Human multiple myeloma cell lines were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum, 2 mmol/l L-glutamine, 5 U/ml penicillin, and 5 μg/ml streptomycin. PCNY-1, MM1.S, MM1.R, KMS-11 and ARP-1 cells were kindly provided by Hearn Cho (New York University School of Medicine, New York, NY, USA). All cells were maintained in an incubator with a humidified atmosphere of 5% CO₂ at 37°C. Sulforaphane, erysolin, erucin, arsenic trioxide and N-acetyl-l-cysteine (NAC) were purchased from Sigma (St. Louis, MO, USA). Bortezomib was acquired from LC Laboratories (Woburn, MA, USA).

For dose response assays, 5,000 cells per well were plated in 96-well culture plates. After overnight incubation, the cells were treated with indicated compounds. Following a 72-h incubation period, cellular proliferation was assessed using a tetrazolium dye reduction assay (CellTiter 96 Aqueous Non-Radioactive Cell Proliferation assay; Promega, Madison, WI, USA) carried out according to the manufacturer’s instructions as previously described (24). Absorbance was recorded on a microplate reader at 495 nm. Cellular proliferation was expressed as a percentage of vehicle-treated cells which were defined as 100% viable. Drug interaction was analyzed by Calcusyn software (Biosoft, Cambridge, UK). This software determines the interaction of two drugs through calculations of the combination index (25) based upon the multiple drug effect equation of Chou and Talalay (26,27). Denotation of CI values as follows: >1, antagonism; 1, additivity; <1, synergy. GI₅₀ values, which represent the concentration necessary to cause 50% growth inhibition, were calculated from dose-response curves generated from cell proliferation assays.

Cellular levels of glutathione (GSH) levels were determined using the HT Glutathione Assay kit from Trevigen (Gaithersburg, MD, USA) as previously described (22). Briefly, 24 h after seeding MM cells at a density of 1×10⁶, cells were exposed to indicated compounds for 3 h at 37°C. The cells were collected and washed with PBS. An equal number of cells from each treatment group were aliquoted for use in cellular GSH quantification per the manufacturer’s instructions.

Cells were harvested in extraction buffer [1% Triton X-100, 50 mmol/l Tris, 2 mmol/l EDTA, 150 mmol/l NaCl (pH 7.5)] containing protease inhibitor cocktail (Roche Applied Science, Indianapolis, IN, USA) and phosphatase inhibitor cocktail (Sigma) after two washes with ice-cold phosphate buffered saline (PBS). The lysates were centrifuged at 10,000 × g at 4°C for 10 min in a microcentrifuge. Protein supernatants were measured with a protein assay kit (Bio-Rad, Hercules, CA, USA). Proteins were separated by 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto polyvinylidene difluoride membranes (Polyscreen; Perkin-Elmer, Waltham, MA, USA). Antibodies against cleaved forms of poly(ADP-ribose) polymerase (PARP), caspase-3 and -4 were obtained from Cell Signaling Technology (Danvers, MA, USA). Phospho-PERK, phospho-eIF2α, CHOP and HSP90 antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-actin (Sigma) was used as a control. Immunoreactive bands were visualized using enhanced chemiluminescence detection reagent (Perkin-Elmer) and X-OMAT processing.

Cells were treated with indicated concentrations of ATO and/or sulforaphane for 24 h. Total RNA was isolated from lysed cells with the RNeasy Mini kit (Qiagen, Germantown, MD, USA). XBP1 splicing was assessed by semi-quantitative RT-PCR as described previously (28,29). cDNA was produced from total RNA preps using ImProm-II Reverse Transcription system (Promega). Primers spanning the fragment of XBP1 containing the intron targeted by Ire1α were used: 5′-TACGGGAGAAAACTCACGGC-3′ and 5′-GGGTCCAACTTGTCCAGAATGC-3′. The thermal PCR cycling conditions are as follows: 95°C for 5 min, 95°C for 1 min, 58°C for 30 sec, 72°C for 30 sec, and 72°C for 5 min with 35 cycles of amplification. PCR products were digested with Pst1 (New England Biolabs, Ipswich, MA, USA) before being separated on a 2.0% agarose/1X TAE gel and visualized by ethidium bromide.

Commercially available lentiviral particles obtained from GenTarget (San Diego, CA, USA) expressing Gaussia luciferase (Gluc) were introduced into KMS-11 and ARP-1 MM cells by infection. For infection, cells were cultured in 6-well tissue culture plates to a density of 3×10⁶ cells/ml and then diluted to 1×10⁶ cells/ml in complete media. Lentiviral particle was added at a ratio of 100 μl viral particles to 1 ml of cells. After 24 h, equal parts of fresh media containing puromycin selection were added. After 72 h, efficacy of transduction was assessed by RFP fluorescence. After infection, cells were maintained in media containing 3 μg/ml puromycin to establish stable clones. For secretion assays, 100,000 cells/well were plated in 96-well culture plates. Cells were immediately treated with indicated concentrations of compounds. Following 24-h incubation, expression and secretion of Gluc was monitored using The BioLux Gaussia Luciferase Assay kit (New England Biolabs) according to the manufacturer’s instructions through measurements of luciferase activity as indicated by relative light units (RLU) on a microplate luminometer (Molecular Devices, Sunnyvale, CA, USA). Percent reduction in Gluc secretion was determined by the following equation: (RLU of treated cells/RLU ratio of the untreated, DMSO control cells) × 100.

Unless otherwise noted, experiments were performed in triplicate. The data are presented as the average ± SEM. P-values were determined by a two-sided Student’s t-test with unequal variance, with P<0.05 considered significant.

In order to enhance the potential clinical efficacy of ATO in the treatment of MM, we wanted to assess its interactive potential with BTZ, a current standard of care in the treatment of MM. As shown in Fig. 1, low, subclinical doses of BTZ and ATO as single agents have limited effect on cellular proliferation in MM. However, when used in combination, BTZ and ATO markedly inhibit cellular proliferation in both KMS-11 and ARP-1 cells (Fig. 1).

Given that recent studies indicate that isothiocyanates possess anti-proteasomal activity, we hypothesized that isothiocyanates should have biological effects similar to BTZ. In order to test this hypothesis, we examined the effect of isothiocyanates on TNFα induced proteasomal degradation of Iκβ. Upstream activation signaling by TNFα-stimulation causes phosphorylation of Iκβ which is ultimately targeted for degradation by the 26S proteasome (30). As previously reported (31) and shown in Fig. 2, proteasome inhibitors such as BTZ prevent TNFα-induced Iκβ proteasomal degradation. Similarly, sulforaphane and erysolin pretreatment also prevent Iκβ degradation after TNFα stimulation. Erucin pretreatment only minimally inhibits Iκβ degradation.

Because these data along with other published studies suggest that isothiocyanates inhibit proteasomal mediated protein degradation, we wanted to investigate their ability to potentiate the growth inhibitory effects of ATO. As shown in Tables I and II, sulforaphane and erysolin display a greater than additive effect (i.e., synergize) with ATO in 3 MM cell lines. Consistent with its inability to inhibit TNFα induced degradation of Iκβ (Fig. 2), erucin interaction with ATO was found to be antagonistic (Table II). Due to its potency, we focused further experiments on sulforaphane.

As shown in Fig. 3, ATO caused modest growth inhibition in PCNY-1 MM cells when employed as a single agent. Similarly, concentrations up to 5 μM sulforaphane had a minimal effect on the proliferation of MM cells (Fig. 3). However, when used in combination, the ability of these compounds to reduce proliferative capacity was dramatically enhanced. Combination index analysis demonstrates the relationship between 0.5 μM ATO and 3 μM sulforaphane to be strongly synergistic (Table I). Similar effects were observed in 4 out of 5 MM cell lines examined (Fig. 3 and Table I). The synergistic relationship was not observed in MM1.R cells which are a subclone of the MM.1 human MM cell line selected for resistance to glucocorticoid therapy (32).

As we previously demonstrated in leukemic cells (22), sulforaphane also acts as an ATO sensitizing agent through depletion of intracellular GSH in MM cells (Fig. 4A). In this capacity, cellular depletion of glutathione by sulforaphane renders cells largely incapable of protection against ATO’s ability to generate ROS. In order to understand the biological consequences of enhanced ROS, we evaluated the induction of apoptosis via cleavage of effector caspase-3 and poly(ADP-ribose) polymerase (PARP), indicators of apoptosis. As shown in Fig. 4B, higher levels of cleaved caspase-3 and PARP protein were present in KMS-11 cells exposed to the combination treatment of 3 μM sulforaphane and 1 μM ATO than with either agent alone. Similar results were observed in ARP-1 cells (data not shown). These data indicate that combinatorial sulforaphane/ATO treatment enhances the apoptotic response. Interestingly, cleavage of the ER stress specific caspase-4 is also enhanced in response to combination treatment (Fig. 4B).

ROS play a critical role in the apoptotic response of combined ATO and sulforaphane treatment as demonstrated by the fact that preincubation with the ROS scavenger N-acetyl cysteine (NAC) partially attenuated the apoptotic induction of combination treatment (Fig. 4B). Pretreatment with NAC had no effect on apoptosis alone. However, in combination with ATO and sulforaphane, NAC partially inhibited PARP, caspase-3 and -4 cleavage to levels comparable to treatment with ATO alone.

Hypothesizing that sulforaphane’s anti-proteasomal activity also is integral to its ability to augment ATO cytotoxicity in MM cells, we investigated whether combination treatment resulted in enhanced ER stress due to perturbations in protein processing. Consistent with this notion, we observed upregulation of HSP90, a general marker for ER stress (33), in KMS-11 cells co-treated with ATO and sulforaphane (Fig. 5A). Additionally, activation of the PERK pathway, a key component of the unfolded protein response (UPR), was enhanced upon co-treatment. As shown in Fig. 5A, PERK phosphorylation was elevated after treatment with ATO and sulforaphane with pathway activation demonstrated by increased expression of downstream mediators CHOP as well as phosphorylation of eIF2. Consistent with upregulation of the PERK arm of the UPR response, parallel activation of the IRE1 arm of the UPR response was observed through enhanced splicing of the UPR transcription factor XBP1 upon treatment with ATO and sulforaphane (Fig. 5B). Similar results were observed in ARP-1 MM cells (data not shown).

B cells synthesize and secrete immunoglobulin protein. Blocking or decreasing protein processing in the secretory pathway is another hallmark of ER stress that is specifically relevant to the biology of MM cells (34). In order to assess the effects of ATO and sulforaphane on protein secretion, we employed the reporter protein Gaussia luciferase (Gluc). Gluc is a naturally secreted luciferase that can be easily monitored through extracellular release of luciferase activity in real time, and has been developed as a sensor of ER stress (35). As shown in Fig. 6, clones of KMS-11 and ARP-1 stably expressing Gluc show a notable reduction in Gluc secretion when treated with 1 μM ATO or 3 μM sulforaphane alone. Consistent with our studies of additional ER stress markers, combination ATO and sulforaphane treatment inhibits protein secretion in a fashion that is greater than treatment with single agent alone. Altogether, these data suggest that when ATO and sulforaphane are administered together, ER stress mediated pathways are enhanced.