Trabectedin Is Active against Malignant Pleural Mesothelioma Cell and Xenograft Models and Synergizes with Chemotherapy and Bcl-2 Inhibition In Vitro

Malignant pleural mesothelioma (MPM) represents a highly lethal malignancy of the mesothelium with increasing incidence, which is in the vast majority of cases caused by asbestos exposure (1). It has been estimated that within the next 30 years 250,000 people in Europe will die of MPM, and 2,500–3,000 new cases are diagnosed each year in the United States (2, 3).The long latency period (even 20–40 years) between the first exposure to asbestos and the diagnosis of MPM suggests that multiple genetic alterations are needed for malignant transformation of the mesothelial cells (4). There exist three histologic subtypes of MPM, epithelioid, sarcomatoid, and biphasic, the latter two being especially refractory to current therapy options (5, 6). The expected median survival of patients suffering from MPM is between 4 and 12 months (7). This adverse prognosis is also caused by the distinct resistance against systemic therapy options. Hence, the current preferred chemotherapy (pemetrexed/cisplatin) prolongs median survival only by a few months (8). Also aggressive cytoreductive therapy, including macroscopic complete resection by extrapleural pneumonectomy or pleurectomy/decortication in combination with chemotherapy and radiotherapy (termed trimodality therapy), has been shown to prolong survival only in selected patients with early MPM (6). However, due to the limited diagnostic options and treatment modalities at the disseminated stage, better understanding of the molecular factors contributing to the development of MPM are urgently needed to identify new molecular therapy targets.

Recently, the anticancer potential of marine-based cytotoxic compounds came into focus and is currently evaluated in various malignancies (9). Trabectedin (also called ET743, tradename: Yondelis) is an antineoplastic substance originally isolated from the Caribbean marine tunicate Ecteinascidia turbinate (10) exerting considerable antitumor activity in murine and human tumors in vitro and in vivo including melanoma, non–small cell lung cancer (NSCLC), ovarian cancer, as well as anaplastic meningioma (11–15). Recently, a preliminary pharmacokinetic study in xenograft models has been published suggesting selective accumulation of trabectedin in MPM tissue (16). Trabectedin was active in early clinical trials against several solid tumors, including melanoma, NSCLC, breast, prostate, and renal cancers (17). It has been approved for treatment of advanced soft tissue sarcoma (STS) and in patients with platinum-sensitive, recurrent ovarian cancer in combination with PEGylated liposomal doxorubicin (18, 19).

The detailed mode of action of trabectedin is not fully understood and the antitumor effects of the drug are based on multiple mechanisms. First, it binds into the minor DNA groove and interacts with repair mechanisms. Interestingly, the nucleotide excision repair (NER) pathway, known to protect against cisplatin-induced DNA adducts (20), sensitizes against trabectedin (21). In addition, trabectedin selectively interferes with the activity of several oncogenic transcription factors and, consequently, blocks expression of several oncogenes and multidrug resistance factors (22). Besides targeting the malignant cells, trabectedin might also interfere with the cancer microenvironment (17, 21) and especially depletion of tumor-associated macrophages might be central to its anticancer effects (23). However, the preclinical activity of trabectedin in MPM has not been investigated in detail so far. Therefore, we aimed to test in vitro and in vivo activity against MPM cell lines, primary cell cultures, and a tumor xenograft. Moreover, the feasibility of a combination setting with the standard anti-MPM chemotherapeutics and experimental bcl-2 inhibitors was evaluated.

SPC111 and SPC212 cells (from biphasic MPM) were kindly provided by Prof. R. Stahel (Zurich, Switzerland). The cisplatin-resistant subline P31res1.2 was established from the parental P31 by in vitro cisplatin selection (ref. 24; kindly donated by Prof. K. Grankvist, Umea, Sweden). Other cell lines obtained from international sources were: M38K (provided by Prof. V.L. Kinnula, Helsinki, Finland) and I2 (Prof. A. Catania, Milan, Italy). The MPM primary cell cultures VMC6, 12, 14, 20, 23, 28, 33, 40, and 45 were established at the Institute of Cancer Research, as well as Meso49, 53, 62, and 80 at the Division of Thoracic Surgery, both Medical University of Vienna (Vienna, Austria; ref. 25). NP2 and NP3 cell lines were established from nonmalignant pleural tissue samples obtained during pneumothorax operations. Cell line sources and histologic subtypes of the respective tumors are listed in Supplementary Table S1. Cell line P31 and its cisplatin-resistant derivative P31res1.2 were maintained in minimal essential medium (MEM) with 10% heat-inactivated FBS. All other cell lines were cultured in RPMI medium with 10% FBS. Cell lines were kept in a humidified atmosphere with 5% CO₂ and were regularly checked for Mycoplasma contamination. Cell lines were authenticated by array comparative genomic hybridization or DNA fingerprinting.

Trabectedin was obtained from Zelita/Pharmamar and cisplatin from Sigma. The bcl-2 protein inhibitors obatoclax and ABT-199 were purchased from Selleck Chemicals Inc.

To determine cell viability, 3-(4,5 dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT)-based survival assays (EZ4U; Biomedics) were performed as published previously (25–27). Mesothelioma cell cultures were seeded in 96-well plates at a density of 2 × 10³/100 μL per well 24 hours before drug treatment. Drug exposure time was 72 hours. Interactions between drugs were tested on the basis of calculating the combination index (CI) according to Chou and Talalay (28) with CalcuSyn software (Biosoft). CI values <0.9, from 0.9 to 1.1, or >1.1 represented synergism, additive effects, and antagonism of the two investigated substances, respectively.

Cells were seeded into 6-well plates at densities of 2–5 × 10³ cells/well 24 hours before treatment. After 7 days, the drug-containing medium was removed and cell clones were washed, fixed, and stained as published previously (29). The results were quantified with ImageJ software.

For the spheroid growth assay, 5 × 10³ cells were seeded in triplicate in serum-free DMEM/Ham F-12 medium (Biochrom) supplemented with 20 ng/mL basic fibroblast growth factor (bFGF, Eubio, Austria), 20 ng/mL EGF (Sigma) and 2% B27 supplement (PAA Laboratories) in ultra-low attachment 24-well plates (Corning) as described before (26). Trabectedin was added at 1 or 5 nmol/L. Ninety-six hours later, the entire wells were photographed and all spheroids measured with ImageJ software. Only spheroids with a diameter over 100 μm were evaluated to exclude cell clumps from the analysis. Self-renewal potential was tested by disintegration of spheroids to single-cell solution after treatment with trypsin (0.1%) and EDTA (0.01%) for 10 minutes. Cells were then resuspended in DMEM/FBS medium, washed once in serum-free medium, and passed through a 70-μm filter (BD Biosciences) followed by replating in spheroid growth medium in ultra-low attachment plates as described above.

MPM cells were seeded onto 6-well plates at a density of 5 × 10⁵ per well and, following 24-hour drug exposure, collected and lyzed. Proteins were extracted and processed for SDS-PAGE and immunoblotting was performed as published previously (30). The following primary antibodies were used: PARP (detecting full-length and cleaved PARP), cleaved PARP, bax, bim, bak, Mcl-1, bcl-2, bcl-x_L, cyclin A2, and cyclin E1 (all from Cell Signaling Technology), p53 (DO1; Neomarkers), cyclin B1, cyclin D1 (both from Santa Cruz Biotechnology) and β-actin (Sigma) at 1:1,000 dilutions in phosphate-buffered saline with 3% BSA. Horseradish peroxidase–coupled anti-rabbit and anti-mouse antibodies (3% BSA, Santa Cruz Biotechnology) were used at 1:10,000 dilutions and developed by Western blotting with Luminol Reagent Kit (Santa Cruz Biotechnology).

For apoptosis induction experiments, cells were seeded on 24-well plates and treated with trabectedin. After 48 hours of treatment, cells were fixed with 4% buffered paraformaldehyde and labeling of terminal deoxynucleotidyl transferase–mediated dUTP nick end labeling (TUNEL) was performed according to the manufacturer's instructions (Roche Diagnostics). Quantification was done by direct counting of the TUNEL-positive cells in at least five 20× microscopic fields.

SPC111, SPC212, P31, and P31res1.2 cells (2 × 10⁴ cells/well) were seeded in 24-well plates and allowed to recover for 24 hours. Cells were treated with trabectedin, cisplatin, obatoclax, and their combinations for the indicated exposure times. Next, cell culture supernatants were supplemented with 2 μL/mL Hoechst 33258 /propidium iodide (HOE/PI) mix [Hoechst 33258 (1 mg/mL) and PI (2.5 mg/mL)] for at least 1 hour at 37°C. While Hoechst 33258 stains DNA and is taken up by living and dead cells, PI enters dead cells only. Consequently, blue Hoechst 33258 fluorescence allows identification of cells in mitosis or early apoptosis (condensed chromatin but PI-negative), while PI positivity indicates dead cells and, in connection with chromatin condensation in Hoechst staining, late apoptosis. Live photomicrographs of control and treated cells were taken after 24 and 48 hours using fluorescent equipment on the Nikon Eclipse Ti inverted microscope system. Four optical fields per well of two experiments performed in duplicate were captured with 4,6-diamidino-2-phenylindole (DAPI), Cy3 and phase-contrast filters settings, and evaluated (mitosis, early/late apoptosis).

Cells were seeded onto 6-well plates at a density of 2 × 10⁵ cells per well 24 hours before drug treatment. After 48 hours of exposure, the cells were trypsinized, pelleted, and processed for cell-cycle analyses by PI staining and flow cytometry using fluorescence-activated cell sorting (FACSCalibur; Becton Dickinson) according to the standard protocols. The results were evaluated using ModFit software (Becton Dickinson).

For measurement of transmigration through a porous membrane, cells were seeded in density of 4 × 10⁴ cells into Boyden chambers with 8-μm pore size. Subsequently, these chambers were placed into 24-well plates containing 1 mL medium with 10% FBS. After seeding, cells were treated with trabectedin for 24 hours. Thereafter, chambers were removed and treatment medium was replaced by fresh medium with 10% FBS. Those cells which had been migrated to the bottom of the wells were further incubated for additional 7 days and stained with crystal violet. Finally, 2% SDS was used to resolubilize the dye and absorbance was measured at 562 nm on a microplate reader.

Cells (5 × 10⁵/well) were seeded into 6-well plates. When confluency was reached, scratches were applied with a pipette tip in the shape of a cross. After washing and medium replacement, the indicated drug concentrations were added. Wells were photographed after 0, 4, 8, 24, and 48 hours and closure of the artificial gap was calculated from the microphotographs using ImageJ software (NIH, Bethesda, MD)

Total RNA was isolated by TRIzol/chloroform (Life Technologies), and quantity as well as integrity of the RNA samples was checked on an Agilent 2100 Bioanalyzer. Gene expression arrays were performed using 4 × 44K (Design 014850) whole genome oligonucleotide-based gene expression arrays (Agilent) as published previously (27). Feature extraction and data analysis were carried out using the Feature Extraction, GeneSpring, and Ingenuity Pathway Analysis (IPA) software packages. Data analysis with the GeneSpring software (Agilent Technologies) was performed using the default parameters (Guided Workflow).

Six- to 8-week-old female CB-17 scid/scid (SCID) mice were purchased from Harlan Laboratories (San Pietro al Natisone). The animals were kept in a pathogen-free environment and every procedure was done in a laminar airflow cabinet. The experiments were performed according to the regulations of the Ethics Committee for the Care and Use of Laboratory Animals at the Medical University Vienna (proposal numbers BMWF-66.009/0157-II/10b/2008 and BMWF-66.009/0084-II/3b/2013), the U.S. Public Health Service Policy on Human Care and Use of Laboratory Animals as well as the United Kingdom Coordinating Committee on Cancer Prevention Research's Guidelines for the Welfare of Animals in Experimental Neoplasia.

SPC111 cells (1 × 10⁶) in 100 μL serum-free medium were injected intraperitoneally into SCID mice (in total n = 10) as published previously (26). Three weeks after inoculation, mice were assigned into treatment groups, each consisting of 5 mice: (1) treated with 0.9% NaCl; (2) trabectedin (0.15 mg/kg dissolved in 0.9% NaCl) as a single shot treatment. All substances were applied intravenously via tail vein injection. Performance status and body weight were checked every second day. The experiment was terminated after 36 days from transplantation when the first animal had to be sacrificed (by cervical dislocation after anesthesia with 5 mg/kg Rompun and 100 mg/kg Ketavet) due to signs of distress. Tumor nodules were harvested by a thoracic surgeon (M.A. Hoda) using magnifying spectacles blinded with regard to trabectedin/solvent treatment. All nodules from each mouse were pooled and weighed.

Unless stated otherwise, data are presented as means ± SD of at least three experiments performed in triplicate. Statistical significance between treatment groups was analyzed with Prism 5.0 (GraphPad) or SPSS17 software (SPSS Inc.) using Student t test, χ² test, or one-way ANOVA with Tukey multiple comparison test as appropriate. In all cases, P < 0.05 was considered statistically significant.

Initially, trabectedin responsiveness of international epithelioid (P31 and the cisplatin-resistant subline P31res1.2) and biphasic (SPC111, SPC212) MPM cell models as well as normal pleural cell cultures (established from pneumothorax patients) was tested in short term (72-hour MTT assay) and long-term (10 days clonogenic assay) exposure tests. Trabectedin was active against all MPM cell lines at low nanomolar or even picomolar concentrations (Fig. 1A; Supplementary Table S1) and even 0.1 nmol/L suppressed clonogenicity by more than 50% (Fig. 1B). In comparison, 2.5 μmol/L cisplatin were needed to reduce clonogenicity to a comparable level in SPC111 and SPC212 cells. The parental P31 cell model was even more responsive to 0.1 nmol/L trabectedin than to 2.5 μmol/L cisplatin treatment while acquired cisplatin resistance of P31res1.2 cells did not alter trabectedin response. On the basis of these initial promising results, two different panels of primary MPM cell cultures established from patient biopsies or surgical specimens were tested (epithelioid and nonepithelioid in Supplementary Fig. S1A and S1B, respectively). Viability of all malignant cell models was reduced by trabectedin with IC₅₀ values ≤ 2.6 nmol/L and frequently even in the picomolar range (Supplementary Table S1). In contrast, primary cultures of nonmalignant mesothelial cells were comparably less sensitive with a mean IC₅₀ value of 3.39 nmol/L. Interestingly, we observed a tendency towards trabectedin hypersensitivity in the cisplatin-resistant P31res1.2 as compared with the parental cell line P31 being one of the least responsive MPM cell model in the tested panel (Fig. 1A; Supplementary Table S1). In addition, no significant relation was found between trabectedin and cisplatin response suggesting a different cellular mode-of-action. Although cell models derived from the major histologies of MPM did not significantly differ concerning responsiveness to both drugs, a trend towards enhanced trabectedin activity against nonepithelioid MPM cell cultures was observed (Supplementary Fig. S1C).

To investigate the effect of trabectedin on 3D cell growth and on the stem cell-like MPM cell subpopulation, nonadherent spheroids were generated (Fig. 1C). All four MPM cell lines tested formed a low number (<8) of large spheroids in every control well based on early fusion of multiple small spheroids. In biphasic MPM cell models (SPC111, SPC212), trabectedin widely impaired spheroid formation already at a concentration of 1 nmol/L (Fig. 1C, bottom, and SPC111 as well as SPC212 representatively in Fig. 1D). P31 and P31res1.2 cell spheroids were still formed at 1 nmol/L but with decreased number (Fig. 1C, bottom) and diameter (Fig. 1C, top). Of note, sensitivity was slightly higher in the P31res1.2 subline with acquired cisplatin resistance. Spheroid formation was completely impaired at 5 nmol/L in case of both the biphasic and the epithelioid cell lines. Moreover, the self-renewal capacity of spheroids was blocked completely in the presence of trabectedin.

To investigate apoptosis induction, HOE/PI staining (Fig. 2A and B) and TUNEL assays (Fig. 2C) were performed. Already in phase-contrast images a massive reduction of viable SPC111 cells and a loss of cell–cell contact as a consequence of a 24-hour exposure to 2.5 nmol/L trabectedin was observed. Hoechst 33258 dye staining elucidated frequent chromatin condensation indicative for apoptotic cell death induction. Indeed, a high percentage of cells with condensed chromatin double stained with PI confirmed late-stage apoptotic cell death (Fig. 2A). Total apoptotic count is shown in Fig. 2B indicating significantly enhanced apoptosis rates already at 1 nmol/L trabectedin in all four cell lines tested. Relevant numbers of necrosis were not found. These results were confirmed by TUNEL assay detecting apoptosis-related DNA strand breaks (Fig. 2C). The comparably weaker apoptosis induction in TUNEL assay might be based on the loss of nonadherent late apoptotic cells during washing and fixation steps. To dissect the molecular mechanisms involved in trabectedin-induced apoptosis, Western blot analysis was carried out after 24 hours drug exposure (Fig. 2D). PARP cleavage, slightly detectable already at 1 nmol/L but massively enhanced at 10 nmol/L trabectedin, confirmed the involvement of caspases in the apoptotic process. Furthermore, upregulation primarily of the proapoptotic bcl-2 family members bim and bax was detected at 1 nmol/L trabectedin, while at the massively cytotoxic dose of 10 nmol/L protein levels sharply dropped. With regard to the antiapoptotic family members, trabectedin treatment resulted in reduced expression of bcl-2 in the P31 model and of Mcl-1 in all MPM cell lines tested. P53 was only detectable in SPC111 and SPC212 and was slightly upregulated after trabectedin exposure while P31 and P31res1.2 cells lacked p53 expression.

Cell invasion, measured by migration of cells through Matrigel-sealed Boyden chamber membrane pores, was weakly but significantly inhibited at 0.5 and 1 nmol/L trabectedin treatment in three of the four tested MPM cell lines (Fig. 3A). The observed effects are considered widely independent of cell growth inhibition, as trabectedin at least at 0.5 nmol/L did not induce major cytotoxicity during the 28-hour observation time. In contrast to invasion, cell migration, analyzed by in vitro wound closure assay, was not significantly inhibited up to 1 nmol/L trabectedin with the exception of SPC111 at 8 and 12 hours exposure (Supplementary Fig. S2). Accordingly, also in videomicroscopy rather enhanced than reduced cell movement was observed in trabectedin-treated cells before undergoing cell death (Supplementary Video SV1).

In HOE/PI staining a clear-cut reduction of cells in mitotic phase of the cell cycle was detected already from doses of 1 nmol/L onwards (Fig. 3B). Consequently, we investigated the impact of trabectedin on cell-cycle distribution by PI staining and FACS analysis at low trabectedin concentrations (Fig. 3C). Especially, in the two biphasic cell models an accumulation of cells in the G₂–M and a decrease of the G₀–G1 phase were observed already at a concentration of 0.5 nmol/L. This effect was also visible but less pronounced in the less responsive p31 and p31res1.2 cells. Together with the HOE/PI data, the FACS analyses suggest an accumulation of cells in G₂ phase of the cell cycle. Accordingly, expression of cyclin D1 tended to be reduced after trabectedin treatment at 1 nmol/L, while cyclin A2 (representative for the G₂–M transition) was distinctly increased in line with the accumulation of cells in G₂ phase (Fig. 3D). At the highly cytotoxic trabectedin concentration (10 nmol/L), cyclin E1 levels (G₁–S) were elevated and cyclin A2 upregulation lost.

SCID mice carrying human SPC111 xenografts were treated with a single injection of 0.15 mg/kg trabectedin or solvent intravenously (compare Materials and Methods). Trabectedin was generally well tolerated with no major toxicity during the observation period. Mice were sacrificed 15 days after drug administration and all tumor nodules collected from the peritoneal cavity (representative animals from each group are shown in Fig. 4A). The mean tumor weight was distinctly reduced (by 43%) indicating that even single administration of trabectedin led to a significant reduction of the intraperitoneal tumor burden (Fig. 4B). This activity was comparable with the one observed in case of cisplatin monotherapy for the identical MPM model tested in frame of a previous study (26).

Next, we tested whether trabectedin might be a feasible option for combination approaches with the standard-of-care mesothelioma drugs pemetrexed and cisplatin. While the multitargeted antimetabolic drug pemetrexed showed mainly additive to antagonistic effects with trabectedin, cisplatin exerted primarily synergistic activities. This was already clearly observable in HOE/PI staining with distinctly enhanced apoptotic rates at a combination of 1 nmol/L trabectedin and 5 μmol/L cisplatin (Fig. 5A and B). Predominantly synergistic effects were confirmed by dose–response curves established by viability assays following 72-hour drug exposure in all four MPM cell lines tested (Fig. 5C). Accordingly, combination indices (CI values) for several combination settings were distinctly lower than 0.9 indicating marked synergism in several cases (Supplementary Fig. S3).

To uncover mechanisms underlying the differences in intrinsic trabectedin sensitivity, genome-wide gene expression signatures of the investigated MPM cell lines were established. When comparing MPM cell models categorized according to trabectedin sensitivity into high (h)- and low (l)-responsive subgroups (dichotomized by the median IC₅₀ value, Supplementary Table S1), distinct gene expression differences regarding apoptosis-related genes were observed. Analyses of both the “KEGG_Apoptosis” and the “BIOCARTA_Death Pathway” gene sets depicted the strongest expression differences (4.4-fold up in the low-responsive subgroup) for the anti-apoptotic gene BCL2 (Supplementary Table S2). Accordingly, a significant inverse correlation between bcl-2 mRNA expression and trabectedin response was observed (Supplementary Fig. S4A). Also analysis of the data by Ingenuity software suggested deregulation of a bcl-2 pathway network between the two subgroups (Supplementary Fig. S4B). Accordingly, we decided to test whether pharmacologic inhibition of the antiapoptotic bcl-2 family members by obatoclax (31) would synergize with trabectedin against MPM cells. HOE/PI staining indicated distinctly higher rates of apoptosis in the combination groups, with the most prominent effects seen in the less trabectedin-responsive MPM cell models SPC212 and p31 (Fig. 6A). Likewise, in 72-hour drug exposure MTT assays a clear synergism (CI values < 0.9) between trabectedin (at low nanomolar to picomolar concentrations) and obatoclax was observed in all cell models tested (Fig. 6B and Supplementary Fig. S5A). Furthermore, the combination of trabectedin with obatoclax enhanced cleavage of PARP and caspase-7 as well as upregulated proapoptotic bim in SPC111 and SPC212 cells (Fig. 6C). Comparable observations were made in the P31 cell models with the difference that in this MPM cell model caspase-7 was undetectable at the protein level (Supplementary Fig. S5B). Interestingly, obatoclax alone tended to enhance bcl-2 protein levels. This effect was attenuated or inhibited by trabectedin coexposure especially in the SPC111 and the P31res1.2 cell models (Fig. 6C and Supplementary Fig. S5B).

Regarding temporal resolution, MPM cell death was monitored over 48 hours by videomicroscopy (the 24- to 49-hour exposure time frame is shown in Supplementary Video SV1). Trabectedin as single drug (2.5 nmol/L) led, in addition to occasional apoptotic events, to reduced cell proliferation and enhanced cell volumes well in agreement with the loss of mitosis and the G₂ cell cycle arrest (compare Fig. 3B and C). Obatoclax alone (500 nmol/L) led to a dramatic change in cell shape and gain of a more fibroblastoid and highly migratory phenotype. In the combination group, massive cell death occurred especially after around 30–48 hours of exposure. As obatoclax is a relatively unspecific bcl-2 inhibitor, we tested the combination of trabectedin also with the more selective compound ABT-199 (32). This combination again led to mostly additive to synergistic activities in the MPM cell lines tested (Supplementary Fig. S6).

Chemotherapy still remains the main backbone of systemic MPM therapy. Platinum in combination with pemetrexed has so far proven to be the most effective first-line treatment combination. However, median survival rates remain poor and chemotherapy in this setting can only improve survival by a few months (8). In terms of second- or even third-line treatment, no drugs, either classic cytotoxic or targeted agents, have been effective so far. Therefore, an unmet need for the discovery of more potent agents still exists.

In this preclinical study, we show that the marine-derived alkaloid trabectedin (i) reduces MPM cell growth in a dose-dependent manner in both monolayer and multicellular spheroid cell cultures; (ii) induces apoptosis accompanied by changes in the expression of proapoptotic and antiapoptotic regulators; (iii) is able to weakly but significantly inhibit cell invasion without influencing the migratory potential of MPM cells; (iv) alters MPM cell mitosis rates and leads to perturbation of the cell cycle; (v) strongly reduces tumor growth in vivo in an orthotopic MPM xenograft model in SCID mice; and (vi) acts synergistically with cisplatin and bcl-2 inhibitors in vitro. In addition, trabectedin activity was distinctly higher in case of MPM cell explants as compared to those from nonmalignant pleural tissue. This corresponds well with preliminary published data on selective accumulation of trabectedin in MPM as compared with plasma and nonmalignant tissue (16).

No preclinical data about trabectedin as an anti-mesothelioma strategy have been published so far with the exception of an AACR 2016 meeting abstract indicating activity of trabectedin against patient-derived mesothelioma xenografts (33). In addition, a few mesothelioma cases within two phase I trials were reported. Interestingly, in two of these MPM cases, clinical response (partial responses and stable disease) was seen (34, 35), giving a first hint for clinical activity in this notoriously unresponsive tumor type. Consequently, a phase II trial (trial registration ID: NCT02194231) has been launched recently (36) and preliminary results were presented on ASCO 2015 indicating clinical benefit especially for patients with nonepithelioid histology (37). This is in good accordance with our preclinical data, demonstrating a tendency towards enhanced activity in primary cell cultures derived from patients with biphasic or sarcomatoid MPM. Of note, the standard MPM chemotherapeutic agent cisplatin is known to be more efficient in epithelioid MPM subtypes. This implies that cisplatin and trabectedin might represent a feasible combination therapy strategy for MPM. Indeed in several tumor types such as ovarian cancer, breast cancer and diverse sarcomas (38–41) combination of platinum compounds with trabectedin has proven to be synergistic both in vitro and in vivo. In accordance, phase I studies including both patients with sarcomas and diverse carcinomas have reported that the combination of cisplatin and trabectedin is feasible, well tolerated, and showed antitumor effects in heavily pretreated patients (42, 43). Nevertheless, our attempts to combine trabectedin and cisplatin in vivo resulted, despite indications of distinct anti-MPM activity, in premature experiment termination due to local toxicity at the injection sites and distinct weight loss of mice. Consequently, alternative application routes using osmotic pump–mediated infusion, more closely mimicking the clinical form of trabectedin application via infusion, are currently established.

At the molecular level, the synergism between trabectedin and cisplatin might be caused by the specific modes of action and interaction with repair pathways characteristic for these two DNA-interacting compounds (21). While trabectedin exerts its activity at least in part by a mode-of-action involving nucleotide excision repair (NER), this repair mechanism is well known to protect against cisplatin-induced DNA damage by recognizing and removing DNA platinum adducts. Cancer cells which are deficient in NER exert high sensitivity to cisplatin but show resistance to trabectedin treatment (44, 45). In contrast, mismatch repair mechanisms are contributing to cisplatin- but not trabectedin-mediated cell death while homologous recombination repair (HR) protects against both compounds (21). Accordingly, trabectedin activity was not reduced but in selected cases even enhanced in cisplatin-resistant cancer sublines (45) and vice versa (46). This holds also true for an MPM cell model with acquired cisplatin resistance in our study (p31 and p31res1.2), exhibiting mild hypersensitivity against trabectedin in vitro and overexpression of several NER components in gene expression arrays. This makes sense as MPM represents a very treatment-refractory tumor type rarely responding to platinum-based drugs used in clinical routine. Probably reflecting its proinflammatory origin, DNA repair mechanisms play an integral role in the resistance of MPM to cytotoxic agents (47). With regard to NER, polymorphisms in two genes (ERCC1, XPD) influenced platinum-treatment efficacy and toxicity in a series of 133 mesothelioma patients (48). Together, this indicates that the opposite interaction with NER processes might be central to the synergism between trabectedin and cisplatin also in case of MPM.

To determine major players in regulating trabectedin response in our collection of MPM primary cell cultures, gene signature differences between the more and less responsive MPM subgroup have been compared. Concerning several apoptosis-related signatures, the gene most strongly upregulated in the less trabectedin-sensitive MPM cell cultures was BCL2, an antiapoptotic member of the bcl-2 family of cell death regulators. Consequently, we investigated whether a combination of trabectedin with bcl-2 inhibitors in vitro might be a feasible strategy for treatment of MPM. Indeed, two small-molecule bcl-2–interacting drugs with different specificities (obatoclax and ABT-199; refs. 31, 32) demonstrated synergistic activities in multiple cell biological assays. Western blots confirmed that several members of the bcl-2 family are altered by this novel anticancer drug combination in a synergistic manner. Thus, a clear-cut activation of the proapoptotic bim and in some cases also bax already at low trabectedin concentrations was further enhanced by administration of obatoclax. In contrast, expression of the antiapoptotic bcl-2, bcl-x_L and Mcl-1 family members was distinctly reduced by trabectedin. This implicates a shift towards cell death induction well in accordance with the data derived from diverse cell death assays including PARP and caspase cleavage. Accordingly, it was reported that siRNA-mediated knockdown of bcl-x_L and Mcl-1 was sufficient to induce MPM cell death without the need for an additional chemotherapeutic compound (49). In addition, the anticancer activity of the BH3 α-helix mimetic JY-1-106 was found to be mediated by disruption of bcl-x_L and Mcl-1 protein–protein interactions with Bak in diverse cancer cell lines including MPM (50). Altered expression of bcl-2 family members including bim was also reported as a mechanism underlying apoptosis resistance in MPM spheroids (51). This corresponds well to the destruction of spheroid growth, representing a surrogate marker for the cancer stem cell compartment, of our MPM cell lines already at 1 nmol/L trabectedin. However, this promising in vitro synergism of trabectedin and bcl-2 inhibition needs to be further confirmed in in vivo models.

In summary, we present evidence that trabectedin is active as a single agent against human MPM in vitro and in vivo by directly targeting antiapoptotic mechanisms. Accordingly, this effect can be synergistically enhanced either by chemotherapeutic agents inducing apoptosis (like cisplatin) or by agents targeting antiapoptotic bcl-2 family members (like obatoclax and ABT-199) in vitro. Together with the published early indication of clinical responses (34, 35), our data suggest the development of trabectedin especially within combination approaches for treatment of this devastating disease.

No potential conflicts of interest were disclosed.

Conception and design: M.A. Hoda, M. Grusch, W. Berger

Development of methodology: M.A. Hoda, Y. Dong, K. Schelch, V. Laszlo, J. Ozsvar, M. Grusch, B. Hegedüs

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): M.A. Hoda, C. Pirker, K. Schelch, P. Heffeter, T. Klikovits, V. Laszlo, B. Döme, B. Hegedüs, W. Berger

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): M.A. Hoda, C. Pirker, Y. Dong, K. Schelch, S. van Schoonhoven, T. Klikovits, B. Döme, B. Hegedüs, W. Berger

Writing, review, and/or revision of the manuscript: M.A. Hoda, C. Pirker, Y. Dong, K. Schelch, P. Heffeter, K. Kryeziu, T. Klikovits, B. Döme, M. Grusch, B. Hegedüs, W. Berger

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): K. Kryeziu, S. van Schoonhoven, T. Klikovits, A. Rozsas, M. Grusch

Study supervision: W. Klepetko, B. Hegedüs, W. Berger

We thank Mirjana Stojanovic, Barbara Dekan, and Gerhard Zeitler for their assistance with the in vitro and in vivo experiments.

This research was supported by the Funds of the Oesterreichische Nationalbank, Anniversary Fund (No. 14574; to M.A. Hoda) and by the Initiative Cancer Research of the Medical University Vienna (to W. Berger).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.