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Prior studies have noted that inhibitors of MEK1/2 enhanced geldanamycin lethality in malignant hematopoietic cells by promoting mitochondrial dysfunction. The present studies focused on defining the mechanism(s) by which these agents altered survival in carcinoma cells. MEK1/2 inhibitors (PD184352; AZD6244 (ARRY-142886)) interacted in a synergistic manner with geldanamycins (17AAG, 17DMAG) to kill hepatoma and pancreatic carcinoma cells that correlated with inactivation of ERK1/2 and AKT and with activation of p38 MAPK; p38 MAPK activation was ROS-dependent. Treatment of cells with MEK1/2 inhibitors and 17AAG reduced expression of c-FLIP-s that was mechanistically connected to loss of MEK1/2 and AKT function; inhibition of caspase 8 or over-expression of c-FLIP-s abolished cell killing by MEK1/2 inhibitors and 17AAG. Treatment of cells with MEK1/2 inhibitors and 17AAG caused a p38 MAPK-dependent plasma membrane clustering of CD95 without altering the levels or cleavage of FAS ligand. In parallel, treatment of cells with MEK1/2 inhibitors and 17AAG caused a p38 MAPK-dependent association of caspase 8 with CD95. Inhibition of p38 MAPK or knock down of BID, FADD or CD95 expression suppressed MEK1/2 inhibitor and 17AAG lethality. Similar correlative data were obtained using a xenograft flank tumor model system. Our data demonstrate that treatment of tumor cells with MEK1/2 inhibitors and 17AAG induces activation of the extrinsic pathway and that suppression of c-FLIP-s expression is crucial in transduction of the apoptotic signal from CD95 to promote cell death.
In the United States, hepatoma is diagnosed in ~ 19,000 patients per annum with ~ 17,000 deaths from the disease, with a 5 year survival rate of less than 10%. Hepatoma is a leading cause of diagnosed cancer in Africa and Asia and represents the fifth most commonly diagnosed malignancy in the World (1). In the United States, pancreatic cancer is diagnosed in ~ 37,000 patients per annum with ~ 34,000 deaths every year (2). Pancreatic cancer has a 5 year survival rate of less than 5%. These statistics emphasize the need to develop novel therapies against these lethal malignancies.
The Raf/mitogen-activated protein kinase (MAPK) kinase 1/2 (MEK1/2)/extracellular signal–regulated kinase 1/2 (ERK1/2) pathway is frequently dysregulated in neoplastic transformation, including hepatocellular carcinoma (3). The MEK1/2-ERK1/2 module comprises, along with c-Jun NH2-terminal kinase (JNK1/2) and p38 MAPK, members of the MAPK super-family (4, 5). These kinases are involved in responses to diverse mitogens and environmental stresses, including DNA damage, osmotic stress, and hypoxia, among others, and have also been implicated in multiple cellular functions, including proliferation, differentiation, and cell survival processes. Although exceptions exist, activation of the ERK1/2 pathway is generally associated with cell survival whereas induction of JNK1/2 and p38 MAPK pathways generally signals apoptosis. There is also evidence that the net balance of signals in terms of amplitude and duration between the cytoprotective ERK1/2 and the stress-related JNK1/2 and p38 MAPK pathways determines whether a cell lives or dies following various insults. Although the mechanism(s) by which ERK1/2 activation promotes survival is not known with certainty, several downstream anti-apoptotic effector proteins have been identified, including direct inactivation of pro-apoptotic proteins such as caspase-9, BAD and BIM, and increased expression of anti-apoptotic proteins such as BCL-XL, MCL-1 and c-FLIP proteins (6–11). In view of the importance of the MEK1/2-ERK1/2 pathway in neoplastic cell survival, MEK1/2 inhibitors have been developed by several pharmaceutical companies and have entered clinical trials, including PD184352 (CI-1040), the second generation Pfizer MEK1/2 inhibitor PD 0325901 and the Astra Zeneca drug AZD6244 (ARRY-142886) (12, 13).
Heat shock protein 90 (HSP90) is a chaperone protein involved in the proper folding and intracellular disposition of multiple proteins involved in cell signaling and survival (14, 15). Tumor cells generally have higher rates of protein synthesis than non-neoplastic cells and disruption of HSP90 function in tumor cells (e.g., by benzoquinoid ansamycin antibiotics such as geldanamycin (16)) has been shown to induce improper folding of diverse proteins, including Raf-1, B-Raf, AKT, ERBB family receptors, among numerous others, culminating in their proteasomal degradation (17). These events have been shown to induce apoptosis or, alternatively, to increase the susceptibility of tumor cells to established cytotoxic agents (18, 19). Such considerations have led to the development of clinically relevant HSP90 antagonists, such as 17-allylamino-17-demethoxygeldanamycin (17AAG), which has both superior pharmacokinetic and reduced normal tissue toxicity characteristics compared with geldanamycin (20, 21). Many studies have argued that inhibition of the PI3 kinase – AKT pathway, rather than the Raf-MEKl/2-ERKl/2 pathway, represents a key component of 17AAG toxicity and sensitization effects in tumor cells (22–27). Free plasma concentrations of 17AAG in patients have been noted to be in the low 1 to 5 μmol/L range for up to 12 h after drug infusion, which is significantly higher than the required concentration of drug to inhibit HSP90 function (25, 26).
The goal of the present studies was to determine whether, and by what mechanism(s), clinically relevant MEK1/2 inhibitors might enhance the activity of clinically relevant geldanamycins (17AAG, 17DMAG) against human hepatoma and other GI and GU tumor cells in vitro and in vivo. Our results indicate that clinically relevant MEK1/2 inhibitors interact synergistically with 17AAG and 17DMAGto induce CD95 (FAS receptor) –dependent cell death.
Total BAX, cleaved caspase 3, Phospho-/total-ERKl/2/5, Phospho-/total-JNKl-3, Phospho-/total-p38 MAPK, Anti-S473 AKT and total AKT antibodies were purchased from Cell Signaling Technologies (Worcester, MA). Active BAX specific antibody (6A7) for immunoprecipitation was purchased from Sigma (St. Lois, MO). The c-FLIP-s/L and all the secondary antibodies (anti-rabbit-HRP, anti-mouse-HRP, and anti-goat-HRP) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The JNK inhibitor peptide (JNK IP), caspase inhibitors (zVAD, IETD, LEHD) and 17AAG was supplied by Calbiochem (San Diego, CA) as powder, dissolved in sterile DMSO, and stored frozen under light-protected conditions at −80°C. Enhanced chemiluminescence (ECL) kits were purchased from Amersham Enhanced ChemiLuminescence (ECL) system (Bucks, England) and NEN Life Science Products (NEN Life Science Products, Boston, MA). Trypsin-EDTA, RPMI medium, penicillin-streptomycin were purchased from GIBCOBRL (GIBCOBRL Life Technologies, Grand Island, NY). BAX/BAK −/−, BIM −/− and BID −/− fibroblasts were kindly provided by Dr. S. Korsmeyer (Harvard University, Boston, MA). HuH7, HEPG2 and HEP3B (hepatoma), pancreatic (PANC1, Mia Paca2), colorectal (SW480, HCT116), and prostate (DU145, PC3) cancer cells were obtained from the ATCC (Rockville, MD). Commercially available validated short hairpin RNA molecules to knock down RNA/protein levels were from Qiagen (Valencia, CA): CD95 (SI02654463; SI03118255); FADD (SI00300223; SI03648911); BID (SI02654568; SI02661911). The dominant negative p38 MAPK and activated MEK1 EE recombinant adenoviruses were kindly provided by Drs. K. Valerie, VCU and J. Moltken (University of Cincinnati), respectively. The proprietary drug 17DMAG was supplied by the Dr. David Gius, Radiation Oncology Branch, Radiation Oncology Sciences Program, National Cancer Institute, National Institutes of Health, Bethesda, Bethesda, MD. Other reagents were of the highest quality commercially available (11, 27, 28).
All established cell lines were cultured at 37 °C (5% (v/v CO2) in vitro using RPMI supplemented with 5% (v/v) fetal calf serum and 10% (v/v) Non-essential amino acids. For short-term cell killing assays and immunoblotting, cells were plated at a density of 3 × 103 per cm2 and 36 h after plating were treated with various drugs, as indicated. In vitro small molecule inhibitor treatments were from a 100 mM stock solution of each drug and the maximal concentration of Vehicle (DMSO) in media was 0.02% (v/v). For adenoviral infection, cells were infected 12 h after plating and the expression of the recombinant viral transgene allowed to occur for 24 h prior to any additional experimental procedure. Cells were not cultured in reduced serum media during any study.
Unless otherwise indicated in the Figure Legend, cells were treated with either vehicle (VEH, DMSO), or the combination of MEK1/2 inhibitor PD184352 (PD184; 1 (μM) or PD98059 (PD98; 20 μM) as indicated, and geldanamycin (17AAG; 1 μM or 17DMAG; 0.25 (.μM) or both agents combined. For SDS PAGE and immunoblotting, cells were lysed in either a non-denaturing lysis buffer, and prepared for immunoprecipitation as described in (27, 28) or in whole-cell lysis buffer (0.5 M Tris-HCl, pH 6.8, 2% SDS, 10% glycerol, 1% β-mercaptoethanol, 0.02% bromophenol blue), and the samples were boiled for 30 min. After immunoprecipitation, samples were boiled in whole cell lysis buffer. The boiled samples were loaded onto 10–14% SDS-PAGE and electrophoresis was run overnight. Proteins were electrophoretically transferred onto 0.22 μm nitrocellulose, and immunoblotted with indicated primary antibodies against the different proteins. All immunoblots were visualized by ECL. For presentation, immunoblots were digitally scanned at 600 dpi using Adobe PhotoShop CS2, and their color removed and Figures generated in Microsoft PowerPoint. Densitometric analysis for E.C.L. immunoblots were performed using a Fluorochem 8800 Image System and the respective software (Alpha Innotech Corporation, San Leandro, CA) and band densities were normalized to that of a total protein loading control.
We generated and purchased previously noted recombinant adenoviruses to express constitutively activated and dominant negative AKT and MEK1 proteins, dominant negative caspase 9, the caspase 9 inhibitor XIAP, the endogenous caspase 8 inhibitor c-FLIP-s, the polyoma virus caspase 8 inhibitor CRM A, and mitochondrial protective protein BCL-XL (Vector Biolabs, Philadelphia, PA) (27). Unless other wise stated, cells were infected with these adenoviruses at an approximate multiplicity of infection (m.o.i.) of 50. As noted above, cells were further incubated for 24 h to ensure adequate expression of transduced gene products prior to drug exposures.
Approximately 10 nM of a defined pre-validated siRNA (Ambion technologies) was diluted into 50 μl growth media lacking FBS and pen-strep. Based on the Manufacture’s instructions, an appropriate amount of Lipofectamine 2000 reagent (usually 1 μl) (Invitrogen, Carlsbad, CA) was diluted into a separate vial containing media with lacking FBS or pen-strep. The two solutions were incubated separately at room temperature for 5 min, then mixed together (vortexed) and incubated at room temperature for 30 min. The mixture was added to each well (slide or 12-well plate) containing an appropriate amount (~ 0.5 ml) of pen-strep- and FBS-free medium. Cells were incubated for 2–4 h at 37 deg C with gentle rocking. Media was then replaced with 1 ml of 1× pen-strep and FBS containing media.
Plasmid DNA (0.5 μg/total plasmid transfected) was diluted into 50 μl of RPMI growth media that lacked supplementation with FBS or with penicillin-streptomycin. Lipofectamine 2000 reagent (1 μl) (Invitrogen, Carlsbad, CA) was diluted into 50 μl growth media that lacked supplementation with FBS or with penicillin-streptomycin. The two solutions were then mixed together and incubated at room temperature for 30 min. The total mix was added to each well (4-well glass slide or 12-well plate) containing 200 μl growth media that lacked supplementation with FBS or with penicillin-streptomycin. The cells were incubated for 4 h at 37°C, after which time the media was replaced with RPMI growth media containing 5% (v/v) FBS and 1× pen-strep (27, 28).
Cells were harvested by trypsinization with Trypsin/EDTA for ~10 min at 37 °C. As some apoptotic cells detached from the culture substratum into the medium, these cells were also collected by centrifugation of the medium at 1,500 rpm for 5 min. The pooled cell pellets were resuspended and mixed with trypan blue dye. Trypan blue stain, in which blue dye incorporating cells were scored as being dead was performed by counting of cells using a light microscope and a hemacytometer. Five hundred cells from randomly chosen fields were counted and the number of dead cells was counted and expressed as a percentage of the total number of cells counted. For confirmatory purposes the extent of apoptosis was evaluated by assessing Hoechst and TUNEL stained cytospin slides under fluorescent light microscopy and scoring the number of cells exhibiting the “classic” morphological features of apoptosis and necrosis. For each condition, 10 randomly selected fields per slide were evaluated, encompassing at least 1500 cells. Alternatively, the Annexin V/propidium iodide assay was carried to determine cell viability out as per the manufacturer’s instructions (BD PharMingen) using a Becton Dickinson FACS can flow cytometer (Mansfield, MA) (27, 28).
Athymic female NCr-nu/nu mice were obtained from Jackson Laboratories (Bar Harbor, ME). Mice were maintained under pathogen-free conditions in facilities approved by the American Association for Accreditation of Laboratory Animal Care and in accordance with current regulations and standards of the U.S. Department of Agriculture, Washington, DC, the U.S. Department of Health and Human Services, Washington, DC, and the National Institutes of Health, Bethesda, MD. HEP3B cells were cultured and isolated by trypsinization followed by cell number determination using a hemacytometer. Cells were resuspended in phosphate buffered saline and ten million tumor cells per 100 μl PBS were injected into the right rear flank of each mouse, and tumors permitted for form to a volume of ~100 mm3 over the following 3–4 weeks. PD184352 was prepared and administered IP three times daily as described in Hawkins et al (6). The geldanamycin 17AAG was prepared in an identical manner to PD184352 and administered once daily. Both agents were dosed at 25 mg/kg for 30 hours.
Animals were euthanized by CO2 and placed in a BL2 cell culture hood on a sterile barrier mat. The bodies of the mice were soaked with 70% (v/v) EtOH and the skin around the tumor removed using small scissors, forceps and a disposable scalpel. These implements were flame sterilized between removal of the outer and inner layers of skin. A piece of the tumor (~50% by volume) was removed and placed in a 10 cm dish containing 5 ml of RPMI cell culture media, on ice. In parallel the remainder of the tumor was placed in 5 ml of Streck Tissue Fixative (Fisher Scientific, Middletown VA) in a 50 ml conical tube for H&E fixation. The tumor sample that had been placed in RPMI was minced with a sterile disposable scalpel into the smallest possible pieces then placed in a sterile disposable flask. The dish was rinsed with 6.5 ml of RPMI medium which was then added to the flask. A 10× solution of collagenase (Sigma, St. Louis MO; 2.5 ml, 28 U/ml final concentration) and 10× of enzyme mixture containing DNAse (Sigma, St. Louis MO; 308 U/ml final concentration) and pronase (EMD Sciences, San Diego CA; 22,500 U/ml final concentration) in a volume of 1 ml was added to the flask. The flasks were placed into an orbital shaking incubator at 37°C for 1.5 hours at 150 rpm. Following digestion, the solution was passed through a 0.4 μM filter into a 50 ml conical tube. After mixing, a sample was removed for viable and total cell counting using a hemacytometer. Cells were centrifuged at 500 × g for 4 min, the supernatant removed, and fresh RPMI media containing 10% (v/v) fetal calf serum was added to give a final resuspended cell concentration of 1 × 106 cells/ml. Cells were diluted and plated in 10 cm dishes in triplicate at a concentration of 2 × 103 cells/dish for control, and for all other drug exposures 4 × 103 cells/dish.
Fixed tumors were embedded in paraffin wax and 10 μM slices obtained using a microtone. Tumor sections were de-parafinized, rehydrated and antigen retrieval in a 10 mM (w/v) Na Citrate/Citric acid buffer (pH 6.7) heated to 90 °C in a constant temperature microwave oven. Prepared sections were then blocked and subjected to imunohistochemistry as per the instructions of the manufacturer for each primary antibody (P-AKT (S473); P-p38; P-ERK1/2; cleaved caspase 3; c-FLIP-s). The permanently mounted slides were allowed to dry overnight and were photographed at the indicated magnification. The area selected for all photo-micrographs was the proliferative zone, within 2 mm of, or juxtaposed to leading edge of the tumor.
Cells were harvested after GST-MDA-7 treatment by centrifugation at 600 rpm for 10 min at 4°C and washed in PBS. Cells (~1 × 106) were lysed by incubation for 3 min in 100 μl of lysis buffer containing 75 mM NaCl, 8 mM NaH2PO4, 1 mM NaH2PO4, 1 mM EDTA, and 350 μg/ml digitonin. The lysates were centrifuged at 12,000 rpm for 5 min, and the supernatant was collected and added to an equal volume of 2X Laemmli buffer. The protein samples were quantified and separated by 15% SDS PAGE (27, 28).
Comparison of the effects of various treatments was performed using one way analysis of variance and a two tailed Student’s f-test. Differences with a p-value of < 0.05 were considered statistically significant. These values were determined using the statistical programming within SigmaStat and SigmaPlot. Median dose effect isobologram analyses to determine synergism of drug interaction were performed according to the Methods of T-C Chou and P Talalay using the Calcusyn program for Windows (BIOSOFT, Cambridge, UK). A combination index (CI) value of less than 1.00 indicates synergy of interaction between the drugs; a value of 1.00 indicates additivity; a value of > 1.00 equates to antagonism of action between the agents. Data points from all experiments shown are the mean of multiple individual data points summated from the stated number of multiple experiments i.e. (mean data shown = Σ n, all data points, ± SEM) (27, 28).
Initial experiments focused on the regulation of hepatoma and pancreatic carcinoma cell survival following exposure to MEK1/2 inhibitors (PD184352 (CI-1040), AZD6244 (ARRY-142886)) and the geldanamycin 17AAG. Treatment of HuH7, HEPG2 and HEP3B (hepatoma) cells with 17AAG and PD184352 caused a greater than additive induction of cell killing than either individual agent alone within 48h of exposure, as judged in TUNEL, trypan blue and annexin - propidium iodide flow cytometry assays (Figures 1A and 1B). Similar data to that with PD184352 were obtained when the MEK1/2 inhibitor AZD6244 was used (Figure S1). Similar hepatoma cell killing data to that obtained with 17AAG were generated when the HSP90 inhibitor 17DMAG was used in combination with the MEK1/2 inhibitor PD184352; cell killing was blocked by the small molecule caspase 8 inhibitor IETD (Figure 1C) (see also data in reference 28). Using median dose effect analyses we determined using short term cell death and long term colony formation assays whether MEK1/2 inhibitors and 17AAG interacted in a synergistic manner: both PD184352 and AZD6244 enhanced 17AAG lethality in a synergistic manner with combination index (CI) values of less than 1.00 (Figure 1D, data not shown). Similar cell killing data to that generated in hepatoma cells were also observed when pancreatic (PANC1, Mia Paca2), colorectal (SW480, HCT116), prostate (DU145, PC3) and breast (MCF7, MDA-MB-231) cancer cells were treated with 17AAG and the MEK1/2 inhibitor PD184352 (data not shown).
The molecular mechanisms by which MEK1/2 inhibitors and 17AAG interacted to kill hepatoma cells were next investigated in greater detail. Inhibition of caspase 9 function suppressed cell killing and abolished the greater than additive induction of cell killing by MEK1/2 inhibitors and 17AAG (Figure 2A). Inhibition caspase 8 function (using the viral CRM A protein) blocked pro-caspase 9 and pro-caspase 3 cleavage and virtually abolished cell killing by MEK1/2 inhibitors and 17AAG (Figure 2A; Figure S2). We next utilized SV40 Large T antigen transformed mouse embryonic fibroblasts that had been genetically modified to lack expression of pro-apoptotic proteins. MEK1/2 inhibitors and 17AAG enhanced cell killing in wild type cells, whereas killing was significantly reduced in cells lacking expression of BAX, BAK, BIM and BID (Figure 2B). As inhibition of caspase 8 and loss of BID function negatively impacted on MEK1/2 inhibitor and 17AAG -induced killing, we performed additional studies to define the relative role of caspase 8, and molecules upstream of caspase 8 that regulate its function, in the observed drug-induced cell killing process.
Over-expression of the caspase 8 inhibitor c-FLIP-s significantly reduced cell killing caused by MEK1/2 inhibitor and 17AAG treatment in hepatoma and pancreatic carcinoma cells (Figures 3A and 3B). Over-expression of c-FLIP-s abolished the synergistic interaction between PD184352 (CI-1040) or AZD6244 (ARRY-142886) and 17AAG in true colony formation assays (Table S1 and data not shown). Similar colony survival data were also obtained in Panc1 and Mia Paca2 cells (data not shown). In agreement with data in Figure 2 showing that caspase 9 and BAX/BAK/BIM function also played a role in MEK1/2 inhibitor and 17AAG lethality, over-expression of the mitochondrial protective protein BCL-XL or the caspase 9 inhibitor XIAP suppressed cell killing. Treatment of HEP3B cells with MEK1/2 inhibitor and 17AAG caused cleavage of pro-caspase 8 and the pro-apoptotic protein BID, and decreased expression of the caspase 8 inhibitor c-FLIP-s, effects that were prevented by constitutive over-expression of c-FLIP-s (Figure 3B).
Pro-caspase 8 is generally thought to be activated by binding to the FAS associated death domain (FADD) protein which associates in a “DISC” with trimerized/activated death receptors such as TRAIL (DR4/DR5), TNFα or FAS (CD95) (29). Previous studies by this laboratory in primary hepatocytes have strongly linked bile acid toxicity, and its promotion by inhibitors of MEK1/2, to ligand independent activation and plasma membrane localization of CD95 (11, 30, 31). Knock down of BID, FADD or CD95 expression significantly reduced MEK1/2 inhibitor and 17AAG lethality in hepatoma cells (Figures 4A and 4B; Figure S3). Treatment of hepatoma cells with MEK1/2 inhibitor and 17AAG caused enhanced association of pro-caspase 8 with CD95 in immunoprecipitates of CD95 (DISC formation) and reduced the association of c-FLIP-s with CD95 (Figure 4C, upper panel). Treatment of hepatoma cells with MEK1/2 inhibitor and 17AAG caused release of cytochrome c into the cytosol from the mitochondria and decreased mitochondrial levels of cytochrome c; an effect that was suppressed by knock down of CD95 expression (Figure 4C, lower panel).
Based on prior studies in primary hepatocytes with bile acids and CD95 activation, we determined whether treatment of hepatoma cells with MEK1/2 inhibitor and 17AAG elevated the plasma membrane levels/surface density of CD95, indicative of CD95 activation. Treatment of hepatoma cells with PD184352 and 17AAG visibly increased plasma membrane staining for CD95 in HEP3B cells and in HEPG2 cells, an effect that we were also able to quantitate (Figure 4D, data not shown). Collectively these findings demonstrate that treatment of hepatoma cells with MEK1/2 inhibitors and 17AAG promotes CD95 activation, DISC formation with caspase 8 association, and extrinsic pathway activation which leads to BID cleavage, mitochondrial dysfunction, and cell death.
Further studies then attempted to define the changes in signal transduction pathway function which were causal in the regulation of the extrinsic pathway in cells treated with MEK1/2 inhibitors and 17AAG. Combined exposure of hepatoma cells to MEK1/2 inhibitor and 17AAG resulted in a rapid phosphorylation of p38 MAPK within 3h and lasting for ~24h; a rapid dephosphorylation of ERK1/2 over 3h–24h; and a slower modest secondary decline in AKT (S473) phosphorylation that occurred over 6h–24h (Figure 5A). Of note, at the concentration of PD184352 used (1 μM) in our studies, ERK1/2 phosphorylation was not completely suppressed over 24h, The JNK1/2 pathway was not activated under our culture/treatment conditions (data not shown). The changes in signaling pathway activity approximately correlated with the prolonged reduced expression of c-FLIP-s, BCL-XL and XIAP, which was in general agreement with our prior data showing that over-expression of c-FLIP-s, BCL-XL and XIAP protected hepatoma cells from MEK1/2 inhibitor and 17AAG treatment.
We next determined whether constitutive activation of MEK1 and/or AKT could suppress the toxic interaction between 17AAG and the MEK1/2 inhibitor PD98059. PD98059 was chosen for these studies because unlike PD184352 and AZD6244, it is a relatively poor inhibitor of the constitutively activated MEK1 EE protein. Combined expression of activated MEK1 and activated AKT, but not either protein individually, maintained ERK1/2 and AKT (S473) phosphorylation in the presence of the MEK1/2 inhibitor PD98059 and 17AAG and suppressed drug-induced phosphorylation of p38 MAPK (Figure S4). In HEPG2 cells expression of constitutively active AKT more strongly suppressed the lethality of 17AAG and MEK1/2 inhibitor treatment than expression of constitutively active MEK1 whereas in HEP3B cells both constitutively active AKT and constitutively active MEK1 were apparently equally competent at blunting drug toxicity (Figure 5B and Figure S5). In both hepatoma cell types, combined expression of constitutively active AKT and constitutively active MEK1 almost abolished 17AAG and PD98059 -induced cell killing. Expression of constitutively active AKT and constitutively active MEK1 maintained the expression levels of c-FLIP-s and well as those of XIAP and BCL-XL in cells treated with 17AAG and PD98059 (Figure 5B, upper inset immunoblotting panel).
As noted in Figure 5A, the p38 MAPK pathway was rapidly activated within 3h after combined exposure to 17AAG and MEK1/2 inhibitor prior to complete inactivation of ERK1/2 and AKT that occurred 6–12h after exposure, suggesting that even though activated MEK1 and activated AKT can suppress drug-induced p38 MAPK activation, the activation of p38 MAPK was likely to be independent of drug-induced ERK1/2 and AKT inactivation (Figure 5 A; Figure S4). Combined expression of dominant negative MEK1 and dominant negative AKT reduced the phosphorylation of ERK1/2 and AKT, but did not profoundly increase the phosphorylation of p38 MAPK (Figure 5C, blots to the left). Combined expression of dominant negative MEK1 and dominant negative AKT reduced the expression of c-FLIP-s and BCL-XL, but did not significantly enhance basal levels of cell morbidity (Figure 5B; data not shown). Expression of dominant negative MEK1 recapitulated the effects of PD184352 in terms of enhancing 17AAG-stimulated p38 MAPK phosphorylation and enhancing 17AAG-stimulated killing (data not shown). These findings argue that the drug 17AAG must provide an additional “signal” separate from simply suppressing ERK1/2 and AKT function, which is required to cause p38 MAPK activation and to promote tumor cell killing.
Prior studies from this laboratory have demonstrated that reactive oxygen species (ROS) are an important component of 17AAG lethal signaling, including the activation of p38 MAPK (3, 27). Exposure of hepatoma cells to the ROS quenching agent N-acetyl cysteine, that suppresses ROS induction in hepatoma cells, did not significantly modify the inactivation of ERK1/2 or AKT by 17AAG and MEK1/2 inhibitor treatment but did suppress the activation of p38 MAPK by these drugs (Figure 5C, blots to the right; data not shown (27)). Exposure of hepatoma cells to the ROS quenching agent N-acetyl cysteine significantly reduced the lethality of 17AAG and MEK1/2 inhibitor treatment (data not shown). Collectively, the data in Figure 5 argues that loss of ERK1/2 and AKT function and gain of p38 MAPK function play important roles in the lethal actions of 17AAG and MEK1/2 inhibitor treatment in hepatoma cells.
Based on our data in Figure 5A, which demonstrated that p38 MAPK was rapidly activated after combined exposure to 17AAG and MEK1/2 inhibitor, we further investigated whether this signaling pathway played any direct role in the regulation of CD95 and the extrinsic pathway following drug treatment. Exposure of cells to 17AAG and PD184352 increased the association of pro-caspase 8 with CD95 in hepatoma cells (DISC complex formation); an effect that was inhibited by expression of dominant negative p38 MAPK or by expression of dominant negative MKK3 and dominant negative MKK6 (Figure 6A, section (i)). Expression of dominant negative p38 was competent to inhibit stress-induced signaling in this pathway (data not shown). Expression of activated AKT and activated MEK1 also suppressed 17AAG and MEK1/2 inhibitor -induced association of pro-caspase 8 with CD95 (Figure 6A, section (ii)). Expression of neither dominant negative p38 MAPK nor activated AKT and activated MEK1 altered the whole cell expression levels of either CD95 or of FAS ligand (Figure S6). This suggests CD95 activation was p38 MAPK dependent and FAS ligand-independent.
Expression of dominant negative p38 visibly suppressed the drug-induced plasma membrane staining for CD95, which was quantified (Figure 6B). Expression of dominant negative p38 MAPK, but not inhibition of the JNK1/2 pathway, suppressed 17AAG and MEK1/2 inhibitor –induced cell killing in HEPG2 and HEP3B cells (Figure 6C and Figure S7, data not shown). The data in Figure 6A argued that inhibition of p38 MAPK prevented the association of pro-caspase 8 and CD95. MEK1/2 inhibitor and 17AAG-induced activation of BAX and BAK, proteins that act downstream of CD95 to cause mitochondrial dysfunction, was also shown to be p38 MAPK dependent (Figure 6C, upper inset panel). Thus 17AAG and MEK1/2 inhibitors, from a signal transduction standpoint, interact to kill human hepatoma cells in vitro by suppressing AKT and ERK1/2 activity and by activating p38 MAPK, and these pathways regulate cell survival both at the level of CD95 and at the level of the mitochondrion, within the tumor cell.
Finally, as both 17AAG and MEK1/2 inhibitors are under evaluation in the clinic, we tested whether our in vitro findings could be translated into animal model systems. We noted that unselected clones of HEP3B and HEPG2 cells are poorly tumorigenic in the flanks of athymic mice and form tumors that rapidly become necrotic upon growth beyond > 200 mm3, potentially due to a relatively low CD31 staining (data not shown). As such, we chose an in vivo treatment, ex vivo colony formation assay approach to assess tumor cell killing and long-term survival, as well as immunohistochemical parameters. HEP3B tumors exposed to PD184352 and 17AAG in vivo had a lower ex vivo cell colony forming ability than tumor cells exposed to either agent individually that correlated with increased caspase 3 cleavage and reduced phosphorylation of ERK1/2 and AKT in the tumor, and increased p38 MAPK phosphorylation (Figure 6D). The expression of c-FLIP-s was also reduced in HEP3B tumors exposed to 17AAG and PD184352 that were undergoing apoptosis, arguing that this protein is both mechanistically linked to modulation of the killing process in vitro and in vivo, and that c-FLIP-s expression could be used as a surrogate marker for tumor responsiveness to this drug combination in vivo.
Prior in vitro studies from our laboratories in chronic myelogenous leukemia (CML) cells have noted that inhibitors of MEK1/2 enhanced geldanamycin lethality by promoting mitochondrial dysfunction (28). The present studies focused more precisely on defining the mechanism(s) by which these agents altered cell survival in hepatoma and pancreatic cancer cells in vitro.
Our findings demonstrated that combined exposure of tumor cells to 17AAG and MEK1/2 inhibitors (PD98059; PD184352; AZD6244) promoted inhibition of the ERK1/2 and AKT pathways and activation of the p38 MAPK pathway. The reduced activity within the ERK1/2 and AKT pathways lowered the cell death threshold of hepatoma cells at multiple points within the extrinsic and intrinsic apoptosis pathways as judged by suppressed protein levels of c-FLIP-s, BCL-XL and XIAP, whose reduced levels of expression could be rescued by molecular activation of AKT and MEK1. Drug-induced activation within the p38 MAPK pathway was a pro-apoptotic stimulus as judged by p38 MAPK-dependent: CD95 localization in the plasma membrane; CD95 association with pro-caspase 8; and activation of BAX and BAK. Loss of MEK1/2 and AKT pathway function reduced c-FLIP-s expression and in parallel facilitated activation of p38 MAPK. Without suppression of c-FLIP-s levels activation of CD95 was incapable of promoting caspase 8 activation/tumor cell killing, regardless of downstream BAX and BAK activation and inhibition of BCL-XL and XIAP expression. This argues that modulation of c-FLIP-s levels represented a key nodal point proximal to CD95 death receptor activation for the manifestation of 17AAG and MEK1/2 inhibitor toxicity in tumor cells (Figure S8).
HSP90 antagonists, of which the ansamycin analogue geldanamycin and its less toxic derivatives, 17AAG and 17DMAG, represent the prototypes, have become a focus of considerable interest as anti-neoplastic agents, and clinical trials involving 17AAG and 17DMAG have been initiated over the last 5–10 years (e.g. 21). These agents act by disrupting the chaperone function of HSP90, leading to the ultimate proteasomal degradation of diverse signal transduction regulatory proteins implicated in the neoplastic cell survival, including Raf-1, B-Raf, AKT, and ERBB family receptors. Mutant active kinase proteins, including activated B-Raf and Bcr-Abl have been noted to be particularly susceptible to agents that disrupt HSP90 function (e.g. 20). The basis for the tumor cell selectivity of 17AAG is not definitively known however there is evidence that HSP90 derived from tumor cells has an increased affinity for geldanamycins compared with HSP90 protein obtained from normal cells (32). One difficulty with the development of 17AAG has been the limited water solubility of this drug and an analogue of 17AAG, 17DMAG, which is considerably more water-soluble than 17AAG, has been synthesized. MEK1/2 inhibitors were previously shown to enhance the lethality of DMAG in CML cells and evidence from our present analyses indicates that PD184352 also enhances 17DMAG lethality in human hepatoma cells (28).
Whilst some hepatoma tumors have been noted to express mutated active forms of Ras and B-Raf proteins, the penetrance of such mutations within the hepatoma patient population as a whole has not been noted to be as prevalent as the well described high mutational rate of these proteins found in other G.I. malignancies such as pancreatic adenocarcinoma or colorectal carcinoma (33, 34). Of note, however, is that 17AAG and MEK1/2 inhibitors interact to kill pancreatic carcinoma cells. Mutations in PI3 kinase and loss of PTEN function/expression in hepatoma have also been noted (35, 36). These findings would suggest that the lethal interaction of 17AAG with MEK1/2 inhibitors we observe in HuH7, HEPG2 and HEP3B hepatoma cells or in other unrelated epithelial tumor cell types is unlikely to be due to a simple suppression of a small subset of hyper-activated HSP90 client proteins as would be predicted based on expression of, for example, mutated active B-Raf or K-RAS. In contrast to pancreatic or colorectal malignancies, virally induced cancers e.g. by hepatitis B virus, the HEP3B cell line is an example, are more prevalent in liver cancers and the key transforming protein of HBV, pX, has been shown by many groups, including this laboratory, to increase the activities of the ERK1/2, AKT and JNK1/2 pathways and enhance the expression of cell cycle regulatory proteins such as p16, p21 and p27 in primary hepatocytes in a dose-dependent manner (37–39). At present there are no published studies indicating whether pX is an HSP90 client protein. Based on the concept of oncogene addiction, however, hepatoma cells such as HEP3B expressing pX could in theory have higher basal levels of ERK1/2 and AKT activity which would in turn make them more susceptible to cell death processes following inhibition of these signal transduction pathways by 17AAG and MEK1/2 inhibitor exposure. Further studies will be required to determine definitively whether HBV infected hepatoma isolates are more sensitive to the 17AAG and MEK1/2 inhibitor drug combination than those lacking transforming HBV proteins.
The Raf-MEKl/2-ERKl/2 pathway exerts cytoprotective actions in a wide variety of transformed cell types which has lead to the development of multiple pharmacologic inhibitors of the pathway, including inhibitors of Ras farnesylation and geranylgeranylation, the multi-kinase and Raf inhibitor Sorafenib and the MEK1/2 inhibitors PD184352, PD0325901 and AZD6244 (40–42). PD184352 has undergone clinical evaluation in phase I and phase II trials involving patients with advanced malignancies and inhibition of ERK1/2 phosphorylation in tumor tissues and peripheral blood mononuclear cells was observed at higher drug doses indicating that achieving desired pharmacodynamic effects in vivo was feasible. However, the relative pharmacodynamic profile of PD1843 52 was not considered to be optimal and as a single agent the drug did not generate any objective tumor growth delay responses in a phase II trial (43). More potent MEK1/2 inhibitors with superior pharmacokinetic characteristics (PD0325901, AZD6244) are currently undergoing clinical evaluation and encouragingly our present studies demonstrated that AZD6244 and 17AAG were competent to interact in a synergistic fashion to kill tumor cells via an extrinsic pathway-dependent mechanism. Studies beyond the scope of the present manuscript will be required to determine whether PD0325901 and AZD6244 can interact with DMAG in vitro and in vivo to kill human hepatoma and other carcinoma cell types.
We noted that administration of low concentrations of PD184352 or of 17AAG in hepatoma cells resulted in an initial abrogation of ERK1/2 phosphorylation, followed by a gradual recovery towards vehicle control treated levels. On the other hand, co-administration of PD184352 and 17AAG resulted in the profound and sustained dephosphorylation of ERK1/2 throughout the entire measured 24h exposure interval. Similarly, only under conditions of drug co-administration was a more modest AKT (S473) dephosphorylation observed. In view of evidence that the duration of MEK/ERK and AKT signaling plays a critical role in the biological consequences of activation of these pathways it is tempting to speculate that sustained inactivation of both ERK1/2 and AKT signaling partially contributes to the lethality of the PD184352 and 17AAG drug regimen in these cells.
The relative roles of ERK1/2 versus AKT inactivation in the promotion of cell killing by 17AAG and MEK1/2 inhibitor treatment were also noted to be slightly different comparing HEPG2 and HEP3B cells. In HEPG2 cells, expression of constitutively active MEK1 did not significantly protect cells from 17AAG and MEK1/2 inhibitor toxicity whereas expression of activated AKT reduced toxicity by ~50%. In HEPG2 cells expression of activated MEK1 in the presence of activated AKT, however, abolished 17AAG and MEK1/2 inhibitor toxicity. In HEP3B cells, both activated MEK1 and activated AKT each approximately equally contributed to suppressing cell killing induced by17AAG and MEK1/2 inhibitor exposure. There are many examples of this form of cell behavior where in some cell types survival is mediated primarily by the actions of one pathway with a secondary or non-existent protective role for other pathways, and in others where survival is shared between many pathways. In hepatocytes/hepatoma cells, the regulation of c-FLIP protein expression has been linked to both the ERK1/2 and AKT pathways (e.g. 11, 44). Thus in the majority of malignancies, based on tumor cell heterogeneity within the tumor, the likelihood that specific inhibition of only one signaling module will achieve a measurable prolonged therapeutic effect will probably be small, which may explain why even when ERK1/2 phosphorylation was significantly suppressed in patient tumors in the presence of PD184352, little benefit was clinically observed. As 17AAG will inhibit not only the ERK1/2 and AKT pathways, and in the presence of a MEK1/2 inhibitor act to cause prolonged suppression of pathway function, but will, furthermore, also reduce the stability of additional cytoprotective HSP90 client proteins such as HIE la, our data argue that the simultaneous targeting of multiple protective pathways by 17AAG and MEK1/2 inhibitors may represent a ubiquitous and better approach to kill cancer cells (45). In a similar vein to reliance on one pathway for a major cellular effect, resistance to 17AAG and MEK1/2 inhibitor exposure could in theory be mediated by reduced expression levels of the death receptor CD95; indeed, HuH7 cells, which have very low expression of CD95 and were relatively resistant to drug exposure killing, compared to HEPG2 and HEP3B cells (46).
Geldanamycins are known to have the capacity to generate reactive oxygen species in G.I. tumor cells (47); prior studies from our laboratory have also shown 17AAG to induce ROS in primary hepatocytes and hepatoma cells (3, 27, 48). Our data argued that ROS production was a key component in p38 MAPK activation after 17AAG and MEK1/2 inhibitor exposure, together with suppression of ERK1/2 and AKT activity. As AZD6244 has recently been shown to reduce hepatoma growth in vivo, collectively, with our present findings, including our in vivo data using HEP3B, and in Mia Paca2 cells (Unpublished findings), it is tempting to speculate that the 17AAG and MEK1/2 inhibitors could have in vivo potential as a therapeutic tool in the treatment of hepatoma and pancreatic cancer (49). Additional studies of will be required to determine whether and how 17AAG and/or 17DMAG and MEK1/2 inhibitors interact in vivo to suppress tumor cell viability and growth.
PD thanks Dr. Shigang Lin for performing some of the initial work on these studies.
Support for the present study was provided; to P.D. from PHS grants (R01-DK52825, P01-CA104177, R01-CA108325), and The Jim Valvano “V” foundation; to S.G. from PHS grants (R01-CA63753; R01-CA77141) and a Leukemia Society of America grant 6405-97; to PBF from PHS grants (P01-CA104177, R01-CA097318; R01-CA098172; P01-NS031492), and The Samuel Waxman Cancer Research Foundation; to DTC from PHS grant (P01-CA104177). P.D. is The Universal Inc. Professor in Signal Transduction Research and P.B.F. is a SWCRF Investigator.