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1 Departments of Biochemistry,
2 Medicine,
3 Radiation Oncology, and
4 Microbiology and Immunology, Virginia Commonwealth University, 401 College Street, Richmond, Virginia 23298, USA
5 IMBECU-CONICET, Mendoza, Argentina
6 Division of Gastroenterology/Hepatology, Faculty of Medicine, Queen Mary Hospital, University of Hong Kong, Pok Fu Lam Road, Hong Kong, China
7 Departments of Pathology, Neurosurgery and Urology, Herbert Irving Comprehensive Cancer Center, Columbia University Medical Center, College of Physicians and Surgeons, New York, New York 10032, USA
8 Division of Human Gene Therapy, Departments of Medicine, Pathology and Surgery, Gene Therapy Center, University of Alabama at Birmingham, Birmingham, 901 19th Street South, BMR2-502, Birmingham, Alabama 35294, USA
(Requests for offprints should be addressed to P Dent who is now at Department of Biochemistry, Massey Cancer Center, 401 College Street, Box 980058 Virginia Commonwealth University, Richmond, Virginia 23298-0058, USA; Email: pdent{at}hsc.vcu.edu)
This paper was presented at the 2nd Tenovus/AstraZeneca Workshop, Cardiff (2006). AstraZeneca supported the meeting and the Welsh School of Pharmacy, Cardiff University has supported the publication of these proceedings.
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Introduction |
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TopAbstractIntroductionMaterials and methodsResults and discussionReferences |
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Growth factors interact with plasma membrane receptors, which transduce signals through the membrane to its inner leaflet (Sizemore et al. 1999, Fanton et al. 2001, Ludde et al. 2001, Moriuchi et al. 2001, Pruitt & Der 2001, Balmanno et al. 2003, Blalock et al. 2003, Cox & Der 2003). Growth factor signals, via guanine nucleotide exchange factors, can increase the amount of GTP bound to membrane-associated GTP-binding proteins, including RAS (Sklar 1988, Pruitt & Der 2001). There are three widely recognized isoforms of RAS: Harvey (H), Kirsten (K), and Neuroblastoma (N) (Reuther & Der 2000). GTP–RAS can interact with multiple downstream effector molecules including the Raf-1 protein kinase and the phosphatidylinositol 3-kinase (PI3K) lipid kinase. Receptor-stimulated guanine nucleotide exchange of ‘RAS’ to the GTP-bound form permits Raf-1 and P110 PI3K to associate with ‘RAS’, resulting in kinase translocation to the plasma membrane environment where activation of these kinases, via complex mechanisms, takes place. RAS contains a GTPase activity that converts bound GTP to GDP, resulting in inactivation of the RAS molecule. PI3K enzymes are also translocated to the plasma membrane environment via the P85 SH2-domain interaction with phosphorylated tyrosine residues on adaptor proteins and growth factor receptors, e.g. GAB2, IRS-1, and ERBB3 (Hellyer et al. 2001).
Mutation of RAS results in a loss of GTPase activity, generating a constitutively active RAS molecule that can lead to elevated activity within downstream signaling pathways. Approximately, one third of human cancers have RAS mutations, primarily the K-RAS isoform that also leads to a radioprotected phenotype (Sklar 1988, Ellis & Clark 2000). Of note is that some studies suggest that K-RAS and H-RAS have different but over-lapping signaling specificities to downstream pathways as judged by in vitro cell-based studies and in animal knock-out models; thus mutant K-RAS is thought to preferentially activate the Raf-1/extracellular regulated kinase (ERK1/2) pathway, whereas mutant H-RAS is believed to preferentially activate the PI3K/AKT pathway (Yan et al. 1998, Liebmann 2001, Ross et al. 2001). It has been argued that ERK1/2 and PI3K signaling downstream of K-RAS and H-RAS respectively, can in turn control cell growth and cell survival following exposure to multiple growth factors, e.g. Epidermal growth factor (EGF) (Dent et al. 1999, Ludde et al. 2001, Moriuchi et al. 2001).
Loss of the single allele of mutated active K-RAS expression has been shown in HCT116 cells to abolish tumor formation in athymic mice and enhance radio-sensitivity (Baba et al. 2000, Ries et al. 2000). The findings presented in these studies were linked to reduced expression of the paracrine growth factor epiregulin. Repeated irradiation of tumor cells can also increase expression of transforming growth factor 
(TGF) and ERBB1 (Schmidt-Ullrich et al. 1994). Increased proliferative rates and poor prognosis of carcinomas in vivo have also been correlated with increased expression of ERBB1 (Putz et al. 1999).
The studies described herein examine the role of p53, ERBB receptors, ERBB receptor paracrine ligands, and mutated active RAS proteins in the signaling and survival responses of multiple human carcinoma cell lines exposed to ionizing radiation. Radiation exposure causes inhibition of protein tyrosine phosphatase function and the activation of ERBB receptors and downstream signaling pathways, for example, ERK1/2 and AKT. Activation of ERK1/2 promoted the cleavage and release of paracrine ligands, which caused re-activation of ERBB family receptors and intracellular signaling pathways. Expression of activated forms of RAS family members and alterations in p53 mutational status were noted to regulate radiosensitivity in isogenic cells, in part, by modulating the expression of ERBB receptors and ERBB receptor paracrine ligands. These alterations in receptor and ligand expression correlated with changes in the ability of HCT116 cells to activate ERK1/2 and AKT after irradiation, and survive radiation exposure. Thus, tumor cells are dynamic and rapidly adapt to any single therapeutic challenge to maintain tumor cell growth and viability.
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Materials and methods |
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TopAbstractIntroductionMaterials and methodsResults and discussionReferences |
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Anti-phospho-ERK1/2, anti-Raf-1, anti-total phosphortyrosine, anti-total-ERK2, anti-ß-actin antibodies, and protein A/G-conjugated agarose were from Santa Cruz Biotechnology (Santa Cruz). Anti-phospho-Ser473-AKT, anti-phospho-Thr308-AKT, anti-total-AKT, anti-phospho-Ser259-RAF, anti-phospho-Tyr1289 ERBB3, anti-phospho-Tyr1173 ERBB1, and anti-phospho-Tyr1187 ERBB2 antibodies were from Cell Signaling Technology (Beverly, MA, USA). Antibodies to detect total RAS, H-RAS and mutant K-RAS, and the neutralizing anti-heregulin antibody and the neutralizing anti-TGF antibody were from Oncogene Research Products (Cambridge, MA, USA). Antibodies to detect ERBB1, ERBB2, ERBB3, and ERBB4 and for immunoprecipitation of ERBB3 were from Neomarkers Lab Vision Corporation (Fremont, CA, USA). Antibodies for anti-PI3K p85 and anti-PI3K p110 were from Upstate Biotechnology (Lake Placid, NY, USA). Oligonucleotides for semi-quantitative PCR were synthesized by the VCU oligonucleotide core facility and were based on published sequences. Reverse transcriptase-PCR (RT-PCR) was performed using a commercial kit (Quiagen OneStep RT-PCR) and 1 µg RNA in a final volume of 25 µl and following the instructions of the manufacturer (Qiagen). The MEK1/2 inhibitors PD98059, U0126, and PD184352, the PI3K inhibitor LY294002, the ERBB1 inhibitor AG1478, the ERBB2 inhibitor AG825 were from Calbiochem (San Diego, CA, USA; Caron et al. 2005a,b).
Methods
Generation of HCT116 cell lines without mutant K-RAS and expressing mutant H-RAS
HCT116 mutant K-RAS-deleted cells were generated by homologous deletion of the mutant K-RAS allele as described (Shirasawa et al. 1993, Baba et al. 2000, Caron et al. 2005a,b). A plasmid to express mutant active H-RAS (H-RAS V12) was kindly provided by the laboratory of Dr M Wigler (Cold Spring Harbor, NY, USA). HCT116 cell lines were transfected by electroporation at 600 V for 60 ms using a Multi-porator Eppendorff (Hamburg, Germany) with control plasmids (‘C2’ cells) or plasmids to express H-RAS V12 (‘C10’ and ‘C3’ cells). Pools of transfected cells were obtained by puromycin (RAS) selection and individual colonies isolated and then characterized.
Culture of HCT116 cell lines
Asynchronous HCT116 carcinoma cells were cultured in DMEM media supplemented with 10% (v/v) fetal calf serum at 37 °C in 95% (v/v) air/5% (v/v) CO2. Cells were plated at a density 3x103 cells/cm2 plate area and all cells were plated from log-phase cultures. For radiation-induced activations of protein kinases, cells were cultured for 4 days in this media, and 24 h prior to irradiation, were cultured in serum-free DMEM medium. For colony formation assays, cells were plated at low density (250–2000 cells per dish), and 24 h after plating, for the 24 h prior to irradiation, they were cultured in serum-free DMEM medium. Cells were irradiated (1–4 Gy) or as indicated in the text; media was replaced with serum-containing media 24 h after radiation exposure. Plates were washed in PBS 10–14 days after exposure, fixed with methanol, and stained with a filtered solution of crystal violet (5% w/v). After washing with tap water, the colonies were counted both manually (by eye) and digitally using a ColCount TM plate reader. Data presented are the arithmetic mean (± S.E.M.) from both counting methods from multiple studies.
Culture of A431, DU145, and MDA-MB-231 cells
Cells were cultured in RPMI-1640 medium supplemented with 5% (v/v) fetal calf serum at 37 °C in 95% (v/v) air/5% (v/v) CO2. For radiation-induced activation of kinases, cells were cultured for 4 days and for 24 h prior to irradiation were cultured in serum-free medium.
Exposure of cells to ionizing radiation and cell homogenization
Cells were cultured as described above. As indicated, 1 h prior to irradiation, cells were treated with either vehicle (DMSO), U1026 (1 µM), PD184352 (2 µM), or LY294002 (1 µM). Treatment was from a 100 mM stock solution and the maximal concentration of vehicle (DMSO) in media was 0.01% (v/v). Cells were irradiated using a 60Co source at dose rate of 1.8 Gy/min. Cells were maintained at 37 °C throughout the experiment, except during the irradiation itself. Zero time is designated as the time point at which exposure to radiation ceased. After radiation-treatment, cells were incubated for specified times followed by aspiration of media and immediately homogenized in either lysis buffer for immune complex kinase or phosphatase assays (see below), or 1 ml SDS-PAGE lysis buffer (5% (w/v) SDS, 40% (v/v) glycerol, 250 mM Tris–HCl, and 10% (v/v) 2-mercaptoethanol). Homogenates were sonicated, boiled for 10 min, the protein concentration determined by Bradford assay (Coomassie Protein Assay Kit, Pierce Biotechnology, Rockford, IL, USA), and stored frozen (–20 °C) prior to use.
Immunoprecipitation from cell lysates
After irradiation, cells were incubated for specified times, followed by aspiration of media and snap-freezing at 70 °C on dry ice. Cells were lysed and Raf-1 and ERK1/2 activity determined as described by Dent et al. (1995a,b).
Protein tyrosine phosphatase assay
Cellular PTP activity was assessed by an in vitro assay with autophosphorylated EGFR as substrate (Dent et al. 2003a,b). EGFR was purified from A431 cells by affinity chromatography on lentil lectin Sepharose as previously described (Tomic et al. 1995).
TGF ELISA/concentration of release assay
ELISA kits for mammalian TGF were obtained from Oncogene. Assays were performed according to the manufacturer’s protocol with slight modifications. Briefly, cells were grown to 60% confluence in RPMI medium alone or supplemented with AG1478, PD98059, 60 min before exposure to 60Co-radiation, and the media sampled after various intervals of incubation at 37 °C. Medium sample and TGF standard in triplicate were added to the TGF-pre-coated wells and incubated for 4 h at room temperature. Samples were then processed according to the manufacturer’s protocols. Media without cells, and media incubated with EGF were used as a negative control in each assay. All reactions were measured using a spectrophotometric plate reader at a wavelength of 490 nm (BioTek Instruments, Inc., Highland Park, VT, USA).
SDS-PAGE and western blotting
Depending on the protein to be studied, a volume of homogenate containing 10, 20, or 40 µg total protein was loaded in 12% (w/v) acrylamide gels and subjected to SDS-PAGE. Gels were transferred to nitrocellulose and western blotted using specific antibodies. Blots were developed using Western Lightning Chemiluminescence Reagent Plus (Perkin–Elmer Life Sciences, Boston, MA, USA) and Kodak X-ray film, Densitometric analysis for ECL immunoblots, and RT-PCR analyses was performed using a Fluorochem 8800 Image System and software (Alpha Innotech Corporation, San Leandro, CA, USA) and band densities were normalized to that of ß-actin in the same sample and expressed as a percentage of the respective control in each experiment as indicated in the figure legend.
Cell death assays: Wright Giemsa for apoptosis
Cells were plated at 5x104 cells per well in 12-well plates and 24 h later, serum-starved. The plates were mock-exposed or irradiated 24 h later at 1 or 4 Gy and harvested 96 h after irradiation by trypsinization followed by centrifugation onto glass slides (cytospin) at 800 r.p.m. for 10 min. The cells were fixed and stained with a commercial kit (Diff-Quik) following the instruction of the manufacturer (Dade Behring AG, Düdingen, Switzerland). Randomly selected fields of stained cells (~200 cells per field, n=5 per slide) were counted for apoptotic nuclear morphology.
Membrane preparation from HCT116 cells.
Cells were cultured as described above. Cells were scraped into 10 ml 1 mM NaHCO3 pH 7.4, 1 mM Na pyrophosphate, 1 mM Na orthovanadate, 1 mM EDTA, 1 mM EGTA, 0.1 mM PMSF, and incubated for 1 h at 4 °C, 24 h after serum withdrawal/starvation. Crude membrane preparations were prepared as described by Dent et al. (1995a,b) by sucrose density over-layer centrifugation.
Statistical analysis
Comparison of the effects of treatments was performed using one-way ANOVA and a two-tailed t-test. Differences with a P value <0.05 were considered statistically significant. Experiments shown, except where indicated, are the means of multiple individual points from multiple separate experiments (± S.E.M.).
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Results and discussion |
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). Incubation of cells with the reactive oxygen species (ROS) quenching agent N-acetyl cysteine suppressed the reduction in PTPase activity and suppressed the activation of ERBB1. These findings argue that radiation can induce ROS to promote activation of plasma membrane receptors (Leach et al. 2001, Sturla et al. 2005). Downstream of growth factor receptor signaling are RAS family signal transducers and proximal to these proteins in the ERK1/2 pathway are the Raf family protein kinases (Dent et al. 1995a,b). Radiation activated Raf-1, but not B-Raf, in A431 cells in a manner that was also suppressed by incubation of cells with N-acetyl cysteine (Fig. 1B). Of note, Raf-1 becomes tyrosine phosphorylated after radiation exposure, whereas B-Raf lacks the sites of Raf-1 tyrosine phosphorylation (Y340/Y341) (Fig. 1B, inset).
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, inset). However, basal levels of phospho-ERK1/2 and phospho-AKT were near identical in both cell lines (Fig. 1C, left-hand side of graphs). Furthermore, in wild-type, but not C10 cells, the majority of Raf-1 protein was membrane associated, whereas in C10, but not wild-type cells, the majority of AKT was membrane associated, that is, membrane location of Raf-1 and AKT also did not correlate with phosphorylation of Raf-1 S259 (Fig. 1C, left-hand side of graphs). Radiation promoted similar-fold reductions in Raf-1 S259 phosphorylation in both cell lines, which again did not correlate well with downstream alterations in ERK1/2 activity in both lines (Fig. 1C, graphical keys). Activation of ERK1/2 followed a similar pattern in A431 cells after irradiation to that noted in HCT116 cells (Figs 1C and 2A). Collectively, these findings argue that ROS play an important role in the activation of ERBB1 and Raf-1 after ionizing radiation exposure, and that Raf-1 tyrosine phosphorylation but not Raf-1 S259 phosphorylation correlates well with the activation of a key downstream target of Raf-1 signaling, the MEK1/2–ERK1/2 pathway.
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for HCT116 carcinoma cells, radiation also appeared to cause two waves of ERK1/2 activation in A431 carcinoma cells; an initial prompt wave that occurred within 5–15 min exposure and a second delayed wave that occurred 90–360 min after exposure (Fig. 2A). In A431 carcinoma cells, the activation of ERBB1 and ERK1/2 5–15 and 90–360 min after exposure was blocked by the tyrphostin inhibitor of ERBB1, AG1478 (Fig. 2A). In A431 cells, neutralization of the paracrine factor-(TGF) suppressed the activation of ERBB1 60–180 min after a 2 Gy exposure (Fig. 2B, section (i)). In HCT116 cells deleted for K-RAS D13 expression, which are stably transfected to express active H-RAS V12 (C10), neutralization of the paracrine factor heregulin suppressed the activation of ERBB3 60–180 min after a 1 Gy exposure (Fig. 2B, section (ii)). Thus, the second wave of ERBB receptor activation after irradiation of carcinoma cells is due to the actions of paracrine ligands; in a cell-type dependent fashion, multiple ligands can play a role in this process. In general agreement with our findings at the level of the ERBB receptors, incubation of tumor cells with neutralizing antibodies against TGF or against heregulin also suppressed the second, but not the first wave, of radiation-induced signaling pathway activation, either ERK1/2 or AKT as shown in the Figure, in multiple carcinoma cell lines (Fig. 2C).
Paracrine factors are synthesized as pro-forms that are cleaved by cell surface metalloproteases; as such, we predicted that media from irradiated cells should contain cleaved/activated TGF or heregulin in a cell-type dependent fashion, and that could be transferred onto unirradiated cells to induce signaling pathway activation. In agreement with this hypothesis, media transferred from irradiated HCT116 (C10) cells promoted heregulin-dependent activation of AKT in unirradiated HCT116 (C10) cells (Fig. 2D). Furthermore, treatment of HCT116 cells in situ with a neutralizing anti-heregulin antibody also suppressed AKT activation in the irradiated cells (compare with data in Fig. 2C). Similarly, in A431 cells, media transferred from these irradiated carcinoma cells promoted TGF-dependent activation of ERK1/2 in unirradiated A431 cells (Fig. 2D). In A431 and DU145 cells, radiation promoted TGF release into the growth media, as measured in an ELISA assay (Fig. 2E, left panel). Release of paracrine factors, e.g. TGF, was also blocked by an inhibitor of MEK1/2, implying that the initial activation of ERK1/2 signaling promoted paracrine ligand release, of note, the initial ERK1/2 activation was ROS-dependent and paracrine ligand-independent. Incubation of cancer cells with an inhibitor of cell surface metalloproteases e.g. GM6001, abolished the ability of media from irradiated tumor cells to promote AKT activation in non-irradiated cells, suggesting that radiation-induced release of a paracrine ligand had been blocked (Fig. 2E, right panel). Collectively, our data in Figs 1 and 2 demonstrate that radiation generates ROS, which inactivates PTPases that permits activation of ERBB receptors, and which activates ERK1/2 and AKT signaling, all within 30 min of exposure (Fig. 2F). The levels of ROS and pathway activities then subside. Subsequently, radiation-induced ERK1/2 signaling causes activation of cell surface metalloproteases which in turn cleave and activate pro-forms of paracrine ligands. The activated paracrine ligands then feedback onto the irradiated carcinoma cell to re-activate ERBB receptors and the ERK1/2 and AKT signaling pathways, 90–360 min after exposure. Of importance for therapeutics is that radiation-induced TGF release has not only been observed using cell lines, in vitro and in vivo, but has also been observed in androgen-independent prostate cancer patients undergoing palliative irradiation for bony metastases (Hagan et al. 2004). Thus, inhibition of ERBB1 and/or MEK1/2 may prevent radiation-induced TGF release in patients; TGF release being an effect that is likely to promote the growth and invasion of unirradiated tumor cells at sites distant to the radiation exposure (Schmidt-Ullrich et al. 1999).
Based on the findings in Figs 1 and 2, it could be simplistically presumed that inhibiting ERBB1 and/or ERK1/2 activation would result in tumor cell radiosensitization. However, while continuous exposure of A431 and DU145 cells for 24 h to MEK1/2 inhibitor did modestly enhance the ability of radiation to promote apoptosis, removal of the drug 6 h after irradiation, abolished the potentiation of apoptosis at 24 h (Fig. 3A). Furthermore, removal of the MEK1/2 inhibitor 24 and 48 h after irradiation abolished the ability of MEK1/2 inhibition to promote cell killing 105 h after exposure in DU145 cells (Fig. 3B, data not shown). These findings argue that for the inhibition of MEK1/2–ERK1/2 signaling to have any radiosensitizing effect in carcinoma cells requires prolonged pathway inhibition. Based on the findings in Fig. 3A and B, colony formation analyses were performed to examine the impact of ERBB1 and MEK1/2 inhibitors on A431, DU145, and MDA-MB-231 radiosensitivity. Inhibition of ERBB1 or MEK1/2 signaling for the 3 h during and after irradiation had the potential to cause radioprotection, that is, ERK1/2 signaling was toxic to cells (Fig. 3C). Removal of the MEK1/2 inhibitor 48 h after exposure had null effect on cell survival, whereas prolonged exposure to MEK1/2 inhibitor promoted radiosensitization when cells were continually cultured in the drug. Collectively, these findings argue that the duration of kinase inhibitor exposure upon a carcinoma cell can dramatically alter the impact on cell survival after radiation exposure.
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, right RT-PCR and immunoblotting panels).
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, left bar graph). The increased expression of ERBB2 and ERBB3 in p53–/– cells may thus also be a compensatory survival response of cells deleted for p53 expression. No change in the expression of RAS proteins or paracrine ligands was observed comparing HCT116 wild-type with HCT116 p53–/– (Fig. 4A, right PCR data, data not shown). Hence, although HCT116 cells (wild-type; C2; C10; p53–/–) are scientifically defined as being ‘isogenic’, our data demonstrate that these cells display considerably different expression profiles of signaling molecules that in other carcinoma cell types can impact on how the tumor cell may subsequently respond to irradiation.
In general agreement with our data in Fig. 4A, the activation of ERK1/2 and AKT following radiation exposure was noted to be very different in wild-type, C2, C10, and p53–/– HCT116 cell lines. In concurrence with several studies examining H-RAS and K-RAS signaling in general, cells expressing K-RAS D13 generated a stronger ERK1/2 activation after irradiation than cells expressing H-RAS V12, whereas cells expressing H-RAS V12 generated a stronger AKT activation after irradiation than cells expressing K-RAS D13 (Fig. 4B and C). Of additional note, in cells lacking p53 expression, basal levels of phospho-ERK1/2 and phospho-AKT were reduced and the activation of ERK1/2 and AKT following radiation exposure was suppressed (Fig. 4B and C; phospho-ERK1/2 reduced to 0.75±0.05; phospho-AKT reduced to 0.89±0.05, of control). The findings with respect to K-RAS and H-RAS activation correlated with wild-type, but not C10, HCT116 cells having Raf-1 translocated to their plasma membrane and C10, but not wild-type, cells having PI3K and AKT associated with their plasma membrane (Fig. 1C). Thus, by translocating either Raf-1 (via K-RAS D13) or PI3K and AKT (via H-RAS V12), it is probable that we have predisposed our wild-type and C10 HCT116 cells to preferentially activate ERK1/2 or AKT respectively.
Thus, in ‘isogenic’ HCT116 cells in Figs 2–4 argue that wild-type HCT116 cells expressing epiregulin, K-RAS D13, and low levels of ERBB1-3, promote a stronger activation of ERK1/2 than AKT after irradiation, whereas in HCT116 (C10) cells expressing heregulin, H-RAS V12 and higher levels of ERBB1 and ERBB3, promote a stronger activation of AKT than ERK1/2 after irradiation. In HCT116 cells lacking p53 function, an approximately 80% reduction in ERBB1 expression correlated with reduced activation of ERK1/2 and AKT following irradiation, suggesting that loss of p53 function may have the potential to either enhance radiosensitivity via loss of AKT and ERK1/2 activation or also promote radioresistance by loss of p53-induced apoptosis after DNA damage.
To further investigate the relative roles of ERBB1 and ERBB2 in HCT116 cell signaling after radiation exposure, cells were incubated with the ERBB1 inhibitor AG1478 or with the ERBB2 inhibitor AG825. Inhibition of ERBB1, but not ERBB2, suppressed radiation-induced tyrosine phosphorylation of ERBB1, ERBB2, and ERBB3 in HCT116 (C10) cells (Fig. 5A). Similarly, incubation with AG1478 abolished radiation-induced activation of ERK1/2 and AKT in HCT116 (C10) cells (Fig. 5B). Our findings in HCT116 (C10) cells are in general agreement with data obtained in other human carcinoma cell lines e.g. wild-type HCT116, A431, MDA-MB-231, and DU145 with respect to ERBB1 activation being a central player in radiation-induced ERK1/2 and AKT signaling responses (Fig. 5C).
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and 5, changes in RAS family member activity and p53 activity would be predicted to alter cellular plating efficiency and radio-sensitivity in in vitro colony formation assays. C10 HCT116 cells, which express H-RAS V12 and exhibit strong radiation-induced AKT activation, were noted to have a greater plating efficiency and be more radio-resistant than wild-type HCT116 cells, which express K-RAS D13 and exhibit strong radiation-induced ERK1/2 activation; both cell lines were more radio-resistant than HCT116 cells lacking expression of any mutated RAS protein (Fig. 6A). Loss of p53 function, in our colony assays, surprisingly resulted in a finding that p53–/– HCT116 cells were modestly more radiosensitive than wild-type HCT116 cells; an effect that we presume is due to a loss of radiation-induced ERK1/2 and AKT activation in this cell line (Fig. 6B). Thus, loss of p53 function in HCT116 cells, which would be presumed to be radioprotective is outweighed in the overall survival response by loss of pro-survival AKT and ERK1/2 signaling. HCT116 wild-type cells, in agreement with this line exhibiting a stronger activation of ERK1/2 than AKT after irradiation, were radio-sensitized by a MEK1/2 inhibitor (Fig. 6C). HCT116 (C10) cells, in agreement with this line exhibiting a stronger activation of AKT than ERK1/2 after irradiation, were radiosensitized by a PI3K inhibitor. In HCT116 (C2) cells, radiosensitization by signaling pathway interruption was achieved only when both MEK1/2 and PI3K were inhibited. This would tend to argue that inhibition of both PI3K and MEK1/2 may best achieve radiosensitization in the widest range of tumor cell types, regardless of RAS protein expression.
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Acknowledgements |
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References |
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P<0.05 the greater survival than corresponding irradiated cell value).
P<0.001 greater than mutant K-RAS-deleted cells value). (B) Identical plating efficiency and colony formation studies to those described in (A) were performed using wild-type HCT116 cells, HCT116 (C2) cells and HCT116 p53–/–cells. (C) HCT116 cells were plated and treated with U0126 or LY294002 as indicated, 24 h after plating. Cells were irradiated (5 Gy). Cells were re-plated as single cells for colony formation assays, 96 h after irradiation. Colonies were permitted to form over the following 10 days. The numbers of colonies were expressed asa fraction of the respective mock-irradiated cells and plotted in a semi-log graph. Data are the means of six separate dishes (S.E.M., n=3) (*P<0.05 less than corresponding vehicle treated cell value;