tumor development (7). Studies have shown that FAT10 plays a significant role in the regulation of cell proliferation and high FAT10 expression can promote the proliferation of cancer cells (3, 8), but the mechanism remains unclear. In addition, studies of FAT10 have focused on protein degradation (9–11). Recently, we for the first time showed that FAT10 stabilized β-catenin expression by antagonizing its ubiquitination (12), but the mechanism of this process remains to be clarified. In particular, FAT10–substrate complexes are directly degraded by the 26S proteasome when FAT10 degrades substrates (13). However, it is not known whether FAT10–substrate complexes can be degraded by the 26S proteasome when FAT10 stabilizes substrates. Thus, in-depth research on these will be necessary and useful to further understand FAT10 function.

Eukaryotic translation elongation factor 1A1 (eEF1A1) is an important protein that is involved in protein translation elongation, and eEF1A1 has the effect of triggering the initiation of translation elongation (14, 15). eEF1A1 is a highly conserved GTP-binding protein in eukaryotic cells that plays a significant role in peptide chain extension during protein synthesis (14). Research indicates that eEF1A1 plays an important role in the regulation of various biological characteristics of cells, such as the cell cycle, cell growth, and cell death (16, 17). Studies have shown that eEF1A1 is aberrantly upregulated in many tumor tissues (18, 19). Overexpression of eEF1A1 is closely related to cancer cell proliferation, downregulating eEF1A1 expression inhibits cell proliferation in many tumors (18, 19). Our previous studies showed that eEF1A1 is a substrate of FAT10 and that the downregulation of FAT10 expression reduces eEF1A1 expression in human hepatocellular carcinoma (HCC) cells (20). However, the mechanism of this process is unclear, and it remains unknown how FAT10 impacts and regulates the expression of eEF1A1 to affect the proliferation of tumor cells.

In this study, our results showed that the protein expression levels of FAT10 and eEF1A1 in cancer cells are higher than those in normal epithelial cells. FAT10 can stabilize eEF1A1 expression and thus influence cancer cell proliferation. We subsequently explored the mechanism by which FAT10 stabilizes eEF1A1. Our results showed that FAT10 and Ub compete with one another to bind to eEF1A1 to form FAT10–eEF1A1 and Ub–eEF1A1 complexes, respectively. Overexpression of FAT10 results in a simultaneous reduction of Ub–eEF1A1 complexes and increase of FAT10–eEF1A1 complexes. Finally, we demonstrated the mechanism by which FAT10 stabilizes eEF1A1 in a complex manner.

The HEK293T, HL-7702, and human cancer cell lines SW620, MDA-MB231, PANC-1, Huh-7, HepG2, Hep3B, HeLa, and MHCC97H were purchased from the Shanghai Cell Bank, Type Culture Collection Committee of Chinese Academy of Science (Shanghai, China). All these cells were authenticated using short tandem repeat profiling by the Cell Bank. After resuscitation, only cells within 6 months were used for this study. The cells were cultured in DMEM or MEM (Gibco) supplemented with FBS (Gibco) to a final concentration of 10% and were exposed to antibiotics at 37°C with 5% CO₂. Plasmids and reagents are included in the Supplementary Materials and Methods. The identification of the interference effects are also included in the Supplementary Materials and Methods (Supplementary Fig. S1).

qRT-PCR, confocal microscopy, coimmunoprecipitation (Co-IP) and in vivo ubiquitination assay were performed as previously described (12). The specific primers used for PCR amplification are provided in Supplementary Table S1.

The cells were incubated with 5-ethynyl-20-deoxyuridine (EdU; Ribobio) for 5 hours, and processed according to the manufacturer's instruction. After three washes with PBS, the cells were treated with 300 μL of 1×Apollo reaction cocktail for 30 minutes. Then, the DNA contents of the cells in each well were stained with 100 μL of Hoechst 33342 (5 μg/mL) for 30 minutes and visualized under a fluorescence microscope.

The CCK-8 assay was used to measure cell viability. Exponentially growing cancer cells (100 μL, 5 × 10⁴ cells/mL) were seeded into 96-well plates. The plates were then incubated at 37°C and 5% CO₂ for 48 hours. Subsequently, the cells were washed and 10 μL of CCK-8 (Beyotime Institute of Biotechnology, China) solution was added to each well; after 1.5 hours, the absorbance was measured. The optical density was measured at an absorbance of 450 nm using an iMark microplate reader (Bio-Rad). For each group, mean values of the mean absorbance rates from five wells were calculated.

For in vivo tumorigenicity assays, 1 × 10⁶ cells in 100 μL of PBS were injected subcutaneously into the flanks of nude mice (Shanghai SLAC Laboratory Animal Co., Ltd.). Tumor size was measured with a caliper (calculated volume = shortest diameter/2 × longest diameter/2) at 5-day intervals. After 40 days, photographs were taken, and tumors were harvested and individually weighed after the mice were anesthetized. The data are presented as tumor weight (mean ± SD). The animal work was approved by the Ethics Committee for Animal Experiments of the Second Affiliated Hospital of Nanchang University.

shRNA-resistant FAT10 plasmids (nonsense mutation FAT10 gene expression vector) were constructed by site-directed mutagenesis using a QuickChange Mutagenesis Kit (Agilent Technologies). The plasmid pcDNA3.1(+)-FAT10 was used as a template. HA-eEF1A1 mut plasmids were constructed by site-directed mutagenesis using a Multipoints Mutagenesis Kit (TaKaRa). The plasmid pcDNA3.1-HA-eEF1A1 was used as a template. The mutagenic primers are provided in Supplementary Table S1. The mutant alleles were confirmed by sequencing.

GST, GST–Rpn10 proteins were expressed in Escherichia coli strain BL21 as described previously. Glutathione Sepharose 4B beads (sigma) were incubated with GST, GST–Rpn10 along with purified His-FAT10 (Enzo) or His-Ub (upstate) in incubation buffer (20 mmol/L Tris-HCl, pH 7.4, 0.1% Triton X-100) overnight at 4°C and washed five times with incubation buffer. For other experiments, HA-eEF1A1 were in vitro transcribed and translated using a TNT T7 coupled reticulocyte lysate system (Promega) and purified GST and GST–FAT10 bound on the GSH beads was added. Proteins were eluted by boiling at 95°C for 5 minutes with SDS buffer, resolved by SDS-PAGE, and subjected to Western blot analysis.

For the deficiency of endogenous FAT10 in HEK293T cells, we used the Cas9 design target tool (http://crispr.mit.edu) and choose two of the effective sequences. To design the specific target sequences for sgRNA synthesis, the primers were: FAT10#1 forward (5′ to 3′) CACC gcatgtccgttccgaggaat; FAT10#1 reverse (5′ to 3′) AAAC attcctcggaacggacatgc; FAT10#2 forward (5′ to 3′) CACC gcaatgatcgagactaagac; FAT10#2 reverse (5′ to 3′) AAAC gtcttagtctcgatcattgc, and then the pairs of annealed oligos was ligated into the pX459 (Addgene Plasmid #48139) after being digested with BbsI(Thermo). HEK293T cells were transfected with 2 μg of total plasmid DNA per well in 6-well plates using lipofectamine 3000 (Invitrogen) according to the manufacturer's instructions.

After being transfected with CRISPR-Cas9 plasmids for 2 days, the HEK293T cells were treated with puromycin in 1 μg/mL for 72 hours to enrich for transfected cells. The surviving cells were sorted into single clones into the 96-well plate. The viable clones were grown larger about 1 week and picked up 80% of cells per well for DNA isolation and genotyping. The genomic PCR primers for identifying genotyping of FAT10 knockout (KO) cells were: reverse (5′ to 3′) ctagaggaccagatagatag; forward (5′ to 3′) cctccaatacaataacatgc. The remaining 20% of cells were cultured for 8 to 10 days, and clones propagated from single cell were picked out. The KO efficacy of FAT10 KO HEK293T cells was confirmed by both Western blotting and DNA sequencing.

All data were analyzed using SPSS 19.0 (SPSS, Inc.). The results are presented as the mean ± SEM of three independent experiments. The differences between groups were analyzed using Student t test when two groups were compared or by one-way ANOVA when more than two groups were compared. Furthermore, univariate and multivariate analyses were performed using the logistic regression model. The test results were considered significant at P < 0.05.

Our previous study found that the downregulation of FAT10 expression could reduce eEF1A1 expression in HCC cells (20). To further investigate whether FAT10 regulates eEF1A1 expression in other cancer cells, we first examined the levels of FAT10 and eEF1A1 by Western blot analysis in a variety of cancer cell lines (i.e., HCC, breast cancer, colorectal cancer, pancreatic cancer, and cervical cancer). The results showed that the protein expression levels of FAT10 and eEF1A1 in cancer cells were higher than the levels in normal epithelial cells (Fig. 1A). Subsequently, Western blot analysis and qRT-PCR analyses indicated that the downregulation of FAT10 significantly reduced the protein expression of eEF1A1 in Huh7, SW620, MDA-MB231, PANC-1, and MHCC97H cells. However, the expression of eEF1A1 mRNA was not affected by the downregulation of FAT10 (Fig. 1B). Moreover, MTT and EDU proliferation assays showed that the cell-proliferation capacity of cancer cells with stable FAT10 interference was significantly lower compared to the control group (P < 0.01; Fig. 1C). A tumorigenicity assay in nude mice also indicated that the downregulation of FAT10 significantly inhibited tumor growth (P < 0.01; Fig. 1D). To rule out possible off-target effects associated with shRNAs, we generated a FAT10 cDNA harboring silent mutations in the shRNA-targeting sequence that made the mRNA insensitive to this shRNA (Supplementary Fig. S2A). Rescue experiments showed that shRNA-resistant-FAT10 significantly restored and stabilized the expression of FAT10 in Huh7 and MDA-MB231 cells with FAT10 interference, and the expression of eEF1A1 was enhanced. Moreover, cell proliferation was also restored (P < 0.01; Supplementary Fig. S2B–D). These results suggest that FAT10 regulates eEF1A1 protein expression and influences cancer cell proliferation.

To further validate that FAT10 mediates cancer cell proliferation by regulating eEF1A1, we first increased the expression of eEF1A1 in FAT10 knockdown cancer cells and then analyzed the FAT10 and eEF1A1 protein expression levels and cell-proliferation abilities using Western blots and proliferation assays. Our results showed that the downregulation of FAT10 decreased eEF1A1 expression, whereas the upregulation of eEF1A1 attenuated the loss of eEF1A1 expression in FAT10 knockdown Huh7 cells (Fig. 2A). The experiment further showed that the knockdown of FAT10 dramatically decreased the proliferative ability of Huh7 cells, whereas the upregulation of eEF1A1 rescued the decreased proliferative ability induced by FAT10 knockdown (P < 0.01; Fig. 2B and C). Then, we decreased the expression of eEF1A1 in FAT10-overexpressing HeLa cells and then analyzed FAT10 and eEF1A1 protein levels and cell-proliferation ability. Western blot analysis results showed that overexpression of FAT10 significantly increased the expression of eEF1A1, whereas the knockdown of eEF1A1 expression dramatically inhibited the increase of eEF1A1 expression induced by FAT10 in HeLa cells (Fig. 2D). In addition, the downregulation of eEF1A1 significantly reduced FAT10-enhanced cell proliferation (P < 0.01; Fig. 2E and F).

Finally, to further demonstrate that FAT10 regulates eEF1A1 expression, we constructed FAT10-KO HEK293T cells using the CRISPR-Cas9 system (Fig. 2G; refs. 21, 22). Study has demonstrated that IFNγ/TNFα could mediate the induction of FAT0 expression (11), so we added IFNγ/TNFα to HEK293T cells and FAT10-KO HEK293T cells to investigate the expression of FAT10 and eEF1A1. The increased expression of eEF1A1 was accompanied by FAT10 overexpression in HEK293T cells. However, in FAT10-KO HEK293T cells, FAT10 expression was not detected, whereas the expression of eEF1A1 was unchanged (Fig. 2H). Thus, the above experiments further confirmed that FAT10 could regulate the expression of eEF1A1 and eEF1A1 was key for FAT10-mediated cancer cell proliferation.

Our previous study confirmed a mutual interaction between eEF1A1 and FAT10 (20). The research has shown that FAT10 modifies substrates by covalently binding to lysine sites on the substrates (23). We first used HEK 293T cells to determine which sites in eEF1A1 binding to FAT10. eEF1A1 was divided into various structural domains, C-terminally truncated eEF1A1 mutants and truncated eEF1A1 fragments (24, 25). Co-IP experiments indicated that the eEF1A1 region in aa 284-332 and aa 394-462 contains the FAT10 interaction region (Fig. 3A and D). The 14 lysines in the binding region of eEF1A1 were mutated to arginine (Fig. 3E), and the various mutant plasmids were constructed. Co-IP experiments revealed that the binding of eEF1A1 to FAT10 was weakened with an increase in the number of mutated sites. The binding of eEF1A1 to FAT10 disappeared when all 14 of the sites were mutated (Fig. 3F). Taken together, these results demonstrate that the binding of FAT10 with eEF1A1 occurs through the 14 lysine sites located in two regions of eEF1A1 (284-332 and 394-462).

To further explore the role of the lysine binding sites of FAT10 and eEF1A1 in the regulation of eEF1A1 function, Huh7 cells were simultaneously transiently transfected with an shFAT10 plasmid and an HA-eEF1A1 wt or HA-eEF1A1 14mut plasmid. The results revealed that in the wild-type eEF1A1 group, when FAT10 expression was reduced, the level of eEF1A1 protein was lower, and tumor cell proliferation also declined. However, in the eEF1A1 mutant group, the eEF1A1 protein expression level and tumor cell proliferation were not significantly changed (Fig. 3G and H). These results demonstrated that the abrogation of the FAT10–eEF1A1 interaction influences tumor cell proliferation.

Research has suggested that eEF1A1 is degraded by the ubiquitin–proteasome system (UPS) and that its degradation also needs the lysine sites combined with Ub (26). However, the exact lysine residues were not specified. First, we determined that the degradation of eEF1A1 protein in Huh7 and Hep3B cells occurs via the UPS. Then, we identified the interaction between eEF1A1 and Ub by Co-IP and confocal microscopy in Huh7 and Hep3B cells (Fig. 4A and Supplementary Fig. S3A). Subsequently, we further explored which lysine residues in eEF1A1 interacted with Ub. Interestingly, our study provides convincing evidence that the lysines of eEF1A1 that bind to Ub are the same as the lysines of eEF1A1 that bind to FAT10 in HEK293T cells (Fig. 4B–E and Supplementary Fig. S3B and S3C). In addition, Co-IP results and the functional studies further verified that Ub binds to eEF1A1 to influence eEF1A1 function (Fig. 4F). Finally, our results further confirmed that FAT10 and Ub bind to the same lysine residues on eEF1A1 (Fig. 4G).

The above experiments revealed that FAT10 and Ub bind to the same lysine sites on eEF1A1. We further found that FAT10 and Ub compete with each other to bind to eEF1A1 in Huh7 and HEK 293T cells by GST pulldown assays (Fig. 4H and Supplementary Fig. S3D). Moreover, our research showed that when increasing amount of FAT10 plasmid were added, the level of FAT10–eEF1A1 complexes gradually increased, and the level of Ub–eEF1A1 complexes gradually decreased. In contrast, as the amount of Ub increased, the level of FAT10–eEF1A1 complexes gradually decreased, and the level of Ub–eEF1A1 complexes gradually increased in Huh7 and HEK293T cells (Fig. 4I and Supplementary Fig. S3E). These experiments demonstrated that FAT10 and Ub bind competitively with eEF1A1, and overexpression FAT10 led to increase in FAT10–eEF1A1 complexes and decrease in Ub–eEF1A1 complexes.

Our previous study showed that FAT10 regulates β-catenin expression by antagonizing its ubiquitination (12). Thus, we speculated that FAT10 might also regulate eEF1A1 expression by antagonizing its ubiquitination. As expected, our results confirmed this. First, we found that FAT10 was involved in the degradation process of eEF1A1 in Huh7 and MDA-MB231 cells (Fig. 5A and B and Supplementary Fig. S4A and S4B). Second, the degradation dynamics assay showed that the half-life of the ectopically expressed eEF1A1 significantly increased in the FAT10-overexpressing cells compared with that in the control cells (P < 0.01; Fig. 5C and D and Supplementary Fig. S4C and D). Third, Co-IP and in vivo ubiquitination assay results revealed that the downregulation of FAT10 increased the ubiquitination level of eEF1A1, and that FAT10 overexpression decreased the ubiquitination level of eEF1A1 in Huh7 and MDA-MB231 cells (Fig. 5E and F and Supplementary Fig. S4E and S4F). Finally, in FAT10-KO HEK293T cells, Co-IP results showed that the ubiquitination level of eEF1A1 was unchanged when IFNγ/TNFα was added. However, with the restoration of FAT10 expression with HA-FAT10 in FAT10-KO HEK293T cells, the ubiquitination level of eEF1A1 decreased significantly (Fig. 5G).

The ubiquitination level of substrate is determined by the expression relationship between UBLs and Ub–substrate complexes, and in turn determines the expression level of substrate (27–31). Our data showed that overexpression FAT10 led to decrease in Ub–eEF1A1 complexes. Hence, we speculated that FAT10 antagonizing eEF1A1 ubiquitination is due to the decrease of Ub–eEF1A1 complexes. First, we investigated the expression of eEF1A1 with an increase in Ub–eEF1A1 complexes. The results showed that following the increase in exogenous ubiquitin, Ub–eEF1A1 complexes increased and endogenous eEF1A1 proteins decreased in a dose-dependent manner in HEK293T cells (Fig. 5H). Second, the expression of eEF1A1 was investigated when Ub–eEF1A1 complexes were reduced by induced FAT10 overexpression. The results showed that as Ub–eEF1A1 complexes decreased, eEF1A1 expression was enhanced following FAT10 overexpression in HEK293T cells (Fig. 5I). Finally, we investigated the expression of eEF1A1 when Ub–eEF1A1 complexes were unchanged. Unlike wild-type (WT) FAT10 HEK293T cells, the induction of IFNγ/TNFα in FAT10-KO HEK293T cells did not cause FAT10 expression. Without a change in Ub–eEF1A1 complexes, the expression of eEF1A1 also remained unchanged. However, the restoration of FAT10 expression with the Flag-FAT10 plasmid in FAT10-KO HEK293T cells reduced Ub–eEF1A1 complexes and increased eEF1A1 expression (Fig. 5J and K). Hence, we suggest that FAT10 antagonizing eEF1A1 ubiquitination for increasing its expression is because a reduction in Ub–eEF1A1 complexes.

In addition, our study found that FAT10 and Ub bind competitively with eEF1A1 to form Ub–eEF1A1 and FAT10–eEF1A1 complexes, respectively. Researches have reported that FAT10–p62 complexes are directly degraded by proteasome when FAT10 degraded p62 (1, 11, 23). Thus, we constructed HA-FAT10–p62 and HA-FAT10–eEF1A1 fusion plasmids. FAT10–p62 complex is took as a positive control to observer whether FAT10–eEF1A1 complexes are degraded by the proteasome. In FAT10-KO HEK293T cells, our results showed that HA-FAT10–p62 expression significantly increased with MG132. However, the expression of HA-FAT10–eEF1A1 remained unchanged irrespective of MG132 (P < 0.01; Fig. 6A). These results indicated that HA-FAT10–p62 was degraded by the proteasome, whereas HA-FAT10–eEF1A1 was not. In addition, research has reported that when FAT10 degrades substrates, FAT10 can be recognized by the only FAT10 receptor Rpn10 within 26S proteasome, leading to the degradation of FAT10–substrate complexes. Thus, a reduction in Rpn10 leads to the accumulation of FAT10–substrate complexes (32). To further confirm that FAT10–eEF1A1 could not be degraded by the proteasome, our study first demonstrated that Rpn10 could bind with FAT10 (Fig. 6B and C). Then, we investigated the degradation rates of FAT10–p62 and FAT10–eEF1A1 in response to altered expression levels of Rpn10 in FAT10-KO HEK293T cells. The results showed that the half-life of ectopically expressed HA-FAT10-p62 decreased significantly in Rpn10-overexpressing cells compared with control cells. The inhibition of Rpn10 expression significantly reduced HA-FAT10–p62 degradation rate. However, altering Rpn10 had no effect on the degradation rate of HA-FAT10–eEF1A1 (P < 0.01; Fig. 6D). Finally, we observed whether Rpn10 expression affects the accumulation of FAT10–p62 and FAT10–eEF1A1 in HEK293T cells. Compared to HEK293T cells expressing normal levels of Ub, the Co-IP and Western blot analysis results in HEK293T cells expressing low levels of Ub showed reduced expression of Rpn10 led to increase in FAT10–p62 accumulation and p62 expression. In contrast, when the expression of Rpn10 increased, FAT10–p62 accumulation and p62 expression decreased. However, altering Rpn10 did not affect the accumulation of FAT10–eEF1A1, and the expression of eEF1A1 remained unchanged (Fig. 6E). Hence, these results demonstrated that FAT10–eEF1A1 complexes are not degraded by the proteasome.

Finally, we further observed the relationship of the FAT10–eEF1A1 complexes and eEF1A1 expression in HEK293T cells. First, Co-IP and Western blotting showed that when FAT10 expression decreased, the FAT10–eEF1A1 complexes and eEF1A1 expression also decreased. In contrast, with a gradual increase in FAT10 expression, the expression of FAT10–eEF1A1 and eEF1A1 complexes also increased in a dose-dependent manner (Fig. 6F). Moreover, we observed the expression of FAT10–eEF1A1 complexes and eEF1A1 when the influence of Ub on eEF1A1 degradation was excluded. Our results showed that with unchanged Ub–eEF1A1 complexes, the expression of eEF1A1 increased with the increase in FAT10–eEF1A1 complexes by inducing FAT10 expression (Fig. 6G and H). In addition, to further verify this phenomenon, FAT10-KO HEK293T cells expressing low levels of Ub were treated with IFNγ/TNFα. Under the circumstance of unchanged Ub–eEF1A1 complexes, FAT10–eEF1A1 complexes were not detected, and the expression of eEF1A1 remained unchanged. However, owing to the restoration of FAT10 expression by transfection with the Flag-FAT10 plasmid, FAT10–eEF1A1 complexes increased, as did the expression of eEF1A1 (Fig. 6I and J). Thus, our results demonstrated that the expression of eEF1A1 also increased with the increase in FAT10–eEF1A1 complexes.

Tumor proliferation plays an important role in the tumorigenesis and development of tumor (33–35). To understand the molecular mechanism of tumor proliferation will contribute to the prevention and treatment of tumor (36, 37). eEF1A1 is a protein that triggers the initiation of protein translation elongation, and involved in the control of various cellular processes (16, 18). Studies have shown that eEF1A1 is high expression in many tumors, and overexpression of eEF1A1 promotes cancer cell proliferation and cancer development (16, 18). In this study, our data also found that eEF1A1 is upregulated in a variety of cancer cells, including HCC, breast cancer, colorectal cancer, pancreatic cancer, and cervical cancer. In addition, we also found that downregulation of eEF1A1 could significantly reduce cancer cell proliferation ability in vitro and in vivo. Study has shown that blood POZ containing gene type 2 (BPOZ-2) directly binds to eEF1A1 to regulate eEF1A1 expression (38). Our previous study found that FAT10 also could bind to eEF1A1 and influence eEF1A1 expression in HCC cells (20). Here, our further study found that the expression levels of FAT10 and eEF1A1 were upregulated in cancer cells. FAT10 inhibition can reduce eEF1A1 expression and decrease the cancer cell proliferation. Moreover, the upregulation of eEF1A1 rescued the decreased cell proliferation induced by FAT10 knockdown, whereas eEF1A1 inhibition significantly decreased FAT10-enhanced cell proliferation. Finally, we used FAT10-KO HEK293T to demonstrate that FAT10 could regulate the expression of eEF1A1. Therefore, we reveal a novel mechanism by which FAT10 promotes cancer cell proliferation by upregulating eEF1A1 expression.

In addition, we further investigated the mechanism by which FAT10 regulates the eEF1A1 expression. In this study, we found that FAT10 could stabilize the eEF1A1 by antagonizing its ubiquitination. Generally, UBL stabilizing substrate expression is due to that UBL could compete with Ub to bind with the same lysine of substrate to form Ub–substrate complexes and UBL–substrate complexes respectively, and overexpression UBL leads to decrease Ub–substrate complexes, antagonizing substrate ubiquitination for increasing substrate expression (27, 31, 39–42). FAT10 also belongs to UBLs and can target proteins for degradation (23, 43). Interestingly, we recently found that FAT10 could stabilize β-catenin by antagonizing its ubiquitination. The knockdown of FAT10 could increase levels of β-catenin ubiquitination, whereas increased FAT10 reduced the levels of β-catenin ubiquitination (12). But the detail process by which FAT10 antagonizs substrate ubiquitination remains unclear. For example, it is not clear whether FAT10 could compete with Ub to bind to the same lysine sites of substrate to form Ub–substrate complexes and FAT10–substrate complexes, respectively. It is also not known whether FAT10 antagonizing substrate ubiquitination is caused by a decrease in Ub–substrate complexes. Here, we investigate the mechanism of FAT10 antagonizing eEF1A1 ubiquitination by these problems in detail. Our results showed that the binding of FAT10 and Ub with eEF1A1 occurred through the same lysine sites of eEF1A1. FAT10 and Ub competitively bound to eEF1A1 to form Ub–eEF1A1 complexes and FAT10–eEF1A1 complexes, respectively. FAT10 overexpression resulted in a decrease in Ub–eEF1A1 complexes. Finally, we further found that FAT10 overexpression led to a decrease in Ub–eEF1A1 complexes, the ubiquitination level of eEF1A1 was decreased and eEF1A1 expression was enhanced. Thus, we demonstrate that FAT10 antagonizes eEF1A1 ubiquitination for increasing its expression through decreasing Ub–eEF1A1 complexes.

Another important question is that FAT10–substrate complexes are directly degraded by the 26S proteasome when FAT10 degrades substrates (44), but it is not known whether FAT10–eEF1A1 complexes can be degraded by the 26S proteasome when FAT10 stabilizes eEF1A1. The research has reported that FAT10 is a unique UBL that mediates proteasome degradation independently of Ub, and attaches to substrate, forming a FAT10–substrate complexes for recognition by the only FAT10 receptor Rpn10 and degradation by the 26S proteasome (9, 13). Hence, in the following experiments, we further focused on whether FAT10–eEF1A1 complexes are degraded by the proteasome. Surprisingly, our results demonstrated that FAT10–eEF1A1 complexes are not degraded by the proteasome. This conclusion is based on the following observations. First, the expression of FAT10–eEF1A1 was not affected by the presence or absence of MG132, which indicated that FAT10–eEF1A1 was not degraded by the proteasome. Second, our data further found that altering Rpn10 expression had no effect on the degradation rate and accumulation of FAT10–eEF1A1. Interestingly, we then further found that the expression of eEF1A1 also increased with the increase in FAT10–eEF1A1 complexes. In conclusion, our results show that FAT10 competes with Ub for binding to eEF1A1 to form FAT10–eEF1A1 and Ub–eEF1A1 complexes, respectively. Overexpression of FAT10 resulted in a simultaneous reduction in Ub–eEF1A1 complexes and an increase in FAT10–eEF1A1 complexes. It is the mechanism by which FAT10 stabilizes eEF1A1 in a complex manner. On one hand, FAT10 inhibits eEF1A1 ubiquitination through reduced Ub–eEF1A1 complexes. On the other hand, eEF1A1 expression increases with the increase of FAT10–eEF1A1 complexes, which are not degraded by the proteasome.

In summary, we demonstrated that FAT10 could promote cancer cell proliferation by stabilizing eEF1A1 expression. Further study found that FAT10 stabilizes eEF1A1 in a complex manner (Figure 7). These findings may propose a new mechanism in which FAT10 stabilizes substrate. In future, in-depth studies on what makes FAT10–eEF1A1 complexes resistant to degradation by proteasomes are necessary. Such efforts would provide a new perspective on biological functions of FAT10 and yield novel insights into the regulation of protein.

No potential conflicts of interest were disclosed.

Conception and design: X. Liu, L. Chen, J. Ge, J. Shao

Development of methodology: X. Liu, L. Chen, J. Ge, C. Yan, Z. Huang, J. Hu, C. Wen, M. Li, D. Huang, J. Shao

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): X. Liu, L. Chen, J. Ge, C. Yan, Z. Huang, J. Hu, Y. Qiu, H. Hao, R. Yuan, J. Lei, X. Yu, J. Shao

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): X. Liu, L. Chen, J. Ge, C. Yan, Z. Huang, J. Hu, C. Wen, M. Li, D. Huang, J. Shao

Writing, review, and/or revision of the manuscript: X. Liu, L. Chen, J. Shao

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): X. Liu, J. Ge, C. Yan, Y. Qiu, H. Hao, R. Yuan, J. Lei, X. Yu, J. Shao

Study supervision: J. Shao

This work was supported by grants from the National Natural Science Foundation of China (81060196 and 81360325), the Project of Jiangxi Provincial Department of Science and Technology (20121BBG70029, 20133BBG70031, and 20152ACB20020), the Project of Jiangxi Provincial Department of Education (KJLD12053).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.