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Cisplatin and its analogues are the most commonly used agents in the treatment of head and neck squamous cell carcinoma (HNSCC). In this study, we investigated a possible role of epidermal growth factor receptor (EGFR), phosphorylation and degradation in cisplatin-induced cytotoxicity. Cisplatin treatment led to an increase in initial EGFR phosphorylation at the Y1045, the binding site of ubiquitin ligase, Casitas B-lineage lymphoma (c-Cbl), followed by ubiquitination in the relatively cisplatin-sensitive cell lines. However, cisplatin-resistant cell lines underwent minimal EGFR phosphorylation at the Y1045 site and minimal ubiquitination. We found that EGFR degradation in response to cisplatin was highly correlated with cytotoxicity in seven head and neck cancer cell lines. Pretreatment with epidermal growth factor (EGF), enhanced cisplatin-induced EGFR degradation and cytotoxicity, whereas erlotinib pretreatment blocked EGFR phosphorylation, degradation, and cisplatin-induced cytotoxicity. Expression of a mutant Y1045F EGFR, which is relatively resistant to c-Cbl mediated degradation, in Chinese hamster ovary cells and the UMSCC11B human head and neck cancer cell line protected EGFR from cisplatin-induced degradation and enhanced cell survival compared to WT-EGFR. Transfection of WT-c-Cbl enhanced EGFR degradation and cisplatin-induced cytotoxicity compared to control vector. These results demonstrate that cisplatin-induced EGFR phosphorylation and subsequent ubiquitination and degradation is an important determinant of cisplatin sensitivity. Our findings suggest that treatment with an EGFR inhibitor before cisplatin would be antagonistic, as EGFR inhibition would protect EGFR from cisplatin-mediated phosphorylation and subsequent ubiquitination and degradation, which may explain the negative results of several recent clinical trials. Furthermore, they suggest that EGFR degradation is worth exploring as an early biomarker of response and as a target to improve outcome.
Cisplatin and its analogues are among the most commonly used and effective agents in the treatment of head and neck cancers (1–4), as well as several other solid tumors, including those of the lung, testis, bladder and ovary (5–7). Cisplatin induces cytotoxicity via the production of DNA damage caused by formation of cisplatin-DNA adducts (8), which leads to irreparable DNA damage and, ultimately, cell death.
Cisplatin-DNA adduct formation might trigger downstream effects that could potentiate cisplatin toxicity. One such potential downstream effect of cisplatin-induced DNA damage that could affect cell survival occurs through the degradation of the EGF receptor. Cancer cell survival has been shown to be maintained by EGFR independent of its kinase activity. Weihua et al. found that knockdown of EGFR with siRNA led to cell death in an autophagic process (9). Nishikawa and Donato reported a central role for EGFR expression in cisplatin-mediated cytotoxicity (10–11). Furthermore, Niwa et al. used an antisense approach to target EGFR to inhibit growth of HNSCC, finding that treatment with a combination of docetaxel and EGFR antisense oligonucleotides resulted in increased cytotoxicity and reduced tumor volumes as compared with monotherapy (12). Using this approach Lai et al. showed that intratumoral EGFR oligonucleotide injection is safe and resulted in antitumor activity in patients with advanced HNSCC (13). Therefore, it is hypothesized that the agents that can induce or potentiate EGFR degradation would sensitize cells driven by EGFR.
We recently found that treatment of head and neck cancer cells with gemcitabine caused EGFR phosphorylation and subsequent degradation, and that inhibition of EGFR degradation using the proteasome inhibitor MG-132 increased cell survival (14). As cisplatin is the most commonly used chemotherapeutic agent in the treatment of HNSCC, we sought to understand the possible role of EGFR phosphorylation and degradation in cisplatin-induced cytotoxicity using specific genetic techniques rather than potentially non-specific proteasomal inhibitors. We found that treatment with cisplatin, like gemcitabine, led to an increase in EGFR phosphorylation followed by degradation in sensitive cell lines, and that this degradation strongly correlates with cytotoxicity. We then investigated the implications of cisplatin-induced EGFR degradation in the rational design of combination therapy involving EGFR inhibitors and chemotherapy.
Cisplatin was acquired from Bedford Laboratories (Bedford, OH, USA). Phospho-epidermal growth factor receptor (pY845EGFR), c-Cbl and GAPDH antibodies were purchased from Cell Signaling Technology (Beverly, MA). Phospho Y1045 (pY1045EGFR) antibody was obtained from Millipore (Billerica, MA). EGFR (Sc-03), and ubiquitin antibodies were acquired from Santa Cruz Biotechnology (Santa Cruz, CA). Erlotinib was kindly provided by Genentech (San Francisco, CA). EGF was obtained from Sigma (St. Louis, MO).
The Chinese hamster ovary (CHO) cell line was purchased from the American Type Culture Collection (Manassas, VA). The human head and neck squamous cell carcinoma cell lines UMSCC1, 10B, 11B, 12, 17B, 29 and 33 and cervix squamous cell carcinoma ME-180Pt cells were kindly provided by Dr. Thomas E. Carey (University of Michigan, Ann Arbor, MI). All cell lines were grown in RPMI 1640 supplemented with 10% cosmic calf serum (Hyclone, Logan, UT). For all in vitro experiments, cells were released from flasks using PBS containing 0.01% trypsin and 0.20 mmol/L EDTA, and 6×105 cells were plated onto 100-mm culture dishes two days before any treatment.
Cells were scraped into PBS containing a sodium orthovanadate and protease inhibitor mixture (Roche Diagnostic Co., Indianapolis, IN). Cells were incubated for 15 min on ice in Laemmli buffer (63 mM Tris-HCl, 2% (w/v) SDS, 10% (v/v) glycerol, and 0.005% (w/v) bromphenol blue) containing 100 mM NaF, 1 mM Na3Vo4, 1 mM phenylmethylsulfonyl fluoride, and 1 μg/ml aprotinin. After sonication, particulate material was removed by centrifugation at 13,000 rpm for 15 min at 4°C. The soluble protein fraction was heated to 95°C for 5 min, then applied to a 4–12% bis-tris precast gel (Invitrogen, Carlsbad, CA) and transferred onto a PVDF membrane. Membranes were incubated for 1h at room temperature in blocking buffer consisting of 3% BSA and 1% normal goat serum in Tris-buffered saline [137 mM NaCl, 20 mM Tris-HCl (pH 7.6), 0.1% (v/v) Tween 20]. Membranes were subsequently incubated overnight at 4°C with 1 μg/ml primary antibody in blocking buffer, washed, and incubated for 1h with horseradish peroxidase-conjugated secondary antibody (Cell Signaling, Danvers, MA). After three additional washes in Tris-buffered saline, bound antibody was detected by enhanced chemiluminescence plus reagent (Amersham Biosciences, Piscataway, NJ). For quantification of relative protein levels, immunoblot films were scanned and analyzed using ImageJ1.32j software (NIH, Bethesda, MD). Unless otherwise indicated, the relative protein levels shown represent a comparison to untreated controls.
Following treatments, cells were trypsinized, washed twice with 1x PBS, and cell lysates were prepared by incubation for 30 min on ice in fresh lysis buffer [1% Triton X-100, 0.1% sodium dodacyl sulfate, 0.15M sodium chloride, 0.01M sodium phosphate, pH 7.2 1 mmol/L phenylmethylsulfonyl fluoride, 2 μg/mL aprotinin, 0.2 mmol/L sodium orthovanadate, 50 mM sodium fluoride, 2 mM EDTA and 20 mM ammonium molybdate]. Immunoprecipitation of EGFR was performed as described previously (14).
Total RNA was isolated from UMSCC11B cells using the RNA easy mini kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. RNA (1μg) was reverse transcribed to cDNA using the High Capacity cDNA Archive kit (Applied Biosystems, Foster City, CA) and purified (Millipore Centrifugal Filter Units, Billerica, MA). Diluted cDNA was used to amplify GAPDH (GAPDH F: 5′ GAG TCA ACG GAT TTG GTC GT 3′ and GAPDH R: 5′ TTG ATT TTG GAG GGA TCT CG 3′) and EGFR (EGFR F: 5′ CTC AGC CAC CCA TAT GTA CC 3′ and EGFR R: 5′ CGT CCA TGT CTT CTT CAT CC 3′) by quantitative reverse-transcription PCR (qRT-PCR) using SYBR green chemistry (Applied Biosystems, Warrington, UK). The PCR products were resolved by electrophoresis on 1.5% agarose gels and melting curve analysis was carried out to confirm the specificity of the product. The δδCt method was used to analyze the data as described (15–16), and GAPDH was used as control.
Clonogenic assays were performed using standard techniques (17). The fraction surviving each treatment was normalized to the survival of the control cells. Cisplatin cell survival curves were fitted using the equation SF=(C50)m/[(C50)m+Cm], where SF is the surviving fraction, C is the cisplatin concentration, C50 is the concentration of cisplatin that produces a 50% cell survival and m is the slope of the sigmoid curve. The effects of EGF, or erlotinib, on cisplatin-induced clonogenic death were calculated by comparing the ratio of the areas under the respective cell survival curves (AUCs).
A site directed mutagenesis approach was used to create the Y1045F substitution in the pEYFP-N1 vector. The mutation was confirmed by sequencing of the plasmid in the DNA sequencing core at University of Michigan. Wild-type EGFR in the pEYFP-N1 vector was provided by Dr. Zhixiang Wang (University of Alberta). The wild-type c-Cbl construct was provided by Dr. Nancy Lill, University of Iowa. UMSCC11B and CHO cells were transiently transfected with the constructs using Lipofectamine (Invitrogen, Carlsbad, CA) according the manufacturer’s instructions.
Live cell imaging of CHO and UMSCC11B cells transfected with wild-type EGFR and Y1045F EGFR constructs was performed using an Olympus DP70 camera fitted in an Olympus 1X-71 microscope. All the images were captured at 60X magnification and processed similarly in Adobe Photoshop by removing the unsharp mask.
Results are presented as mean ± SE of at least three experiments. Student’s t test was used to assess the statistical significance of differences. A significance level threshold of p < 0.05 was used.
Our initial goal was to determine if there were a correlation between cisplatin-induced cytotoxicity and EGFR degradation. Cisplatin cytotoxicity was assessed by clonogenic survival analysis in seven HNSCC cell lines, UMSCC1, 10B, 11B, 12, 17B, 29, 33, and in the cisplatin-resistant cervical carcinoma ME-180Pt cell line. We selected ME-180Pt cells for use along with the head and neck cancer cell lines because platinum resistance in this cervical cancer line has previously been reported to be associated with higher expression of EGFR (10–11). Our results revealed that the UMSCC17B and 11B lines were the most sensitive, and UMSCC1 & ME-180Pt were the most resistant, with the other cell lines displaying intermediate sensitivity (Figure 1A). Based on these results and clinically achievable levels of cisplatin (18), we selected a 2-hr exposure to 10 μM cisplatin to assess the effects of cisplatin treatment on EGFR protein level by immunoblotting relative to untreated controls in each of the eight cell lines. We found a direct relationship between relative EGFR levels at 72 hrs and surviving fraction after cisplatin treatment (R2=0.97) (Figure 1A). For example, in response to 10 μM cisplatin, the relatively cisplatin-sensitive UMSCC11B cell line showed a 95% decrease in EGFR levels at 72 hrs, whereas the relatively cisplatin-resistant ME-180Pt line showed almost no change in EGFR levels at 72 hrs (Figure 1B).
We then sought to determine whether cisplatin, like gemcitabine, caused initial EGFR phosphorylation followed by subsequent degradation in head and neck cancer cell lines. We chose UMSCC11B as a representative cisplatin-sensitive and UMSCC1 and ME-180Pt as cisplatin-resistant cell lines for further studies. We observed increased phosphorylation of EGFR at both the Y845 and Y1045 positions in (sensitive) UMSCC11B cells. We also noted a moderate increase in phosphorylation of EGFR at the Y845 site, but little to no increase in phosphorylation of the Y1045 site in (resistant) UMSCC1 and ME-180Pt cells. The pY845 and pY1045 EGFR levels decreased by 72 hrs in the cisplatin-sensitive UMSCC11B cell line. However, in cisplatin-resistant UMSCC1 and ME-180Pt, pY845 EGFR levels remained high at 72 hrs. These results showed that phosphorylation of EGFR at Y1045 was more strongly correlated with cisplatin cytotoxicity than the Y845 site and suggested to us that the Y1045 site might be critical for the eventual degradation of EGFR in response to cisplatin. To determine if this observed EGFR phosphorylation affected signaling, we analyzed the downstream mediators of EGFR signaling in these cell lines. As anticipated, pERK levels were significantly increased in UMSCC11B by 2 hrs and then decreased at 72 hrs in a similar manner to pEGFR levels. In cisplatin-resistant UMSCC1 and ME-180Pt, we noticed an initial moderate increase in pERK levels, which did not decrease appreciably by 72 hrs, also consistent with pEGFR levels.
In order to determine whether the decrease in EGFR levels was due to decreased production or increased degradation, we assessed ubiquitination of EGFR at 2 and 6 hrs after treatment with cisplatin in cisplatin-sensitive and resistant cell lines. In the sensitive cell line UMSCC11B, we observed EGFR binding to ubiquitin in response to cisplatin (Figure 1C), whereas in the relatively resistant cell line ME-180Pt, there was no increase in ubiquitination of EGFR at 2 and 6 hrs after cisplatin treatment. This lack of ubiquitination was congruent with the lack of EGFR degradation in these cells. UMSCC1 cells showed a slight increase in ubiquitinated EGFR at 6 hrs, but not as substantial as was observed in UMSCC11B cells. Therefore, ubiquitination of EGFR in the three cell lines was directly correlated with levels of Y1045 phosphorylation of EGFR in response to cisplatin. We further carried out EGFR transcript analysis in UMSCC1, ME-180Pt and UMSCC11B cells. Real time PCR results showed an increase in EGFR transcript levels in response to cisplatin at both 24 and 72 hrs as compared to controls in UMSCC11B (Figure 1D). These results suggested that the decrease in EGFR levels was occurring at the protein level, and its transcript level might be undergoing a partially compensatory upregulation. In the case of UMSCC1 and ME-180Pt, EGFR transcript levels did not increase at 24 and 72 hrs but in fact slightly decreased for ME-180Pt (Figure 1D). We also analyzed the effects of cisplatin on EGFR degradation after blocking protein synthesis using cyclohexamide (50 μg/ml) in UMSCC11B cells. EGFR degradation was more rapid in the case of cells treated with combination of cisplatin and cyclohexamide compared to cyclohexamide alone (Supplementary figure 1), confirming that EGFR degradation was occurring earlier than it was detected through immunoblotting, and suggesting that the increase in transcription of EGFR is partially compensating for the loss of protein.
We next hypothesized that if EGFR phosphorylation were critical for its eventual degradation, then pretreatment with EGF would enhance cisplatin-induced EGFR degradation while erlotinib would protect EGFR from degradation. For these experiments, ME-180Pt (resistant) cells were treated with EGF (10 ng/ml) for 1 hr before cisplatin was added to the cells. As we anticipated, EGF induced rapid EGFR degradation by 24 hrs after treatment with cisplatin (Figure 2A). Cell survival data for EGF and cisplatin combination showed a significant decrease in clonogenic survival with combined cisplatin and EGF as compared to cisplatin alone in ME-180Pt cells (Figure 2B). Conversely, pretreatment of UMSCC11B (sensitive) cells with erlotinib (3 μM) for 2 hrs before the addition of cisplatin protected EGFR from cisplatin-induced degradation at 72 hrs, supporting a critical role for EGFR phosphorylation in cisplatin-induced EGFR degradation (Figure 2C). Likewise, pretreatment with erlotinib significantly protected the UMSCC11B cells from cisplatin-induced cytotoxicity (Figure 2D). Erlotinib pretreatment also protected UMSCC1 cells from cisplatin-induced cytotoxicity (Supplementary figure 2). These findings emphasize the important role of initial EGFR phosphorylation followed by degradation in cisplatin cytotoxicity.
We next hypothesized that blocking EGFR phosphorylation at the Y1045 residue, which is necessary for binding with the ubiquitin ligase c-Cbl, and thus required for EGFR degradation, would protect cells from cisplatin-induced toxicity. We initially tested our hypothesis in a system with no endogenous expression of EGFR so as to avoid confounding effects of native EGFR. We selected CHO cells, which have been shown to be EGFR-negative (19), but upon transfection with EGFR constructs are able to not only express EGFR but also to activate normal downstream signaling events (20–21). We utilized wild-type EGFR and Y1045F EGFR constructs in the N1-EYFP vector backbone to monitor changes in EGFR levels and localization in response to cisplatin in live cells. CHO cells were transfected with either wild-type EGFR or Y1045F EGFR constructs, treated with cisplatin, and assessed for EGFR levels at 24 and 72 hrs after treatment (Figure 3A). We detected a more rapid disappearance of wild-type EGFR when compared to Y1045F EGFR, verifying that Y1045 site is important for EGFR degradation. Immunoblotting data also confirmed these results (Figure 3A, middle panel). We then assessed the role of EGFR degradation in cisplatin toxicity. As expected, expression of the Y1045F-EGFR construct in CHO cells induced significant protection from cisplatin cytotoxicity as compared to wild-type EGFR expressing cells (DMF 1.5) (Figure 3B).
Next, we sought to extend our findings on the critical role of the 1045 phosphorylation of EGFR to a head and neck cancer cell line. We transfected UMSCC11B cells with wild-type or Y1045F EGFR constructs and treated cells with cisplatin to monitor the effects on transfected EGFR in these cells. Similar to our results in CHO cells, we found a more rapid decrease in wild-type EGFR as compared to Y1045F EGFR in response to cisplatin (Figure 3C). In addition, although the high endogenous levels of EGFR make it more difficult to detect a difference in EGFR levels after transfection of the mutant, we found that Y1045F-transfected cells were protected from cisplatin-induced EGFR degradation (Figure 3C). Importantly, cells expressing the Y1045F EGFR construct were significantly protected from cisplatin-induced cytotoxicity (Figure 3D). These results, especially when combined with those obtained in the CHO system described above, strengthen the importance of Y1045 phosphorylation of EGFR in cisplatin-induced EGFR degradation and resulting cytotoxicity.
To further assess the role of EGFR degradation in cisplatin-induced cytotoxicity in head and neck cancer, we decided to accelerate ubiquitination by enhancing the ability of c-Cbl to bind EGFR. Therefore, we transfected UMSCC11B cells with a control vector and wild-type c-Cbl constructs and assessed the effects on EGFR degradation and cytotoxicity. We found that cells transfected with wild-type c-Cbl showed enhanced EGFR degradation as compared with control vector transfected cells (Figure 4A). Wild-type c-Cbl transfected cells had a greater sensitivity to cisplatin than cells transfected with control vector construct (Figure 4B). We further found that after cisplatin treatment, wild-type c-Cbl expressing cells underwent increased ubiquitination of EGFR compared to cells expressing the control vector (Figure 4C). These results confirm that a sufficient degree of EGFR degradation (in this case, achieved by enhancing c-Cbl activity to degrade approximately 90% of EGFR) increases cisplatin-mediated cytotoxicity.
In this study, we demonstrate that cisplatin-induced cytotoxicity in head and neck cancer is critically dependent upon phosphorylation and subsequent degradation of EGFR. This conclusion is supported by our findings that: 1) decreased EGFR levels after cisplatin treatment correlate with decreased clonogenic survival; 2) stimulation of EGFR phosphorylation and degradation (by treatment with EGF) prior to cisplatin treatment increases cytotoxicity; and 3) inhibition of EGFR phosphorylation and degradation (by pretreatment with erlotinib) before cisplatin diminishes cytotoxicity. Using distinct genetic methods of preventing and accelerating EGFR degradation, we then confirmed that preventing degradation decreases cisplatin cytotoxicity, whereas promoting EGFR degradation increases cisplatin cytotoxicity. These findings elucidate a new step in the process by which cisplatin produces cell death and suggest that enhancing EGFR degradation may be a novel way of overcoming cisplatin resistance in head and neck cancer cells. Furthermore, our findings provide insight into maximizing the therapeutic benefit of EGFR inhibitors when used in combination with chemotherapy.
Substantial evidence now shows that radiation and chemotherapy induce EGFR phosphorylation and activation of downstream survival pathway. For example, Benhar et al. reported that cisplatin-induced EGFR phosphorylation (22) coincides with DNA adduct formation, suggesting that a DNA damage sensor activates (directly or indirectly) EGFR and downstream cell survival pathways. Others have suggested that cisplatin induces EGFR activation via a p38 mitogen activated kinase, and that this activation leads to downstream effectors such as protein kinase B/AKT (23). In addition, phosphorylated EGFR can undergo nuclear translocation and interact with DNA protein kinase (24) and can also mediate DNA repair (25). Likewise, ionizing radiation produces EGFR activation and nuclear translocation, which stimulates repair mechanisms (24). Nuclear localization of EGFR has also been found to be important for its role as a transcription factor that activates genes required for cell growth and proliferation (26). Inhibition of this activation of downstream survival and repair signals by treatment with an EGFR inhibitor after chemotherapy causes dramatic potentiation of cell death (14, 27), which has clinical implications as described below.
In contrast to EGFR activation after chemotherapy and radiation, the role of treatment-induced EGFR degradation has been much less appreciated. Our prior finding that gemcitabine causes EGFR degradation and cell death (14) and the results of our current study show that radiation (unpublished data) and chemotherapy-induced degradation causes cell death downstream of the initial DNA damage. Our findings are in agreement with a recent study demonstrating that loss of EGFR can induce autophagic cell death (9), and that antisense oligonucleotides against EGFR can produce clinical responses (13). Thus, the degradation of EGFR might decrease the ability of cells to repair damaged DNA, leading to increased cytotoxicity.
Our studies are consistent with several preclinical studies that have demonstrated a direct correlation between EGFR activation and sensitivity to cisplatin (28–31). In particular, Christen et al. reported that stimulation of ovarian carcinoma cells with EGF sensitized cells to cisplatin, independent of the mitogenic effects of EGF (32). Similarly, Nishikawa et al. found that the cisplatin-resistant cervical carcinoma cell line ME-180Pt (used in our study) was sensitized to cisplatin by EGF-pre treatment (10). Although the mechanism by which EGF sensitized cells to cisplatin was not elucidated, we would hypothesize that EGF increased EGFR activation and subsequent degradation.
As we now know that both cisplatin treatment and EGF stimulation can cause EGFR phosphorylation leading to receptor degradation, it is of interest to compare the mechanisms of these two processes. EGFR is internalized and degraded following stimulation with EGF via binding with c-Cbl at the phosphorylated Y1045-EGFR residue (33), which is a docking site for the ubiquitin ligase c-Cbl (33–34). Similarly, in response to treatment with cisplatin, we found that EGFR is phosphorylated at the Y1045 site, ubiquitinated and degraded via c-Cbl, as occurs in degradation after ligand binding. Thus it appears that EGFR phosphorylation, ubiquitination and degradation after EGF stimulation resemble that resulting from cisplatin treatment, although differences might still emerge with further study.
Irrespective of the exact mechanism of cisplatin-induced EGFR degradation, our results suggest that that the schedule of EGFR inhibition and chemotherapy can have a crucial impact on the effectiveness of therapy. We have found that the activation of EGFR is necessary for its subsequent degradation and resultant chemosensitization. Under these conditions, the addition of an EGFR inhibitor prior to cisplatin treatment prevents EGFR activation, thereby resulting in a cytoprotective effect. In contrast, treatment with chemotherapy or radiation first, leading to EGFR activation, followed by an EGFR inhibitor (depriving the cell of downstream “life” and DNA repair signaling) stimulates degradation and has a synergistic cytotoxic effect (27, 35). This is consistent with previous studies examining combination treatment with EGFR inhibitors and chemotherapy, where it has been suggested that EGFR phosphorylation in response to chemotherapeutic agents (29, 36) or anti-metabolites (37–38) is necessary for the chemosensitization benefit of added EGFR inhibitors. Thus, our findings may provide the underlying biological rationale to the paradoxical clinical observations that while EGFR inhibitors after chemotherapy can increase treatment efficacy, the converse schedule could actually be antagonistic (39).
It may be possible to develop new methods of potentiating cisplatin-induced cytotoxicity by discovering new ways of increasing cisplatin-induced EGFR degradation. Under conditions of cellular stress, the molecular chaperone heat shock protein 90 (Hsp90) regulates the stability of several oncoproteins (40). Yang et al (41) have reported that association of mutant EGFR with Hsp90 decreases its downregulation mediated by c-Cbl binding. It is also known that ErbB-2 can bind to both EGFR and Hsp90 directly. Whether an EGFR-Hsp90 interaction (directly or via ErbB-2) is involved in cisplatin-induced EGFR activation and degradation is not known at this time. In addition to EGFR and stress-related molecular interactions, it would also be interesting to explore if the interaction between EGFR and DNA-PK plays a role in cisplatin-induced cell death.
In summary, we show here that EGFR phosphorylation and degradation is a determinant of cisplatin cytotoxicity in head and neck cancer cells, and that this degradation is mediated by c-Cbl. At present, the mechanism of cisplatin-induced signaling in EGFR degradation is not known. Regardless of the mechanism of degradation, our results suggest that the incorporation of EGFR inhibitors into therapy may be most effective following cisplatin administration, and may even be antagonistic if given prior to cisplatin. Thus, our results indicate that EGFR inhibitors should be used after chemotherapy to facilitate EGFR degradation and chemosensitization. These findings also indicate that agents that facilitate EGFR degradation should be explored in overcoming cisplatin resistance.
We thank Dipankar Ray for helpful discussion, Mary Davis for her careful review of the manuscript, and Steven Kronenberg for assistance in making figures.
Grant support: This research is supported by NIH R01CA131290, NIH through the University of Michigan Head and Neck Specialized Program of Research Excellence grant P50CA097248, Michigan Institute for Clinical and Health Research, University of Michigan Cancer Center support grant 5 P30 CA46592 (M.K. Nyati), and Howard Hughes Medical Institute training fellowship (S.M.Hiniker).