New Results

HuR -dependent SOD2 protein synthesis is an early adaptation of ovarian cancer cells in response to anchorage independence

Yeon Soo Kim, Jaclyn E. Welles, Priscilla W. Tang, Zaineb Javed, Amal T. Elhaw, Karthikeyan Mythreye, Scot R. Kimball, Nadine Hempel
doi: https://doi.org/10.1101/2021.02.19.432010

Abstract

The presence of malignant ascites is a common feature of advanced stage ovarian cancer. During metastasis, as cells detach from the tumor and extravasate into the peritoneal fluid, ovarian cancer cells must adapt to survive the loss of anchorage support and evade anoikis. An important pro-survival adaptation in this context is the ability of tumor cells to increase their antioxidant capacity and restore cellular redox balance. We previously showed that the mitochondrial superoxide dismutase SOD2 is necessary for ovarian cancer cell anoikis resistance, anchorage-independent survival and spheroid formation, and intraperitoneal spread in vivo. We now demonstrate that the upregulation of SOD2 protein expression is an early event initiated in response to anchorage independence and occurs at the post-transcriptional level. SOD2 protein synthesis is rapidly induced in the cytosol within 2 hours of matrix detachment. Polyribosome profiling demonstrates an increase in the number of ribosomes bound to SOD2 mRNA, indicating an increase in SOD2 translation in response to anchorage-independence. Mechanistically, we find that anchorage-independence induces cytosolic accumulation of the RNA binding protein HuR/ELAVL1, leads to HuR binding to SOD2 mRNA, and that the presence of HuR is necessary for the increase in SOD2 mRNA association with the heavy polyribosome fraction and SOD2 protein synthesis. Cellular detachment activates the stress-response protein kinase p38 MAPK, which is necessary for HuR-SOD2 mRNA binding and the rapid increase in SOD2 protein expression. Moreover, HuR is necessary for optimal cell survival during early stages of anchorage independence. These findings uncover a novel post-transcriptional stress response mechanism by which tumor cells are able to rapidly increase their mitochondrial antioxidant capacity to adapt to stress associated with anchorage-independence.

Introduction

Despite significant improvements in our understanding of the genomic and RNA expression signatures of ovarian cancer, the 5-year survival rate of patients has remained relatively low (Torre et al, 2018). Epithelial ovarian cancer (EOC), which accounts for 90% of ovarian cancer cases, remains the most deadly gynecological malignancy, partly due to the asymptomatic progression to advanced disease. About 80% of patients with high grade serous adenocarcinoma, the most common subtype of EOC, are diagnosed at stage III and stage IV. At this stage the 5-year survival rate remains less than 30% (Torre et al., 2018), and is characterized by significant tumor spread throughout the peritoneal cavity and the accumulation of malignant ascites (Ahmed & Stenvers, 2013). This peritoneal fluid is thought to further facilitate transcoelomic metastasis of ovarian cancer by providing a medium in which extravasating cells disseminate throughout the peritoneal cavity where they eventually invade the mesothelial wall of the peritoneum, and neighboring organs including the omentum, liver, intestines, and pleural fluid.

Disseminated ovarian tumor cells undergo stress adaptation to survive in non-adherent conditions. One adaptation to evading anchorage-independent cell death, known as anoikis, is by increased antioxidant capacity to reduce the detachment-induced surges in reactive oxygen species (ROS) (Jiang et al, 2016; Schafer et al, 2009). In vivo studies have demonstrated that increased antioxidant enzyme expression and small molecule antioxidant treatment promotes the metastatic spread of melanoma and breast cancer cells (Davison et al, 2013; Piskounova et al, 2015), suggesting that the maintenance of redox homeostasis is a key adaptation during metastasis. Similarly, we have shown that ovarian cancer cells increase their mitochondrial antioxidant capacity after matrix detachment, by upregulating the expression and activity of the deacetylase sirtuin 3 (SIRT3), and its target protein the mitochondrial superoxide dismutase SOD2 (Kim et al, 2020). Our previous work demonstrated that SIRT3-dependent SOD2 deacetylation is necessary for SOD2 activity and that upregulation of both SIRT3 and SOD2 are necessary for anoikis resistance and in vivo transcoelomic spread of ovarian cancer cells (Kim et al., 2020).

SOD2 is a nuclear encoded mitochondrial protein that is responsive to stress-activated transcriptional regulation (Kim et al, 2017). An extensively studied stress-response transcription factor implicated in the regulation of antioxidant responses of tumor cells is Nrf2/NFE2L2, which has been implicated with the increases in SOD2 expression observed in breast cancer and clear cell ovarian carcinomas (Hart et al, 2016; Hemachandra et al, 2015; Konstantinopoulos et al, 2011). SOD2 transcription is also induced by the sirtuin regulated transcription factor Foxo3A (Kenny et al, 2017), and by NF-κB, which has also been implicated in the upregulation of SOD2 transcription in response to matrix detachment of breast cancer cells (Kamarajugadda et al, 2013). Although much emphasis has been placed on the SIRT3-dependent regulation of SOD2 activity and stress response pathways leading to SOD2 transcription in cancer, whether translational regulation contributes to SOD2 expression in tumor cells has not been investigated in depth.

Posttranscriptional and translational regulatory mechanisms are crucial for fine-tuning of gene expression. In particular, the interplay between mRNAs, miRNAs, and RNA-binding proteins has been implicated in cancer development and metastasis (Audic & Hartley, 2004; van Kouwenhove et al, 2011; Wurth & Gebauer, 2015). Among RNA binding proteins, HuR exerts many effects such as increased mRNA stability and translational efficiency by binding to the AU-and U-rich elements (AREs) in the 3’ UTR of target mRNAs (Abdelmohsen & Gorospe, 2010). Following activation HuR translocates from the nucleus to the cytoplasm, where it binds and regulates mRNAs that encode proteins involved in oncogenic signaling pathways (Epis et al, 2011; Mazan-Mamczarz et al, 2008), anti-apoptosis (Filippova et al, 2011), cell cycle (Lal et al, 2014; Wang et al, 2000b), and chemoresistance (Raspaglio et al, 2010), all of which converge on accelerated tumorigenesis and aggressive phenotypes (Wang et al, 2013). Importantly, HuR can translocate to the cytoplasm upon genotoxic or environmental stimuli in cancer cells (Lafarga et al, 2009; Lal et al., 2014), which suggests that HuR-dependent translation may be a critical stress adaptation utilized by cancer cells. HuR expression analyses across different malignancies including ovarian cancer showed that its cytoplasmic accumulation is correlated with advanced stages and poor prognosis (Denkert et al, 2004a; Denkert et al, 2004b; Miyata et al, 2013; Mrena et al, 2005). However, it is not known if HuR is involved in ovarian cancer metastatic spread, or specifically upregulated as an adaptation to anchorage independent stress.

Interestingly, a transcriptome-wide RNA-binding analysis identified multiple HuR binding sites in the 3’ UTR of SOD2 mRNA (Lebedeva et al, 2011), but the functional consequences of these sites related to SOD2 mRNA translation remain elusive. Given that HuR is a stress responsive RNA binding protein promoting cell survival, we investigated if SOD2 mRNA is a target of HuR in tumor cells. In the present work, we show for the first time that SOD2 mRNA is a target of HuR in ovarian cancer and that the interaction of HuR with SOD2 mRNA is required for rapid de novo SOD2 protein synthesis after matrix detachment in a p38 MAPK dependent manner. Our study provides evidence for a novel mechanism of SOD2 regulation in response to acute stress associated with anchorage-independence. It demonstrates that tumor cells are able to increase their antioxidant defenses at multiple levels beyond transcription, allowing for rapid adaptations to stress associated with different stages of tumor metastasis.

Results

SOD2 protein expression increases rapidly in response to anchorage independence

In previous work, we demonstrated that SOD2 activity increases in ovarian cancer cells within 2 hours in ultra-low attachment (ULA) cell culture conditions, which is followed by increased transcription within 24 hours in anchorage-independence (Kim et al., 2020). In addition to the rapid increase in SOD2 superoxide dismutase activity, which we reported to be dependent on SIRT3 (Kim et al., 2020), it was noted that the cytosolic SOD2 protein pool rapidly increases following detachment (Fig 1A). Subcellular fractionation demonstrated an average 4.7-fold increase in OVCA433 cytosolic SOD2 expression after 0.5 hour of cell detachment compared to attached cells, while a 1.5-fold increase was observed after 2 hours in anchorage-independent conditions in OVCAR10 cells (Fig 1A&B). In OVCA433 anchorage-independent cultures SOD2 mRNA increases trailed the changes in SOD2 protein expression, suggesting that the rapid rise in SOD2 protein following detachment is likely independent of increases in transcription in this cell line (Fig 1C, Supp Fig 3B). To determine if the observed increase in cytosolic SOD2 represents a newly synthesized SOD2 protein pool, cells were treated with the protein synthesis inhibitor cycloheximide, which decreased total SOD2 protein levels under anchorage independence (Fig 1D). 35S-Met/Cys incorporation assays demonstrated a global increase in protein synthesis immediately following detachment (Suppl. Fig 1A), and subsequent immunoprecipitation of SOD2 confirmed enhanced synthesis of SOD2 protein in anchorage independence. After 2 hours of incubation, ovarian cancer cells demonstrated 1.8-fold (OVCA433) and 2.4-fold (OVCAR10) increases in 35S-Met/Cys incorporation into the SOD2 protein compared to attached conditions, which was abrogated by cycloheximide.

Figure 1.
Figure 1.

A. The cytosolic SOD2 protein pool increases rapidly in response to anchorage-independence (a-i), compared to attached culture conditions (A). Cells were maintained for indicated times in ULA plates and SOD2 protein expression assessed following cellular fractionation and immunoblotting.

B. Fold change in SOD2 cytosolic protein expression in response to anchorage-independent (a-i) culture was quantified using densitometry, normalized to β-tubulin loading control and expressed relative to attached (A) culture conditions (n=4, one-way ANOVA, OVCA433 P=0.0015, OVCAR10 P=0.0744, Dunnett’s multiple comparison test *P<0.05; **P<0.01).

C. Fold change in SOD2 mRNA in response to short term anchorage-independent culture was assessed using semi-quantitative real time RT-PCR (n=3-4, one-way ANOVA, OVCA433 P=0.0069, OVCAR10 P=0.2946, Dunnett’s multiple comparison test *P<0.05; **P<0.01).

D. Total SOD2 protein levels were assessed by immunoblotting in response to culture in anchorage-independent conditions and protein synthesis inhibited by cycloheximide (CHX, 20 µg/mL; n=4, one-way ANOVA, P<0.0001, Tukey’s multiple comparison test *P<0.05; **P<0.01).

E35S-Met/Cys incorporation assay followed by SOD2 IP (Suppl. Fig 1B&C), demonstrates increased 35S-Met/Cys incorporation into SOD2 under anchorage independence compared to attached cells, which is abrogated in the presence of cycloheximide (n=4, one-way ANOVA, OVCA433 P<0.0001, OVCAR10 P=0.0057, Tukey’s multiple comparison test **P<0.01; ****P<0.0001).

To confirm that the increases in SOD2 protein expression are due to de novo protein synthesis, ribosome-mediated mRNA translation was assessed using polyribosome profiling. Following centrifugation, sucrose gradients were separated into four fractions and RNA was isolated from each fraction. Fraction 1 contained mRNAs not associated with ribosomes, fraction 2 contained mRNAs associated with one or two ribosomes, fraction 3 contained mRNAs associated with 3-6 ribosomes (referred to hereafter as ‘light polysomes’), and fraction 4 contained mRNAs associated with >6 ribosomes (referred to as ‘heavy polysomes’; Fig 2A). In attached conditions, SOD2 mRNA was primarily found in fractions 2 and 3 (Fig 2B&C), suggesting that SOD2 is translated at a constitutive level in this condition, which is evident by our ability to readily detect SOD2 protein by western blotting. In anchorage independent conditions the relative proportion of SOD2 mRNA shifted to fractions 3 and 4. In particular, anchorage independent cells showed a significant shift towards an enrichment of SOD2 mRNA in the heavy polyribosome fraction 4, demonstrating a larger number of ribosomal units associated with the SOD2 mRNA and an increase in SOD2 mRNA translation in anchorage independent conditions. As a point of comparison, the mRNA of the nutrient stress response protein ATF4 also shifted into fraction 4 in response to anchorage-independence (Supp Fig 2).

Figure 2.
Figure 2.

A. Polyribosome profiling was carried out after OVCA433 cells were cultured in attached (A) and anchorage independent (a-i) conditions (0.5 h) and analyzed by sucrose density gradient centrifugation. Four fractions were collected as indicated, and RNA extracted.

B. Polyribosome profiling demonstrates an increase in the percentage of SOD2 mRNA in the heavy polysomal fraction 4 in response to anchorage independence. Representative image of SOD2 RT-PCR from RNA isolated from each polysomal fraction.

C. Quantification of relative SOD2 mRNA levels in each fraction demonstrates increased proportion of SOD2 in fraction 4 following culture in anchorage independent conditions (n=3; t-test, **P<0.01).

HuR accumulates in the cytosol and binds SOD2 mRNA in response to anchorage-independence

Regulation of gene expression at the translational level is mediated by the interplay between mRNAs and RNA binding proteins. HuR (encoded by the gene ELAVL1) is a major RNA binding protein that has been implicated with alternative splicing, mRNA stability, and translation during stress conditions (Akaike et al, 2014; Lafarga et al., 2009; Lal et al., 2014). HuR recognizes and binds to AU-, U-rich elements in target mRNA transcripts. Screening of publicly available transcriptome-wide data sets analyzing HuR RNA binding by RNA immunoprecipitation sequencing (RIP-seq; ENCODE: ENCSR000CWW, ENCSR000CWZ) and photoactivatable ribonucleoside-enhanced crosslinking and immunoprecipitation (PAR-CLIP; GSE29943) revealed that the SOD2 mRNA contains multiple binding sites for HuR (Fig 3A) (Davis et al, 2018; EncodeProjectConsortium, 2012; Lebedeva et al., 2011). Several clusters of HuR binding were identified in the SOD2 3’ UTR within 3.5 kb downstream of the STOP codon (Fig 3A). While the 5’ UTR of SOD2 is less than 75 bp in length, the complete SOD2 3’ UTR spans 13,424 bp (Fig 3A, Variant 1: NM_000636). SOD2 transcripts with variable 3’ UTR lengths have previously been reported (Suppl Fig 3A) (Chaudhuri et al, 2012; Church, 1990). Using RT-PCR we confirmed that OVCA433 and OVCAR10 cells express the longer 3.4 kb 3’ UTR containing the majority of HuR sites identified (Suppl Fig 3B).

Figure 3.
Figure 3.

A. HuR/ELAVL1 binding profiles on the SOD2 mRNA was assessed using ENCODE RIP-seq data sets ENCSR000CWW and ENCSR000CWZ, and PAR-CLIP data set GSE29943.

B. HuR accumulates in the cytosol in response to anchorage-independence (n=4, one-way ANOVA, OVCA433 P<0.0001, OVCAR10 P=0.0248, Dunnett’s multiple comparison test **P<0.01; ***P<0.001).

C. Anchorage-independence induces HuR binding to SOD2 mRNA, as assessed by Ribonucleoprotein Immunoprecipitation and SOD2 RT-PCR following OVCA433 culture in attached or anchorage independent conditions (a-i, 0.5h).

D. HuR knock-down does not affect SOD2 mRNA stability in attached or anchorage-independent conditions, as determined by Actinomycin D treatment (n=4; two-way ANOVA: ns). HuR knock-down was assessed by semi quantitative real time RT-PCR (t-test, ****P<0.0001).

To examine if HuR regulates SOD2 protein expression in response to anchorage independence, cytosolic translocation of HuR in response to culture in ULA plates was first determined. Concurrent with the increases in cytosolic SOD2 protein expression (Fig 1A), HuR cytosolic protein levels increased significantly in OVCA433 within 0.5 hours of anchorage independence and within 2 hours in OVCAR10 cells (Fig 3B). Based on this finding, we next investigated if HuR binds to SOD2 mRNA in anchorage independent conditions using ribonucleoprotein immunoprecipitation to capture the HuR bound mRNAs using HuR antibody (Fig 3C). While SOD2 mRNA could not be detected following HuR IP from attached cells, SOD2 mRNA was readily identified by PCR in HuR immunoprecipitates from both OVCA433 (Fig 3C) and OVCAR10 cells (Supp Fig 3C) under anchorage independent conditions, indicating that matrix detachment causes the binding of HuR to SOD2 mRNA.

Since HuR binds to SOD2 mRNA shortly after matrix detachment, we investigated the functional consequences of the HuR-SOD2 mRNA interaction using siRNA mediated knockdown of HuR/ELAVL1 (Fig 3D & Fig 4A). An established function of HuR as a stress response RNA binding protein is its role in mRNA stabilization within the cytosol (Filippova et al., 2011; Jakstaite et al, 2015). To determine if HuR has an effect on SOD2 mRNA stability, we treated ovarian cancer cells with the transcription inhibitor actinomycin D. Compared to attached conditions, anchorage independence did not significantly alter SOD2 mRNA stability in OVCA433 cells (Fig 3D), while decreased SOD2 mRNA stability in anchorage independence was observed in OVCAR10 cells compared to attached conditions (Supp Fig 3D, two-way ANOVA, P=0.0104), indicating that these cells differ in mechanisms regulating SOD2 mRNA stability. However, HuR knockdown did not significantly alter SOD2 mRNA levels in response to actinomycin D treatment in anchorage independent or attached culture conditions (Fig 3D & Supp Fig 3D), suggesting that increased binding of HuR to SOD2 mRNA does not influence SOD2 mRNA stability.

Figure 4.
Figure 4.

A. HuR/ELAVL1 knock-down abrogates increases in cytosolic SOD2 expression in short term anchorage-independence (a-i, OVCA433 0.5 h; OVCAR10 2 h) compared to attached cultures (A; n=3-4, one-way ANOVA, OVCA433 P=0.012, OVCAR10 P=0.0001; Tukey’s multiple comparison test *P<0.05, **P<0.01, ****P<0.0001).

B. Polysome profiles of OVCA433 cells cultured in attached (A) and anchorage independent (a-i, 0.5 h) conditions following siRNA-mediated HuR/ELAVL1 knockdown.

C. HuR knock-down abrogates a shift of SOD2 mRNA into fraction 4 in response to anchorage independence (a-i). Representative image of SOD2 RT-PCR from polyribosome fractions and quantification of relative SOD2 mRNA levels in each fraction shown (n=3; t-test, *P<0.05).

HuR enhances SOD2 mRNA translation under anchorage independence

We next tested if HuR is necessary for enhanced SOD2 mRNA translation in anchorage independence. Following siRNA-mediated HuR (ELAVL1) knockdown the increase in SOD2 cytosolic protein levels induced by matrix detachment were significantly decreased (Fig 4A). To further demonstrate that increased SOD2 protein synthesis in anchorage independent ovarian cancer cells is HuR dependent, we conducted polyribosome profiling following siRNA mediated HuR knock-down (Fig 4B). In response to culture in anchorage independent conditions, SOD2 mRNA shifted towards the heavy polyribosome fraction (fraction 4) in OVCA433 cells transfected with a scramble control siRNA (Fig 4C), as demonstrated above in un-transfected cells (Fig 2). HuR knockdown abrogated this shift of SOD2 mRNA to the heavy polyribosomal fraction, and anchorage independent cultured cells lacking HuR displayed a similar distribution of SOD2 mRNA in polysomal fractions compared to attached cells (Fig 4C). There was no difference in SOD2 mRNA abundance in the subpolysome fractions (fractions 1 & 2) following HuR knock-down, indicating that a loss of HuR does not lead to a complete loss of SOD2 mRNA translation, and suggests that the primary function of HuR is to enhance SOD2 translation in response to anchorage independence, boosting SOD2 protein levels under these conditions.

Inhibition of p38 MAPK activation in response to anchorage independence abrogates increases in SOD2 protein expression and HuR-SOD2 mRNA binding

HuR can be activated in response to cellular stress via the p38 MAPK stress response kinase pathway (Tran et al, 2003; Wang et al., 2000b). An increase in p38 MAPK phosphorylation was previously reported in ovarian cancer cell lines cultured in anchorage independence for 24-48 h (Carduner et al, 2014). We were able to show that short-term anchorage independence (0.5-2 h) also increased p38 MAPK phosphorylation in OVCA433 and OVCAR10 cell lines (Fig 5A). To determine if the p38 MAPK pathway is involved in the observed increases in cytosolic SOD2 protein expression during this time, cells were treated with the p38 MAPK inhibitor, SB203580. SB203580 inhibited the phosphorylation of the p38 target MAPKAPK2 and abrogated the increases in SOD2 protein expression observed in anchorage independent conditions (Fig. 5B). In addition, the formation of the HuR-SOD2 mRNA complex was monitored in the presence of p38 MAPK inhibition. Similar to Fig 3, anchorage independent conditions increased SOD2 mRNA binding to HuR, while treatment with SB203580 decreased this interaction (Fig. 5C). The above demonstrates a link between p38 MAPK signaling, HuR binding to the SOD2 mRNA and SOD2 expression in response to cellular detachment. p38 MAPK has previously been shown to phosphorylate Thr 118 of HuR (Lafarga et al., 2009; Liao et al, 2011). In the absence of a commercially available phospho-Thr118 HuR specific antibody we were unable to successfully demonstrate that anchorage independence or p38 MAPK inhibition influences phosphorylation of HuR using HuR IP and a pan phospho-Thr antibody (data not shown). In addition, we analyzed the effects of p38 MAPK on HuR cellular localization in matrix detached cells, as this had previously been shown as a mechanism of HuR regulation in response to stress (Slone et al, 2016; Tran et al., 2003; Wang et al., 2000b). In OVCAR10 cells, p38 MAPK inhibition decreased cytosolic HuR accumulation in response to anchorage-independence, while this could not be consistently observed in OVCA433 cells (Fig. 5D). Although it is beyond the scope of the present work, the precise mechanism by which p38 MAPK activates HuR require further attention.

Figure 5.
Figure 5.

A. p38 MAPK (Thr180/Tyr182) phosphorylation is induced in response to culture in anchorage-independent culture conditions (a-i OVCA433 0.5 h, OVCAR10 2 h; n=4, T-test, **P<0.01, ****P<0.0001).

B. p38 MAPK inhibition abrogates a-i induced increases in SOD2 expression (n=3, one-way ANOVA P<0.0001, Tukey’s multiple comparison test **P<0.01, ****P<0.0001).

C. p38 MAPK inhibition abrogates HuR binding to SOD2 mRNA in anchorage-independence, as assessed by RNA immunoprecipitation.

D. Effects of p38 MAPK inhibition on cytosolic HuR levels (n=4-5, one-way ANOVA, OVCA433 P=0.0053, OVCAR10 P=0.0221, Tukey’s multiple comparison test **P<0.01, ****P<0.0001).

The above data demonstrate a novel role for HuR in the rapid upregulation of SOD2 translation in response to detachment of ovarian cancer cells. We previously reported that SOD2 is necessary for survival of cells in anchorage independence (Kim et al., 2020). Similarly, HuR knock-down resulted in an increase in the dead cell fraction of OVCA433 cells cultured for 2 h in anchorage independent conditions (Fig 6). These data demonstrate that HuR contributes to cellular survival in early stages of detachment.

Figure 6.
Figure 6.

HuR/ELAVL1 knock-down increases the dead cell fraction of OVCA433 cells when cultured in anchorage-independence for 2 h. Cells were stained with Ethidium homodimer (dead cells) and Calcein AM (live cells) and fractions of live and dead cells quantified (scale bar = 100μm; n=6; T-test **P=0.004).

Discussion

Recent studies have highlighted that tumor cells need an adequate antioxidant system to deal with intrinsic and extrinsic increases in ROS associated with metastatic progression (Davison et al., 2013; Kim et al., 2020; Piskounova et al., 2015). Tumor cells must therefore readily adapt to increase their antioxidant capacity at the transcriptional and post-transcriptional levels. In line with these findings, we previously showed that SIRT3-mediated deacetylation of SOD2 drives transcoelomic metastasis by increasing mitochondrial antioxidant capacity in anchorage-independent ovarian cancer cells (Kim et al., 2020). The present work demonstrates an additional SOD2 regulatory mechanism during early-stage anchorage independence. We found that ovarian cancer cells rapidly adapt to detachment by increasing SOD2 mRNA translation in a p38 MAPK-HuR-dependent manner.

Aberrant HuR expression has been reported in several malignancies including ovarian cancer, and an increase in cytoplasmic subcellular localization, where HuR acts to enhance translation, is linked with worse prognosis (Denkert et al., 2004a; Denkert et al., 2004b; Miyata et al., 2013). HuR’s pro-tumorigenic function in the cytoplasm involves selective stabilization and/or increased translation of target mRNAs. Previously identified HuR targets not only include mRNAs encoding pro-survival and anti-apoptotic proteins, such as Bcl-2 (Filippova et al., 2011; Ishimaru et al, 2009; Wang et al, 2000a), but also angiogenic factors and proteins that support invasion and metastasis, such as VEGF (Levy et al, 1998; Tran et al., 2003). Similar to our findings (Fig 6), HuR knock-down decreased glioma cell survival in anchorage independence (Filippova et al., 2011). It was found that HuR knock-down increased apoptosis and decreased Bcl-2 mRNA stability and protein expression, and the investigators demonstrated that HuR has the ability to bind the 3’ UTR of Bcl-2 family members (Filippova et al., 2011). Moreover, HuR regulation can interplay with miRNAs to further fine tune expression in cancer, as has been demonstrated in ovarian cancer with miR-200c (Prislei et al, 2013). This continuously growing repertoire of cancer-related mRNAs regulated by HuR further emphasizes the critical role of this RNA binding protein in cancer cells. Moreover, HuR appears to confer survival advantages to cancer cells by rapidly adjusting gene expression for stress adaptation and resistance to cell death. Our data here identify SOD2, an important antioxidant enzyme for maintaining mitochondrial redox homeostasis, as a novel HuR target during early-stage metastasis.

HuR is a predominantly nuclear protein which translocates to the cytoplasm upon extrinsic or intrinsic stimuli and stress signals. Posttranslational modifications of HuR by different signaling pathways have been shown to affect its RNA binding affinity, nucleo-cytoplasmic shuttling, and the stability of HuR protein depending on the location of residues (Abdelmohsen & Gorospe, 2010). In particular, phosphorylation events are directly involved in spatiotemporal regulation of HuR. Among different kinases activated during stress, p38 MAPK-dependent phosphorylation on Thr118 induces cytoplasmic accumulation of HuR and increased p21 mRNA binding after exposure to ionizing radiation (Lafarga et al., 2009) and enhanced mRNA binding upon IL-1β treatment (Liao et al., 2011). Consistent with these previous findings, we found that stress associated with matrix detachment activated p38 MAPK (Fig 5). Importantly, activation of the p38 MAPK pathway increased SOD2 cytosolic protein expression under anchorage independence and we found that the association of HuR with SOD2 mRNA was also p38 MAPK-dependent (Fig 2 & 5). It remains to be determined whether HuR is phosphorylated on Thr118 in anchorage independent cells, or if p38 MAPK indirectly activates HuR to bind SOD2 mRNA. Moreover, cytosolic HuR accumulation was not affected by the p38 MAPK inhibitor in one of the ovarian cancer cells (Fig 5), raising a possibility that additional stress signaling pathways could contribute to the HuR nucleo-cytoplasmic shuttling in OVCA433 cells.

The p38 MAPK pathway primarily governs the cellular response to stress and triggers apoptosis upon matrix detachment or oncogene-induced ROS accumulation (Dolado et al, 2007; Owens et al, 2009). During cancer progression, however, elevated levels of intracellular oxidants accelerate proliferation and transition to malignant phenotypes via upregulation of pro-survival pathways (Liou et al, 2016; Weinberg et al, 2010), and cancer cells must activate antioxidant defenses to maintain these elevated ROS at sub-lethal levels. Cancer cells can acquire resistance by uncoupling the p38 MAPK pro-apoptotic pathway from it’s ROS-sensing ability (Dolado et al., 2007). Therefore, cancer cells manipulate the cellular p38 MAPK surveillance program to favor their survival and adapt to a variety of stresses that they encounter during different stages of cancer. Indeed, downstream targets of p38 MAPK, including HIF-1α and uPA (urokinase-type plasminogen activator) (Emerling et al, 2005; Huang et al, 2000) are well-known drivers of angiogenesis and cancer invasion. Here we show that increased cytosolic SOD2 protein expression was also dependent on p38 MAPK activity, demonstrating that cancer cells overcome matrix detachment-induced stress via this pathway. Further investigation of the downstream mediators linking the p38 MAPK pathway to HuR under conditions of anchorage independence require further attention and should unveil novel stress response proteins important for anoikis resistance.

While the transcriptional regulation of antioxidant enzymes has been studied widely in the context of antioxidant response elements and stress response transcription factors such as Nrf2/NFE2L2, fewer studies have focused on translational regulation of these enzymes. In earlier work, the presence of an unknown stress-responsive SOD2 mRNA binding protein was reported across several cells and tissues of different species (Chung et al, 1998; Fazzone et al, 1993). Rat lung extracts contained a redox-sensitive SOD2 mRNA binding protein (Fazzone et al., 1993), and further analysis identified that this is associated with a cis-regulatory region located 111 bp downstream of the stop codon in rat SOD2 mRNA (Chung et al., 1998). The 3’ UTR of human SOD2 mRNA shares ∼75% homology with the rat 3’ UTR. Based on sequence comparison, we found that the sequence of the protein binding region partially overlaps with the first HuR binding sites from the PAR-CLIP analysis (Fig 3A) (Chung et al., 1998; Lebedeva et al., 2011). Among the different SOD2 mRNA splice variants, different 3’ UTRs have been reported (Supp Fig 3A). Variant 2 (NM_001024465) has a short 3’ UTR composed of a spliced region that excludes the majority of the HuR sites identified. Variant 1 (NM_000636) has been annotated to contain a 13.4 kb 3’ UTR. However, past studies have shown that the two most common SOD2 transcripts contain either a short 240 bp or a 3,439 bp segment of this 3’ UTR, which arise from use of a proximal and distal polyadenylation site, respectively (Supp Fig 3A) (Chaudhuri et al., 2012; Church, 1990). Interestingly, Chaudhuri et al. reported that the expression of these two SOD2 transcripts is altered between quiescent and proliferating cells, with the shorter transcript being associated with quiescence and increased protein expression (Chaudhuri et al., 2012). Moreover, radiation increased levels of the shorter SOD2 transcript levels of the 1.5 kb MnSOD transcript, with expression of the longer form remaining unaltered (Chaudhuri et al., 2012). The mechanisms for this radiation induced increase in the short 3’ UTR transcript remain unclear. However, we predict that is likely not HuR-dependent, as only the longer 3.4 kb 3’ UTR contains the majority of identified HuR binding sites, and we verified that ovarian cancer cells used in the present work express the transcript containing this longer 3’ UTR (Supp Fig 3B). It remains to be determined if and how these alternate 3’ UTR SOD2 transcripts are regulated during different sources of stress, and how their transcription co-operates with translational regulation through the activation of cell-specific RNA binding proteins, as well as the interplay with non-coding RNAs, such as miRNAs. A screen for miRNA binding reveals that the SOD2 mRNA contains potential binding sites for miRNAs throughout the length of the 3’ UTR. While most are located toward the far upstream region, several overlap with identified HuR binding sites. The role of miRNAs in regulating SOD2 expression have also been interrogated and several miRNAs identified that either positively or negatively regulate SOD2 levels in cancer (Kim et al., 2017). It remains to be determined if changes in miRNA binding further influence the regulation of SOD2 mRNA translation in anchorage-independence, and if this interplays with the regulation by HuR.

In conclusion, we show for the first time that SOD2 mRNA is an HuR target in anchorage-independent ovarian cancer cells. Although SOD2 regulatory mechanisms have been studied extensively at the transcriptional level, very few studies have investigated how translational regulation contributes to adaptable changes in SOD2 expression. The present findings uncover a novel post-transcriptional stress response mechanism by which tumor cells are able to rapidly increase their mitochondrial antioxidant capacity to adapt to stress associated with anchorage-independence, promoting survival during metastatic progression.

Materials and Methods

Cell Culture and Reagents

OVCA433 and OVCAR10 cells were provided by Dr. Susan K. Murphy (Duke University) and Dr. Katherine Aird (University of Pittsburgh), respectively. OVCA433 and OVCAR10 were grown in RPMI1640 supplemented with 10% FBS at 37 °C with 5% CO2. STR profiling is carried out routinely to validate cell identity, which revealed at the commencement of this work that OVCAR10 cells share the same STR profile as NIH-OVCAR3 cells. It is unclear if the OVCAR10 cell line was initially derived from the same patient as OVCAR3, or if OVCAR10 cells represent a sub-line derived from OVCAR3 cells. The protein synthesis inhibitor cycloheximide (Sigma) was added at a concentration of 20 µg/mL in fully supplemented growth media. For mRNA stability assays, actinomycin D (Sigma) was added at 10 µg/mL. The p38 MAPK inhibitor SB203580 was used at a final concentration of 20 µM.

Cell culture in adherent and ultra-low attachment (ULA) conditions

For attached conditions, cells were plated in 150-mm dishes and grown to ∼80% confluency. For anchorage independent cell culture, cells were trypsinized and seeded at a density (300,000 cells/2 mL media/well) in 6-well ULA (ultra-low attachment) plates (Corning: 3471) and collected at different time points for downstream analyses.

siRNA-mediated HuR/ELAVL1 knock-down

Cells were transfected with scramble non-targeting SMARTpool control (Dharmacon: D-001810-10-05) or HuR (ELAVL1)-specific SMARTpool siRNA oligonucleotides (Dharmacon: L-003773-00-0005) using Lipofectamine RNAiMAX (Invitrogen), and knock-down confirmed by western blotting.

Subcellular Fractionation

Cells in adherent and ULA plates were collected and the cell pellets were washed with ice-cold PBS. The cell pellets were processed as described in Sugiura et al. (Sugiura et al, 2017). Briefly, cells were centrifuged and resuspended in 200-500 µl of ice-cold homogenization buffer (10 mM HEPES pH 7.4, 220 mM mannitol, 70 mM sucrose, Roche protease and phosphatase inhibitor cocktails). The lysates were homogenized by several passages through 27-G needles. Lysates were centrifuged at 800 g for 10 min, followed by centrifugation of the supernatants at 2,500 g for 15 min at 4 °C. The mitochondrial pellets were resuspended in homogenization buffer and the supernatants were centrifuged at 100,000g for 1 h at 4 °C using a Beckman Coulter Optima MAX Ultracentrifuge. Post-centrifugation supernatants containing cytosolic fractions were transferred to new tubes and used for immunoblotting.

Immunoblotting

Protein concentrations were measured using the Pierce BCA protein assay kit. An equal amount of protein lysates was loaded onto 4-20% SDS-PAGE gels. Following electrophoresis, proteins were transferred to PVDF membranes. For detection of proteins, the membranes were incubated with the following antibodies overnight at 4 °C: SOD2 (A-2, Santa Cruz: sc-133134, 1:500 dilution); β-tubulin (9F3, Cell Signaling Technology: 2128, 1:1,000 dilution), ATP5A (Abcam: ab14748, 1:1000 dilution), β-actin (Thermo: AM4302, 1:10,000 dilution), HuR/ELAVL1 (3A2, Santa Cruz: sc-5261, 1:500 dilution), Phospho-p38 MAPK (Thr180/Tyr182, Cell Signaling Technology: 9211, 1:1000 dilution), p38 MAPK (A-12, Santa Cruz Biotechnology: sc-7972, 1:1000 dilution), MAPKAPK-2 (Cell signaling technology: 3042, 1:1000 dilution). The blots were developed using SuperSignal West Femto Maximum Sensitivity Substrate (Thermo: 34096) after incubation with horseradish peroxidase (HRP)-conjugated secondary antibodies (Amersham Biosciences) for 1 h at RT.

Immunoprecipitation (IP)

1-1.5 mg of cell lysates were pre-cleared by incubating with 2 µg normal rabbit IgG (Cell Signaling Technology: 2729S) or normal mouse IgG (Millipore: 12-371) on a rotator for 1 h at 4 °C followed by an additional 1 h incubation with protein A-(Thermo: 20333) or protein G-agarose beads (50 µL; Thermo: 20399) at 4 °C. Following centrifugation at 3000g for 10 min supernatants were transferred to clean tubes and incubated with either IgG or primary antibodies overnight at 4 °C. 50 µL of agarose beads were added to the lysates for 1-2 h at 4 °C and the antibody-bead complexes were washed three times in IP lysis buffer and further processed for downstream assays.

35S Protein Radiolabeling

Cells in adherent and ULA plates were treated with EasyTag Express35S Protein Labeling Mix (Perkin Elmer: NEG772), using 40 µl 35S (440 uCi) per 20 mL media in 150-mm dish, 4 µl 35S (44 uCi) /2 mL media/ well in ULA plates, according to a protocol adapted from Gallagher et al. (Gallagher et al, 2008). Following 2 h incubation in the presence of 35S-L-methionine and 35S-L-cysteine, cells were collected, washed with ice-cold PBS, and harvested using RIPA buffer supplemented with protease and phosphatase inhibitors. The cell lysates were rotated for 30 min at 4 °C, centrifuged at 12,000 rpm for 30 min at 4 °C and supernatants transferred to new tubes. After pre-clearing, the lysates were incubated overnight with 2 µg of normal rabbit IgG or SOD2 antibody (Abcam: Ab13533). Following SOD2 IP, the lysates were resolved in SDS-PAGE gels. The SOD2 band in each lane was cut with a razor blade and weighed. The bands were dissolved in 1 mL of 1X TGS running buffer overnight on a rocker at 4 °C. Next day, dissolved gel pieces were further heated for 20 min at 60 °C. The dissolved radioactive sample solutions were transferred to glass vials containing 10 mL of Opti-Fluor (Perkin Elmer) in duplicate (500 µl per vial). Liquid scintillation counting was performed using a Beckman Coulter Scintillation Counter. The readouts were normalized against the values from untreated samples.

Ribonucleoprotein Immunoprecipitation & RT-PCR

Cells were cultured in attached and anchorage independent conditions as described above. Before harvesting cells, 0.3% formaldehyde was added for 10 min at 37 °C for crosslinking followed by addition of glycine (final concentration 0.25 M) for 5 min for quenching. RNP-IP was performed as described in (Raspaglio et al., 2010; Tenenbaum et al, 2002) with modifications. Briefly, crosslinked cells were lysed in 500-1,000 µl NT1 buffer (100 mM KCl, 5 mM MgCl2, 10 mM HEPES, [pH 7.0], 0.5% Nonidet P40 [NP40], 1 mM dithiothreitol [DTT], 100 units/mL SUPERase·In RNase Inhibitor [Invitrogen: AM2694], protease inhibitors [Thermo: 78429], 0.2% vanadyl ribonucleoside complexes [New England Biolabs: S1402S]). After centrifugation of lysates at 16,000 g for 15 min, the supernatants were used for IP with normal mouse IgG or HuR antibody. The antibody-bead mixtures were washed several times with NT2 buffer (50 mM Tris-HCl [pH 7.4], 150 mM NaCl, 1 mM MgCl2, 0.05% NP40, RNAse inhibitor, protease inhibitor). IP samples for RNA elution were incubated with proteinase K (30 µg/100 µl NT2 buffer with 0.1% SDS) for 30 min at 60 °C. RNA was extracted using TRIzol, followed by cDNA synthesis (Quantabio: 95047) and SOD2 RT-PCR using the PrimeSTAR polymerase (Takara: R010A) with the following cycles: 98°C for 10 sec, 98°C for 10 sec + 60°C for 10 sec + 72°C for 20 sec X 35-38 cycles, followed by a final extension step at 72°C for 2 min. PCR products were analyzed by 2% agarose gel electrophoresis.

Polysome Profiling by Sucrose Density Gradient Centrifugation

Cells in adherent and ULA plates were incubated with cycloheximide (100 µg/mL) for 10 min at 37 °C before harvesting and were washed twice with ice cold 1X PBS containing cycloheximide. The cells were homogenized in 500 µl lysis buffer (50 mM HEPES, 75 mM KCl, 5 mM MgCl2, 250 mM sucrose, 100 ug/mL cycloheximide, 2mM DTT, 20 U/µl SUPERase·In RNase Inhibitor [Invitrogen: AM2694], 10% Triton X-100, 13% NaDOC) and polysome profiling carried out as previously described (Dang Do et al, 2009). Lysates were placed on ice for 10 min and centrifuged at 3000 g for 15 min at 4 °C. 500 µl supernatants were loaded on linear sucrose gradients ranging from 20% to 47% (10 mM HEPES, KCl 75 mM, 5 mM MgCl2, 0.5 mM EDTA) and were separated by ultracentrifugation in a SW41 rotor at 34,000 rpm for 4 h 15 min at 4 °C (Beckman Coulter). Subsequently, four sucrose fractions were collected using a UV/VIS absorbance detector. TRIzol reagent (Invitrogen) was added to each fraction for RNA isolation. Briefly, post-centrifugation at 3,200g for 20 min after addition of 1/5 volume of chloroform, the aqueous layer was transferred, and 1/2 volume of isopropanol was added for overnight precipitation at −20 °C. RNA was pelleted by centrifugation at 4,640 rpm for 55 min at 4 °C. RNA pellets were washed with 70% ethanol twice and dissolved in RNAse-free water. After cDNA synthesis and qPCR reactions, final PCR products were analyzed on 2% agarose gels.

Semi-quantitative real-time PCR

Total RNA was isolated by RNA isolation kit (Zymo Research: R2052) and used for cDNA synthesis (Quantabio: 95047) according to the manufacturer’s instruction. cDNA was mixed with iTaq™ Universal SYBR® Green Supermix (BioRad) and the primers listed in Table 1. Semi-quantitative real time RT-PCR was carried out using a BioRad qRT-PCR machine (BioRad), data normalized to the geometric mean of four housekeeping genes (Table 1), and expressed as fold-change in expression using the 2-ΔΔCT formula.

View this table:
Table 1:

Primers used for RT-PCR and semi-quantitative real time PCR.

Live/dead staining

Live and dead cell fractions of cells cultured for 2 h in anchorage independence was assessed by staining with 4 μM Calcein AM and 4 μM ethidium homodimer (in PBS; Sigma) to visualize live and dead cells, respectively. Cells were exposed to both dyes for 30 min at 37 °C, followed by imaging on a Keyence BZ-X700 fluorescence microscope. The percentage of live and dead cells were quantified using Image J.

Statistical Analysis

All data are representatives of at least three independent experiments. Data are presented as mean ± SEM with individual replicate values superimposed. Statistical analysis was performed using GraphPad Prism Software v9, with statistical tests chosen based on experimental design, as described in figure legends.

Author contributions

Y.S.K. designed the conceptual framework and experiments of the study, carried out the majority of the experiments and data analysis, prepared the figures and wrote the manuscript. J.E.W., P.T., Z.J. and A.E. assisted with experimental execution, and manuscript editing. K.M. and S.R.K. contributed to experimental design, data interpretation and manuscript editing. N.H. supervised and conceived the study, contributed to experimental design, assisted in data analysis, and assisted in writing and editing of the manuscript.

Conflict of interest

The authors have no conflicts of interest.

Acknowledgements

The authors would like to thank Ms. Sara Shimko and Lydia Kutzler for technical assistance. This work was supported by the U.S. National Institutes of Health grants R01CA242021 (N.H.) and R01CA230628 (N.H. & K.M.).

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Posted February 19, 2021.
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Supplementary Material

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HuR -dependent SOD2 protein synthesis is an early adaptation of ovarian cancer cells in response to anchorage independence
Yeon Soo Kim, Jaclyn E. Welles, Priscilla W. Tang, Zaineb Javed, Amal T. Elhaw, Karthikeyan Mythreye, Scot R. Kimball, Nadine Hempel
bioRxiv 2021.02.19.432010; doi: https://doi.org/10.1101/2021.02.19.432010
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