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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Cancer Res. Author manuscript; available in PMC 2011 January 1.
Published in final edited form as:
PMCID: PMC2805033

Pro-apoptotic activity of bortezomib in gastrointestinal stromal tumor (GIST) cells


Gastrointestinal stromal tumors (GISTs) are caused by activating mutations in the KIT or PDGFRA receptor tyrosine kinase genes. Although more than 85% of GIST patients treated with the small molecule inhibitor imatinib mesylate (Gleevec®) achieve disease stabilization, complete remissions are rare and a substantial proportion of patients develop resistance to imatinib over time. We previously reported that upregulation of soluble, non chromatin-bound histone H2AX has an important role in imatinib-induced GIST cell apoptosis and that H2AX levels, in untreated GIST, are maintained at low levels by a pathway that involves KIT, PI3K, and the ubiquitin-proteasome system. Here, we asked whether bortezomib-mediated inhibition of the ubiquitin-proteasome machinery could lead to upregulation of histone H2AX and GIST cell death. We show that bortezomib rapidly triggers apoptosis in GIST cells through a combination of mechanisms involving H2AX upregulation and loss of KIT protein expression. We demonstrate downregulation of KIT transcription as an underlying mechanism for bortezomib-mediated inhibition of KIT expression. Modulation of the nuclear factor-kappa-B (NF-κB) signaling pathway did not appear to play a major role in bortezomib-induced GIST cell death. Importantly, bortezomib induced apoptosis in two imatinib-resistant GIST cell lines as well as a short-term culture established from an imatinib-resistant GIST. Collectively, our results show that inhibition of the proteasome using bortezomib can effectively kill imatinib-sensitive and imatinib-resistant GIST cells in vitro and provide a rationale to test the efficacy of bortezomib in GIST patients.


GISTs are the most common mesenchymal tumors of the gastrointestinal tract. They are caused by activating mutations in the KIT or PDGFRA receptor tyrosine kinase (RTK) genes (13) and can be effectively treated with the small molecule kinase inhibitor imatinib mesylate (Gleevec®) (4). However, although more than 85% of patients with metastatic GIST benefit from imatinib therapy, complete responses are rare, and the majority of patients develop resistance to imatinib during the course of treatment (46).

Various second- and third-line therapies for GIST are being developed, most of them targeting the oncogenically activated kinases KIT and PDGFRA. However, sunitinib (Sutent®), a multi-targeted compound that inhibits VEGFR, RET, CSF-1R, and FLT3 in addition to KIT/PDGFRA, is so far the only FDA-approved compound for the treatment of imatinib-resistant GIST (79). The most prevalent mechanisms of resistance to imatinib involve additional (secondary) mutations of KIT or PDGFRA that affect the conformation of the kinase domain (1012), and only a subset of these imatinib-resistance mutations are sensitive to sunitinib. The efficacy of sunitinib and other ATP-competitive direct inhibitors of KIT/PDGFRA is further hampered by genomic heterogeneity of resistance mutations within each individual patient. Moreover, virtually all patients receiving imatinib or sunitinib for metastatic inoperable GIST have persistent measurable disease indicating that KIT-inhibitory treatments may induce quiescence rather than apoptosis in a subset of cells as suggested recently (13). It is therefore imperative to identify novel strategies for treating GIST, particularly using drugs that induce GIST cell apoptosis.

We have previously shown that upregulation of soluble histone H2AX induces apoptosis in imatinib-treated GIST cells (14). We found that in untreated GIST cells, H2AX levels are regulated by a pathway that involves KIT, PI3K and the ubiquitin-proteasome machinery. Hence, we ask here whether it would be possible to upregulate levels of histone H2AX to induce GIST cell apoptosis by inhibiting its proteasomal degradation.

Bortezomib (Velcade®) is a dipeptide boronic acid inhibitor of the 20S proteasome that is FDA-approved for the treatment of multiple myeloma and mantle cell lymphoma (1517). The therapeutic activity of bortezomib results, in part, from impeding the degradation of pro-apoptotic factors, thereby inducing programmed cell death in neoplastic cells (18). In multiple myeloma, a key consequence of bortezomib treatment appears to be inhibition of the transcription factor nuclear factor-κB (NF-κB) (16, 19).

In the present study, we tested the activity of bortezomib in GIST cells and found profound pro-apoptotic effects at concentrations similar to those showing activity against multiple myeloma in vitro. Bortezomib treatment increased histone H2AX expression and, unexpectedly, also resulted in a significant downregulation of KIT mRNA levels and subsequent protein expression. Inactivation of the NF-κB signaling pathway, however, did not play a major role in GIST cell apoptosis. Hence, our findings suggest that bortezomib has a dual mode of action against GIST cells involving upregulation of pro-apoptotic histone H2AX and downregulation of oncogenic KIT. Most importantly, bortezomib was active against imatinib-resistant GIST cells and a short-term culture derived from an imatinib-resistant GIST in vitro. Future studies are warranted to test the clinical effectiveness of bortezomib in GIST patients.

Materials and Methods

Cell culture and inhibitor treatments

The human GIST cell line GIST882 was derived from an untreated metastatic GIST and maintained in RPMI1640 supplemented with 15 % fetal bovine serum (FBS, Gemini BioProducts, West Sacramento, CA), 1% L-Glutamine, 50 U/ml penicillin (Cambrex, Walkersville, MD) and 50 μg/ml streptomycin (Cambrex) as described earlier (20). GIST430 and GIST48 were established from GISTs that had progressed during imatinib therapy after initial clinical response. As described previously (21), GIST430 and GIST48 cell lines have heterozygous primary KIT exon 11 mutations accompanied by secondary imatinib-resistance mutations in exons 13 and 17, respectively.

The short-term culture GIST004 was established from a patient with a clinically imatinib-refractory GIST that underwent surgery for removal of a progressing lesion at UPMC Presbyterian Hospital, Pittsburgh, PA (IRB #0509050). The fresh tumor specimen was disaggregated with scalpels and incubated overnight at 37 °C in a solution containing DNase and Collagenase B. Following the enzymatic disaggregation, cells were collected by centrifugation, resuspended in RPMI1640 and plated in 35 mm dishes. After growing to near-confluency, cells were treated as indicated for 72 h and processed for protein extraction. Sequencing of the tumor and short-term culture DNA (22) revealed a primary KIT exon 11 deletion (del550–558) as well as a secondary imatinib-resistance mutation in KIT exon 14 (T670I), also known as the “gatekeeper mutation” (23).

Cells were treated with bortezomib (dissolved in DMSO; LC Laboratories, Woburn, MA) at concentrations ranging from 1 nM to 10 μM for up to 72 h. Imatinib mesylate (kindly provided by Novartis Pharma, Basel, Switzerland) was used at a final concentration of 1 μM in DMSO. α-Amanitin (Sigma, St. Louis, MO) was reconstituted in dH2O and used at concentrations ranging from 0.1 μg/ml to 5 μg/ml for up to 72 h. The I-κB inhibitor Bay 11-7082 (Sigma), the HSP90 inhibitor 17-AAG (EMD, San Diego, CA) as well as the proteasome inhibitors MG-132, MG-262 (both Boston Biochem, Boston, MA), epoxomicin (EMD) and ZL3VS (Biomol, Plymouth Meeting, PA) were used at a final concentration of 1 μM in DMSO for up to 72 h or as indicated. Human recombinant TNF-α (Sigma) was reconstituted in PBS and used at 0.1 μM for 24 h.

Immunological and cell staining methods

Protein lysates of GIST monolayer cultures (23) were prepared by scraping cells into RIPA buffer (1% NP-40, 50 mM Tris-HCl pH 8.0, 100 mM sodium fluoride, 30 mM sodium pyrophosphate, 2 mM sodium molybdate, 5 mM EDTA, 2 mM sodium orthovanadate) containing protease inhibitors (10 μg/ml aprotinin, 10 μg/ml leupeptin, 1 μM phenylmethylsulfonyl fluoride). Lysates were incubated for 1 h with shaking at 4°C and then cleared by centrifugation for 30 min at 14,000 rpm at 4°C. Protein concentrations were determined by the Bradford assay (Biorad, Hercules, CA). 30 μg of protein were loaded on a 4–12% Bis-Tris gel (Invitrogen, Carlsbad, CA) and blotted onto a nitrocellulose membrane.

For immunofluorescence analysis, cells grown on coverslips were briefly washed in PBS and fixed in 4% paraformaldehyde in PBS for 15 min at room temperature (RT). Cells were then washed in PBS and permeabilized with 1% Triton-X 100 in PBS for 15 min (RT) followed by washing in PBS and blocking with 10% normal donkey serum (Jackson Immunoresearch, West Grove, PA) for 15 min (RT). Cells were then incubated with primary antibodies overnight at 4°C in a humidified chamber and incubated for another 3 h at 37°C the next morning. After a brief wash in PBS, cells were incubated with FITC-anti-mouse or FITC-anti-rabbit secondary antibodies (Jackson Immunoresearch) for at least 2 h at 37°C, washed with PBS, and counterstained with 4′,6-diamidino-2-phenylindole (DAPI; Vector Laboratories, Burlingame, CA). Cells were analyzed using an Olympus AX70 epifluorescence microscope equipped with a SpotRT digital camera.

Primary antibodies used for immunoblotting and immunofluorescence were PARP (Invitrogen/Zymed Laboratories, South San Francisco, CA), ubiquitin (BD Biosciences, San Jose, CA), mouse monoclonal phospho-H2AX S139 (Upstate/Millipore, Billerica, MA), H2AX (Bethyl Laboratories, Montgomery, TX), phospho-KIT Y703 (Invitrogen/Zymed), KIT (DakoCytomation, Carpinteria, CA), phospho-RNA polymerase II S2 (Covance, Berkley, CA), phospho-RNA polymerase II S5 (Covance), RNA polymerase II (Covance), CBP (Santa Cruz), p300 (Santa Cruz), phospho-I-κB-alpha S32, I-κB-alpha, phospho-NF-κB p65 S536, NF-κB p65 (all Cell Signaling, Danvers, MA) and actin (Sigma).

Apoptotic cells were visualized using the In situ Cell Death Detection Kit (Roche Applied Sciences, Indianapolis, IN) according to manufacturer’s recommendations.

Reverse Transcriptase (RT)-PCR and quantitative real time RT-PCR

For RT-PCR, cells were plated into p60 dishes, grown to 80% confluence and treated with bortezomib (0.01 μM), α-amanitin (1 μg/ml) and solvent controls for 48 h. RNA was extracted using the RNeasy Mini Kit (Qiagen, Valencia, CA) according to the manufacturer’s protocol. RT-PCR was performed using exon-overlapping, mRNA/cDNA-specific primers to KIT (forward: 5′-TCATGGTCGGATCACAAAGA-3′, reverse: 5′-AGGGGCTGCTTCCTAAAGAG-3′; Operon, Huntsville, AL) and β-actin (forward: 5′-CCAAGGCCAACCGCGAGAAGATGAC-3′, reverse: 5′-AGGGTACATGGTGGTGCCGCCAGAC-3′) and the AccessQuick RT-PCR system from Promega (Madison, WI). Cycling conditions were 50°C (45 min, RT reaction), 92°C (2 min, denaturation), 55°C (30 sec, annealing), 62°C (45 sec, extension), 92°C (30 sec, denaturation) for 20 cycles plus a final annealing step for 2 min at 62°C for KIT amplification or an annealing temperature of 44°C for β-actin, respectively, on an Eppendorf Personal Cycler (Eppendorf, Hamburg, Germany).

For quantitative real time RT-PCR (qRT-PCR), cDNA was transcribed by RT-PCR using random primers and KIT cDNA was amplified and measured using TaqMan® gene expression assays (Applied Biosystems, Softer City, CA) and Real Time PCR system LightCycler (Roche, Grenzach-Wyhlen, Germany). β-Actin cDNA, (also measured using Taqman gene expression assay) served as reference gene for relative quantification.

In vitro Proliferation and Apoptosis Assays, Flow Cytometry

Cell viability studies were performed using the CellTiter-Glo luminescence-based assay (Promega, Madison, WI, USA) as described previously (21). For these studies, the cell lines were plated at 15,000 to 40,000 cells per well in a 96-well flat-bottomed plate (Falcon, Lincoln, NJ), cultured in serum-containing media for 1 to 3 days, and then incubated for 72 h with bortezomib, imatinib or DMSO-only solvent control. Luminescence was measured with a Veritas Microplate Luminometer (Turner Biosystems, Sunnyvale, CA), and the data were normalized to the DMSO-only control group. All experimental points were measured in triplicate wells for each plate and were replicated in at least two plates.

Apoptosis was assessed by measuring caspase-3 and caspase-7 activity using the caspase-Glo 3/7 luminescence-based assay (Promega) as described previously (21). Experimental conditions were as described above for the CellTiter-Glo studies.

For flow cytometry, cells were treated with bortezomib at the indicated concentrations for 24 h or 48 h and resuspended in 100 μl of staining solution containing annexin V FITC (525 nm) and 7-AAD (675 nm) in HEPES buffer. After incubation at room temperature for 15 min, annexin V-positive cells were estimated by flow cytometry, as described previously (21). 10,000 events of each sample were acquired on a Beckman Coulter FC500 Flow Cytometer. Doublet discrimination was done with FL2 vs. FL2 peak histogram.


Bortezomib induces apoptosis in GIST cells

To determine whether bortezomib affects GIST cell proliferation and viability, GIST882 cells were treated with bortezomib at increasing concentrations. Substantial inhibition of proliferation was found at concentrations of 0.01 μM and higher (Fig. 1A). To assess induction of apoptosis, GIST882 cells were treated with 0.001 μM to 10 μM bortezomib for 72 h and immunoblotted for PARP. A PARP cleavage fragment indicating apoptosis was detected in cells treated with concentrations as low as 0.01 μM (Fig. 1B, left panels), which is the same concentration that led to a substantial reduction in cell growth and induction of apoptosis in multiple myeloma cells (16). A 72 h time course experiment using this concentration (0.01 μM) demonstrated that induction of apoptosis is time dependent and started at 24 h after beginning bortezomib treatment (Fig. 1B, right panels). Induction of apoptosis was also time dependent, when evaluated using the TUNEL assay (Fig. 1C). Quantification of apoptotic cells revealed a statistically significant increase of apoptotic cells to 14.5%, 48.9% and 60.6% after 24 h, 48 h and 72 h of treatment, respectively, in comparison to control-treated cells that had corresponding apoptotic indices of 1%, 1%, and 2.8%, respectively (p≤0.002, p≤0.0001, p≤0.0006, respectively; Fig. 1C). Taken together, these results show that bortezomib has profound pro-apoptotic activities in GIST cells.

Figure 1
Bortezomib induces apoptosis in GIST cells

Bortezomib upregulates histone H2AX and causes a rapid loss of KIT expression

To determine the mechanism of bortezomib-induced GIST cell apoptosis, we analyzed H2AX levels by immunoblotting and found that both phosphorylated and total histone H2AX protein expression increased in a dose- and time-dependent manner (Fig. 2). The increase of phospho- and total H2AX levels paralleled the increase in general ubiquitylation of proteins (data not shown). We also detected a second, higher molecular weight band (approximately 24 kDa in size) that most likely corresponds to ubiquitylated histone H2AX indicating that increased levels of H2AX after bortezomib treatment are a possible mechanism of apoptosis induction. When downregulating H2AX by small interfering RNA (siRNA), however, the percentage of surviving cells after bortezomib did not increase to the same extent seen after imatinib treatment (data not shown) (14). These results suggest that additional mechanisms may be involved in bortezomib-induced GIST cell apoptosis.

Figure 2
Bortezomib leads to upregulation of H2AX, but downregulates KIT

In line with this notion, we found that KIT protein expression levels were dramatically downregulated in a dose- and time-dependent manner after bortezomib treatment (Fig. 2). As expected, KIT phosphorylation decreased in parallel.

It is known that most GIST cells are dependent on continued KIT expression for their survival as evidenced by the fact that KIT knock-down by small interfering RNA (siRNA) or downregulation of KIT using the HSP90 inhibitor 17-AAG readily causes GIST cell death (14, 21). Hence, downregulation of KIT could play a substantial role in bortezomib-induced apoptosis in GIST.

Bortezomib induces transcriptional downregulation of KIT

To further dissect the role of proteasome inhibition in bortezomib-induced GIST cell apoptosis, we treated GIST cells with several other proteasome inhibitors. Treatment with all compounds tested (MG-132, MG-262, epoxomicin and ZL3VS), which – like bortezomib – predominantely inhibit the chymotrypsin-like activity of the proteasome, led to an increase of phospho- and total H2AX levels as well as downregulation of KIT expression levels (Suppl. Fig. 1). These results suggest that the effect of bortezomib on GIST cells is most likely mediated by its ability to inhibit the chymotrypsin-like activity of the proteasome.

To analyze the events that are involved in the downregulation of KIT expression, we then tested whether bortezomib leads to transcriptional inhibition of KIT. KIT mRNA levels were substantially downregulated in GIST882 cells that were treated with 0.01 μM bortezomib for 48 h as shown by RT-PCR (Fig. 3A, left panels) and quantitative RT-PCR (qRT-PCR; Fig. 3A, right panel). Remarkably, downregulation of KIT transcripts by 0.01 μM bortezomib as determined by qRT-PCR were comparable to treatment of the cells with 1 μg/ml α-amanitin, a potent and selective inhibitor of RNA polymerase (pol) II.

Figure 3
Bortezomib induces transcriptional inhibition of KIT

We then tested the effects of bortezomib on the transcriptional machinery in GIST cells. Immunoblotting for RNA pol II phosphorylation at serine 2 (S2) of the tandem heptad repeats of its C-terminal domain (CTD) showed a substantial decrease of phosphorylation after bortezomib treatment that was dose- and time-dependent (Fig. 3B, left and middle panels). Phosphorylation of RNA pol II (S2) is associated with transcriptional elongation (24). The same pattern was seen when probing for RNA pol II phosphorylation at serine 5 of the CTD, which occurs predominantly during transcriptional initiation. Inhibition of RNA pol II (S2) after bortezomib could also be visualized by immunofluorescence microscopy (Fig. 3B, right panels) showing loss of RNA pol II (S2) phosphorylation.

To directly test the effect of transcriptional inhibition on KIT expression in GIST cells, we performed a dose-response experiment treating GIST cells with increasing amounts of α-amanitin. As expected, α-amanitin led to a dose-dependent decrease of phosphorylation of RNA pol II (S2) and RNA pol II (S5) (data not shown). Interestingly, α-amanitin treatment resulted in complete loss of KIT protein expression at concentrations of 1 μg/ml and higher indicating that KIT is highly regulated on a transcriptional level (Fig. 3C).

In addition to downregulating RNA pol II, bortezomib also led to downregulation of the transcriptional c o-activator CREB-binding protein (CBP) as shown by immunofluorescence staining (Fig. 3D, left panels). 97.8% of DMSO-treated cells showed positivity for CBP in form of so-called nuclear speckles, whereas only 44.1% of cells were positive when treated with 0.01 μM bortezomib (Fig. 3D, right). Negative cells either showed a complete loss of CBP expression or redistribution of CBP into displacement foci that are associated with global transcriptional downregulation. The effect of bortezomib was comparable to treatment of GIST882 cells with 1 μg/ml α-amanitin (Fig. 3D, right; 100% CBP positive in controls to 39.4% in cells treated with α-amanitin). Taken together, the global displacement of RNA pol II and CBP from chromatin throughout the nucleus suggests that bortezomib leads to a general transcriptional shutdown rather than a to specific loss of RNA pol II from the KIT promoter. Our results therefore indicate that bortezomib causes downregulation of KIT by inhibition of the transcriptional machinery.

Control experiments in which we treated GIST882 cells with imatinib or the HSP90 inhibitor 17-AAG revealed a rapid loss of KIT phosphorylation, but no change of KIT protein levels after imatinib (Suppl. Fig. 2A, left panels). In contrast, 17-AAG not only led to a loss of KIT phosphorylation, but also to a loss of KIT protein expression (21) (Suppl. Fig. 2A, right panels). Treatment with imatinib was associated with an inhibition of ongoing transcription (Suppl. Fig. 2A–C), whereas 17-AAG treatment resulted in an increase of RNA pol II phosphorylation at S5 (Suppl. Fig. 2B, right panels) indicating that bortezomib and imatinib differ substantially from 17-AAG with respect to the effects on the transcriptional machinery. In addition, imatinib led to an increase in the abundance of KIT mRNA, whereas 17-AAG resulted in a more than two-fold decrease of KIT transcripts as determined by qRT-PCR (Suppl. Fig. 2D). The most likely explanation for these seemingly discrepant results are effects of these compounds on posttranscriptional modifications that result in differences in mRNA stability.

To explore the role of NF-κB in bortezomib-induced GIST cell death, we treated GIST882 cells with the I-κB inhibitor Bay 11-7082, which had no effect on apoptosis as judged by the lack of a PARP cleavage fragment (Suppl. Fig. 3A) nor on levels of H2AX or KIT protein expression (Suppl. Fig. 3A). Furthermore, treatment of GIST882 cells with bortezomib had no effect on NF-κB signaling as measured by I-κB-alpha and NF-κB phosphorylation in whole cell lysates (Suppl. Fig. 3B). Most importantly, NF-κB p65 was cytoplasmic at baseline levels indicating that it is not constitutively active in GIST882 cells (Suppl. Fig. 3C, D). Treatment with either bortezomib or Bay 11-7082 did not lead to a change in subcellular localization of NF-κB over a period of 72 hours (Suppl. Fig. 3C, D). By contrast, stimulation with TNF-α readily resulted in a nuclear translocation of NF-κB (Suppl. Fig. 3C, D) thus indicating that the pathway is nevertheless functional in GIST882 cells.

Collectively, these findings underscore that bortezomib has unique effects on KIT mRNA levels, global transcriptional activity and KIT protein levels when compared to imatinib or 17-AAG, and that modulation of the NF-κB pathway does not play a major role in bortezomib-induced apoptosis in GIST cells.

Bortezomib induces apoptosis and downregulates KIT in imatinib-resistant GIST cells

We next determined the efficacy of bortezomib in imatinib-resistant GIST cells. GIST48 and GIST 430 are cell lines derived from imatinib-resistant human GISTs (21). Treatment of these cells with bortezomib at concentrations ranging from 0.001 μM to 0.25 μM led to a significant reduction of cellular proliferation with IC50s of 0.003 μM and 0.002 μM for GIST48 and GIST430, respectively (Fig. 4A, B; top left panels). By contrast, the IC50 was not reached when treating GIST48 or GIST430 with imatinib at concentrations ranging from 0.001μM to 10 μM (Fig. 4A, B; top panels). A dramatic, 5-to 12-fold increase in apoptosis was seen in GIST48 and GIST430 cells after 48 h of treatment at concentrations of 0.005 to 0.02 μM (Fig. 4A,B; bottom left panels). These data were confirmed by flow cytometry and annexin V staining at the same concentrations and time points (Fig. 4A,B; right panels).

Figure 4
Bortezomib induces apoptosis and downregulates KIT in imatinib-resistant GIST cells

These preclinical validations of bortezomib as a novel therapeutic option in imatinib-resistant GIST were extended through evaluations in primary cultures established from an imatinib-resistant GIST. Mutational analysis of this tumor revealed a primary KIT exon 11 deletion (del550–558) and a secondary imatinib-resistance mutation in exon 14 (T670I). Short-term cultures were treated with 0.01 μM bortezomib or 1 μM imatinib for 72 h. As shown by immunoblotting, bortezomib induced apoptosis in these cells as indicated by PARP cleavage, whereas imatinib had only a minor effect (Fig. 4C). Moreover, levels of phospho- and total H2AX increased after bortezomib treatment (data not shown). In concordance with our previous results, KIT and phospho-KIT (Y703) were downregulated after bortezomib treatment, whereas levels of KIT protein as well as KIT phosphorylation were largely unchanged when treated with imatinib (Figure 4C).


Although GISTs can be successfully treated with imatinib mesylate, new therapeutic options are needed, because complete responses are rare and more than 80% of the patients develop resistance to the drug over time (46). Based on our previous findings that histone H2AX, which is causatively involved in the induction of apoptosis after imatinib, is regulated by the ubiquitin-proteasome machinery (14), we tested the possibility of using the FDA-approved proteasome inhibitor bortezomib for treating GIST. We found that bortezomib had a potent pro-apoptotic effect on GIST cells at the same concentrations used in multiple myeloma models, a disease where bortezomib has clinical benefit (15, 16). Notably, the bortezomib-induced GIST apoptosis was preceded not only by upregulation of histone H2AX, but also by downregulation of KIT oncogene mRNA levels.

Our current study supports the hypothesis that upregulation of histone H2AX by inhibition of the proteasome can induce apoptosis. However, downregulation of H2AX by siRNA only led to a moderate increase in surviving cells after bortezomib treatment (data not shown), suggesting that additional mechanisms are involved in the induction of apoptosis. Our finding that bortezomib treatment of GIST cells resulted in significant downregulation of KIT protein, accompanied by loss of KIT tyrosine phosphorylation lends important support to this notion. It is known that loss of the KIT protein has a strong apoptotic effect in GIST cells as seen after knocking down KIT with siRNA or treatment with HSP90 inhibitors (14, 21). It is therefore likely that downregulation of KIT contributes importantly to the effect of bortezomib on GIST cells. Interestingly, downregulation of BCR-ABL in CML cells has been described after treatment with various proteasome inhibitors in vitro (25). Although the underlying mechanism was not demonstrated, transcriptional inhibition was suggested as a potential explanation (25).

It is known that GISTs are especially prone to transcriptional inhibition (26). We have shown previously that treatment with the RNA polymerase II inhibitor α-amanitin led to an apoptotic response in GIST882, whereas non-transformed fibroblasts were not affected (14). We show here that bortezomib causes a transcriptional downregulation of KIT (Fig. 3A). We show that bortezomib treatment is associated with an inhibition of the global transcriptional machinery including downregulation of RNA pol II phosphorylation as well as loss of the transcriptional co-activator CBP from chromatin throughout the nucleus. These findings indicate that bortezomib leads to a general transcriptional shutdown rather than a to specific loss of RNA pol II from the KIT promoter. Interestingly, it has been shown in BCR-ABL positive CML cells that combination treatment of bortezomib with flavoperidol leads to transcriptional inhibition and loss of phosphorylation of the CTD of RNA pol II (27). However, the effect of either agent, given singly, was less pronounced (27). The mechanisms by which bortezomib downregulates KIT oncogene transcription are incompletely understood. It is known that the 26S proteasome is associated with transcriptionally active genes and physically interacts with RNA pol II (28). In fact, several studies have shown that the ubiquitin-proteasome machinery is involved in regulation of transcriptional activation (29, 30), with initial binding and subsequent proteasomal degradation of transcriptional activators being necessary to initiate transcription and transcriptional elongation (3134).

The dichotomous effect that bortezomib has on H2AX and KIT expression levels can most likely be explained by differences in the mechanism of protein degradation between H2AX and KIT. H2AX is processed by the ubiquitin-proteasome system, and its levels increase after treatment with a proteasome inhibitor. However, the phosphorylation-dependent degradation of KIT is mediated by the lysosome (35). This process is unlikely to be inhibited by compounds that target the ubiquitin-proteasome machinery. Hence, bortezomib treatment does not affect ongoing KIT degradation, which – together with reduced transcription (Figure 3) – results in a significant loss of KIT protein expression.

Inactivation of NF-κB through retention in the cytoplasm is thought to be a key mechanism of bortezomib-induced apoptosis in multiple myeloma (16, 19). We provide evidence that inhibition of the NF-κB signaling pathway does not appear to play a major role in bortezomib-mediated GIST cell death (Suppl. Fig. 3).

It is likely that transcriptional inhibition of KIT expression is not the only mechanism leading to apoptosis in GIST cells after bortezomib treatment. We are currently testing whether bortezomib is involved in the regulation of chaperone proteins, such as HSP90 and HSP70, as well as chromatin modifiers, such as histone deacetylases, that could also result in the downregulation of KIT protein. Interestingly, bortezomib has been demonstrated to modulate the abundance of DNMT1, a DNA methyl transferase, in acute myeloid leukemia cells leading to changes in DNA methylation and gene expression (36). Although increased DNA methylation would not explain the transcriptional downregulation of KIT, a change in methylation patterns could lead to the re-expression of other, cell cycle inhibitory or pro-apoptotic genes that were epigenetically silenced in GIST cells thus providing additional anti-proliferative or cytotoxic stimuli.

Because most imatinib-resistant GISTs develop secondary mutations within the KIT or PDGFRA kinase domains, novel therapeutic approaches that do not directly target these kinases are particularly important. The example of HSP90 inhibitors, which likewise lead to loss of KIT oncoprotein expression in GIST, supports this notion (21). In that respect, our results that bortezomib is effective against GIST cells harboring various resistance mutations, including the so-called gatekeeper mutation, seem especially promising.

Further support for this notion comes from preliminary experiments, in which we treated KitK641E transgenic mice (37) with bortezomib. After two or four weeks of treatment some resected tumors showed loss of Kit expression, downregulation of cyclin A and upregulation of cleaved caspase 3 by immunoblotting of whole cell lysates (data not shown). These promising results indicate that bortezomib indeed has an effect in vivo. However, a larger cohort of animals needs to be treated for a longer period of time to confirm these preliminary results.

In summary, we have demonstrated that bortezomib induces apoptosis in imatinib-sensitive as well as imatinib-resistant GIST cells. We identified a dual mode of action for bortezomib, including stabilization of histone H2AX as well as transcriptional downregulation of KIT. Our results provide a compelling rationale for clinical trials to test the efficacy of bortezomib in GIST patients.

Supplementary Material


Supplemental Figure 1. Effect of various proteasome inhibitors on expression levels of phospho- and total H2AX as well as phospho- and total KIT.

Immunoblot analysis of GIST882 cells treated with MG-132, MG-262, epoxomicin and ZL3VS (all 1 μM in DMSO) for 24 h and probed for phospho-H2AX (S139), total H2AX, phospho-KIT (Y703) and total KIT. Immunoblot for actin is shown to demonstrate loading. Treatment of GIST882 with bortezomib (0.01 μM, 24 h) is shown as comparison. Bo, bortezomib.


Supplemental Figure 2. Effect of imatinib and 17-AAG on transcription.

(A) Immunoblot analysis of GIST882 cells treated with imatinib (left panels) or 17-AAG (right panels) at 1 μM for the indicated times and probed for phospho-KIT (Y703), KIT, phospho-RNA pol II (S2), phospho-RNA pol II (S5) and total RNA pol II. Immunoblot for actin is shown to demonstrate loading.

(B) Immunofluorescence microscopic analysis of GIST882 cells treated with DMSO, imatinib (1 μM) or 17-AAG (1 μM) for 48 h and stained for CBP. Nuclei stained with DAPI.

(C) Quantification of the percentage of GIST882 cells showing CBP displacement foci, which are associated with a global loss of transcription, after treatment with imatinib (1 μM) or 17-AAG (1 μM) for 48 h (bars, mean and standard error of at least 100 cells of two independent experiments).

(D) Quantitative RT-PCR (qRT-PCR) of KIT mRNA after treatment of GIST882 with DMSO, imatinib (1 μM) or 17-AAG (1 μM) for 48 h (graph on right). Values are normalized against β-actin mRNA.


Supplemental Figure 3. Effect of bortezomib on NF-κB signaling.

(A) Immunoblot analysis of GIST882 cells treated with the I-κB inhibitor Bay 11-7082 at 1μM for the indicated times and probed for phospho-KIT (Y703), total KIT, phospho-H2AX (S139) and total H2AX. Immunoblot for actin is shown to demonstrate loading. Treatment of GIST882 with imatinib (IM; 1 μM, 72 h) or bortezomib (Bo; 0.01 μM, 72 h) is shown as comparison. Bo, bortezomib.

(B) Immunoblot analysis of GIST882 cells treated with bortezomib at 0.01 μM for the indicated times and probed for phospho-I-κB-alpha (S32), total I-κB-alpha, phospho-NF-κB p65 (S536) and total NF-κB p65. Immunoblot for actin is shown to demonstrate loading.

(C) Immunofluorescence microscopic analysis of GIST882 cells treated with DMSO, TNF-α (0.1 μM), bortezomib (0.01 μM) or Bay 11-7082 (1 μM) for 24 h and stained for NF-κB p65. Nuclei stained with DAPI.

(D) Quantification of the percentage of GIST882 cells showing nuclear NF-κB p65 after treatment with DMSO, TNF-α (0.1 μM), bortezomib (0.01 μM; upper panel) or Bay 11-7082 (1 μM; lower panel) for the indicated time points (bars, mean and standard error of at least 100 cells of three independent experiments).


The authors would like to thank Christopher L. Corless, Oregon Health and Science University, Portland, OR, for providing the mutational analysis of GIST004 and Saumendra Sarkar, University of Pittsburgh Cancer Institute, Pittsburgh, PA, for helpful discussions and sharing important reagents. Imatinib mesylate was generously provided by Novartis Pharma, Basel, Switzerland. This work was supported by a Research Scholar Grant from the American Cancer Society (RSG-08-092-01-CCG; to A.D), the GIST Cancer Research Fund (to A.D. and J.A.F.), The Life Raft Group (to A.D., S.B., B.P.R. and J.A.F), the Deutsche Krebshilfe (to S.B.), the Virginia and Daniel K. Ludwig Trust for Cancer Research (to J.A.F.) and the National Institutes of Health (1P50CA127003-02; to J.A.F.). A.D. is supported by the University of Pittsburgh Cancer Institute and in part by a grant from the Pennsylvania Department of Health. The Department specifically disclaims responsibility for any analyses, interpretations or conclusions.


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