Purpose: The transcription factor specificity protein 1 (Sp1) controls number of cellular processes by regulating the expression of critical cell cycle, differentiation, and apoptosis-related genes containing proximal GC/GT-rich promoter elements. We here provide experimental and clinical evidence that Sp1 plays an important regulatory role in multiple myeloma (MM) cell growth and survival.

Experimental Design: We have investigated the functional Sp1 activity in MM cells using a plasmid with Firefly luciferase reporter gene driven by Sp1-responsive promoter. We have also used both siRNA- and short hairpin RNA–mediated Sp1 knockdown to investigate the growth and survival effects of Sp1 on MM cells and further investigated the anti-MM activity of terameprocol (TMP), a small molecule that specifically competes with Sp1-DNA binding in vitro and in vivo.

Results: We have confirmed high Sp1 activity in MM cells that is further induced by adhesion to bone marrow stromal cells (BMSC). Sp1 knockdown decreases MM cell proliferation and induces apoptosis. Sp1-DNA binding inhibition by TMP inhibits MM cell growth both in vitro and in vivo, inducing caspase-9–dependent apoptosis and overcoming the protective effects of BMSCs.

Conclusions: Our results show Sp1 as an important transcription factor in myeloma that can be therapeutically targeted for clinical application by TMP. Clin Cancer Res; 17(20); 6500–9. ©2011 AACR.

Translational Relevance

The transcription factor specificity protein 1 (Sp1) affects growth and metastatic potential of tumor cells. In MM, key genes such as NF-κB p65, IGF-IR, VEGF, and IL-6 contain proximal GC-rich promoter sequences, and their interactions with Sp proteins are critical for their expression. Here, we show that Sp1 plays an important role in myeloma cell growth and survival. Importantly, Sp1-specific inhibitor can induce tumor apoptosis in murine models of MM. Thus, inhibition of Sp1 may be an attractive therapeutic modality in MM, alone or in combination with other agents.

Specificity protein 1 (Sp1) and other Sp and Kruppel-like factor proteins are members of a family of transcription factors that bind GC/GT-rich promoter elements through 3 C2H2-type zinc fingers that are present at their C-terminal domains (1).

Sp1 regulates gene expression both by direct interaction with promoter elements and via protein–protein interactions or interplay with other transcription factors, such as Ets-1, c-myc, c-Jun, Stat1, and Egr-1, and/or components of the basal transcriptional machinery (2). Sp1 has also been linked to chromatin remodeling through interactions with chromatin-modifying factors such as p300 (3) and histone deacetylases (HDAC; ref. 4).

Although Sp1 has been considered as a ubiquitous transcription factor, increasing evidence suggests that it plays a major role in regulating expression of cell differentiation, cell cycle, and apoptosis-related genes affecting cellular growth (5). Sp1 levels and/or activity are increased in multiple cancers including breast (6), colon (7), gastric (8), pancreatic (9–12), and thyroid cancer (13) as compared with normal tissues. Elevated Sp1 expression is inversely correlated with the survival of patients with gastric cancer (14) and identifies advanced stage tumors and predicts a poor clinical outcome in primary pancreatic adenocarcinoma (12). On the other hand, interference of Sp1 activity has been shown to suppress tumor cell growth (15, 16) as well as tumor formation in athymic mice (6, 17), suggesting that Sp1 plays a central regulatory role in controlling the number of pathways of tumor development and progression and thus may be an attractive therapeutic target.

Sp1 could contribute to transformation via regulation of the expression of Sp1-responsive genes including those supporting cell growth (c-jun, Raf, cyclins, E2F1, TGF-β, IEX-1, and TCL1; refs. 18, 19) and apoptosis (Bcl-2, survivin; refs. 5, 20). Sp1 also regulates genes involved in angiogenesis and metastasis, including VEGF, uPA (21), PSA, and MT1-MMP (22). Enhanced Sp1 activity is related to both increased Sp1 gene expression and its posttranslational modification; for example, Sp1 phosphorylation regulates target genes in both positive and negative directions (23, 24).

Although Sp1 has been described to modulate autocrine interleukin (IL)-6 secretion by multiple myeloma (MM) cells affecting its growth (25), its role in MM pathobiology remains unexplored. The promoter of several genes such as NF-κB p65, IGF-IR, VEGF, hTERT, and IL-6 that regulate MM cell growth, cell-cycle progression, survival, and apoptosis contains proximal GC-rich promoter sequences that interact with Sp1 protein for their optimal expression (25–28). In addition, the same kinase pathways that have been shown to increase Sp1 phosphorylation and the transactivation of target genes (21, 22) are also known to mediate proliferation, survival, drug resistance, and migration (ERK, Jak/STAT, PI3K/Akt, and PKC signaling cascades, respectively) in MM.

Here, we report significant role of Sp1 in myeloma cell growth and survival. Our results suggest that Sp1 activity can be therapeutically targeted for clinical application in MM.

Cells

Bone marrow mononuclear cells and primary MM cells from bone marrow aspirates from MM patients following informed consent and Dana-Farber Cancer Institute Institutional Review Board (IRB) approval were isolated by Ficoll–Hypaque density gradient sedimentation. MM patient cells were separated from bone marrow samples by antibody-mediated positive selection using anti-CD138 magnetic-activated cell separation microbeads (Miltenyi Biotech). Bone marrow stromal cells (BMSC) were established as previously described (29). MM cell lines were cultured in RPMI 1640 (Mediatech) supplemented with 10% FBS. The IL-6–dependent MM cell lines INA-6 (kindly provided from Dr R. Burger, University of Kiel, Kiel, Germany) were cultured in the presence of 2.5 ng/mL rhIL-6 (R&D Systems).

Reagents

Terameprocol (TMP), kindly provided by Erimos Pharmaceuticals, was synthesized and dissolved in 30 hydroxypropyl CPE B-cyclodextrin and 25 polyethylene glycol 300 as previously described (30).

Cell proliferation assay

MM cell proliferation was measured by [3H]thymidine (Perkin-Elmer) incorporation assay as previously described (29).

Bromodeoxyuridine staining

The proportion of myeloma cells in S-phase was determined by incorporation of bromodeoxyuridine (BrdUrd). A total of 1 × 106 MM cells were exposed to 10 μg/mL of BrdUrd for 30 minutes. The cells were then harvested and stained with FITC anti-BrdUrd antibody and 7-aminoactinomycin D (7-AAD) using a BrdU Flow kit (BD Bioscience Pharmingen) according to the manufacturer's directions. Cells were analyzed by flow cytometric analysis with a Becton-Dickinson FACScan flow cytometer.

Apoptosis assay

Apoptosis was evaluated by flow cytometric analysis following Annexin-V and propidium iodide (PI) staining.

Sp1 binding activity

The Sp1 binding activity was analyzed using the Transcription Factor ELISA Kit, a DNA-binding ELISA-based assay (Panomics). Sp1 transcription factor binding to its consensus sequence on the plate-bound oligonucleotide was studied from nuclear extracts, following the manufacturer's procedure. Briefly, nuclear proteins were extracted with a Nuclear Extraction kit (Panomics) and quantified using the Bio-Rad Protein Assay Kit (Bio-Rad). A total of 15 μg of nuclear protein from each treatment was analyzed. Sp1 antibody was used as the primary antibody, and anti-rabbit IgG horseradish peroxidase was used as the secondary antibody. The absorbance was measured at a wavelength of 450 nm on a spectrophotometer.

Promoter activity assay

The Sp1 promoter reporter constructs were purchased from Sabiosciences. To examine transcriptional regulation of the Sp1 promoter by TMP, MM cells were transiently transfected with 1 μg of Sp1 reporter plasmid or empty vector control by electroporation using AMAXA technology according to the manufacturer's instructions. Luciferase assays were done with a Luminoskan Ascent 2.4 luminometer and the Dual-Luciferase Reporter Assay System (Promega). Firefly luciferase values were normalized to Renilla luciferase activity and were either expressed as relative luciferase units (RLU) or as mean “fold induction.”

Sp1 knockdown

siRNA.

RNA interference was done using the TranSilent Human Sp1 siRNA (Panomics, Inc.) following the manufacturer's instructions. Nontargeting scrambled negative control siRNA (Panomics, Inc.) was used as a negative control. Briefly, U266 and OPM2 cells were seeded to 80% confluency in 6-well plates in triplicate and transiently transfected with 2 μmol/L of Sp1 siRNA by electroporation using AMAXA technology.

Short hairpin RNA.

Lentiviral short hairpin RNAs (shRNA) were used to knockdown Sp1 expression in MM cells. Scrambled and Sp1 pLKO shRNA vectors were provided by Dr. William Hahn (Dana-Farber Cancer Institute). Recombinant lentivirus was produced by transient infection of 293T cells following a standard protocol, as previously described (31).

Immunoblotting

Whole-cell lysates, nuclear extracts, or cytsolic fractions of lysates (30 μg) were subjected to SDS-PAGE using “Precast Gel” (Bio-Rad Laboratories), transferred to a nitrocellulose membrane (Bio-Rad), and immunoblotted with anti-Sp1 (Abcam) anti-survivin, anti-caspase-3, -7, -8, -9, anti-PARP (Cell Signaling Technology) antibodies. After incubating with secondary antibodies, membranes were developed by enhanced chemiluminescence (GE Healthcare).

In vivo study

Subcutaneous model.

The in vivo efficacy of TMP was tested in a murine xenograft model of MM using U266, MM1S, or OPM2 MM cell lines injected subcutaneously in severe combined immunodeficient (SCID) mice. Following detection of tumor, mice were treated with either vehicle or TMP (50 mg/kg) subcutaneously for 5 consecutive d/wk for 2 weeks. Tumor growth was measured as previously described (32). Excised tumors from mice were immediately fixed and stored in 10% formalin. The fixed tissue was then dehydrated through a series of graded alcohols and xylene and embedded in paraffin. The paraffin tissue blocks were thin sectioned and stained for microscopy with hematoxylin–eosin (H&E) or analyzed by immunocytochemical methods for Ki67, terminal deoxynucleotidyl transferase–mediated dUTP nick end labeling (TUNEL), and caspase-3. The survivin level analysis was conducted following staining with a combination of anti-CD138 and anti-survivin antibody. Appropriate immunofluorescent secondary antibodies were applied along with the nuclear dye 4′,6-diamidino-2-phenylindole. The tissue sections were imaged and the relative amount of survivin was localized within tumor-cell nuclear compartment was determined using an automatated quantitation of antigen expression (AQUA) analysis. The data reflects the results of 12 fields imaged at 200× magnification per tumor.

SCID-hu model.

Six- to 8-week-old male CB-17 SCID mice (Taconic) were housed and monitored in our Animal Research Facility. All experimental procedures and protocols had been approved by the Institutional Animal Care and Use Committee (VA Boston Healthcare System). Human fetal bone grafts were subcutaneously implanted into SCID mice (SCID-hu) as previously described. Four weeks following bone implantation, 3 × 106 INA-6 MM cells were injected directly into the human bone implant. Mouse sera were serially monitored for shuIL-6R by ELISA (R&D Systems, Inc.).

Statistical analysis

The statistical significance of differences was analyzed using the Student t test; differences were considered significant when P ≤ 0.05. Tumor growth inhibition and Kaplan–Meier survival analysis were determined using the GraphPad analysis software.

High Sp1 protein expression and activity in MM

Sp1 is ubiquitously expressed in cells; however, its nuclear localization is important for functional activity. We first evaluated nuclear Sp1 protein levels in MM cell lines and normal peripheral blood mononuclear cells (PBMC). All MM cell lines had high nuclear Sp1 expression, whereas normal PBMCs had predominantly cytoplasmic Sp1 with relatively small amount of nuclear localization (Fig. 1A and B). We further confirmed increased Sp1 nuclear levels and binding activity in MM cells compared with PBMCs and BMSCs using an ELISA-based Sp1 binding assay (Fig. 1C).

Figure 1.

High Sp1 protein expression and activity in MM cells. A, nuclear extracts from 8 MM cell lines were subjected to Western blot analysis using anti-Sp1 and p84 antibodies. B, nuclear and cytoplasmic extracts from MM1S, U266, and PBMCs from 3 healthy donors were subjected to Western blot analysis using anti-Sp1 and GAPDH or p84 antibodies. C, 15 μg of nuclear proteins was analyzed for Sp1 activity using the Sp1 TF ELISA kit that measures Sp1 DNA binding activity. Absorbance was obtained with a spectrophotometer at 450 nm and presented as optical density (OD). D, MM1S and U266 cells were transiently transfected with either negative control (NC)-Luc or Sp1 promoter–driven Luc. After 48 hours from transfection, MM cells were treated with 10 μmol/L of U0126 or LY29004 or control for 30 minutes, washed, and then cultured in absence (−) or presence (+) of BMSCs for an additional 6 hours. The firefly luciferase activity was measured in cell lysate and normalized according to Renilla luciferase activity and expressed as RLUs to reflect the Sp1 promoter activity in the absence or presence of BMSCs. The graph shows 1 of 2 representative experiments carried out in triplicate. Results are shown as mean ± SD.

Figure 1.

High Sp1 protein expression and activity in MM cells. A, nuclear extracts from 8 MM cell lines were subjected to Western blot analysis using anti-Sp1 and p84 antibodies. B, nuclear and cytoplasmic extracts from MM1S, U266, and PBMCs from 3 healthy donors were subjected to Western blot analysis using anti-Sp1 and GAPDH or p84 antibodies. C, 15 μg of nuclear proteins was analyzed for Sp1 activity using the Sp1 TF ELISA kit that measures Sp1 DNA binding activity. Absorbance was obtained with a spectrophotometer at 450 nm and presented as optical density (OD). D, MM1S and U266 cells were transiently transfected with either negative control (NC)-Luc or Sp1 promoter–driven Luc. After 48 hours from transfection, MM cells were treated with 10 μmol/L of U0126 or LY29004 or control for 30 minutes, washed, and then cultured in absence (−) or presence (+) of BMSCs for an additional 6 hours. The firefly luciferase activity was measured in cell lysate and normalized according to Renilla luciferase activity and expressed as RLUs to reflect the Sp1 promoter activity in the absence or presence of BMSCs. The graph shows 1 of 2 representative experiments carried out in triplicate. Results are shown as mean ± SD.

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As activation of Akt and ERK signaling pathways induces nuclear translocation and activation of Sp1 (21) and these pathways are activated in MM cells by their interaction with bone marrow microenvironment, we investigated whether the presence of BMSCs could modulate Sp1 activity in MM cells. Using a plasmid containing Firefly luciferase reporter gene driven by Sp1-responsive promoter, we observed that the interaction between MM cells and BMSCs significantly induced the transcriptional activity of Sp1 in MM cells (Fig. 1D). This increase was completely abrogated by the ERK pathway inhibitor U0126 but not by the Akt inhibitor LY29004.

Sp1 knockdown decreases MM cell proliferation

We have further evaluated the role of Sp1 in MM by analyzing the effect of Sp1 knockdown on MM cell growth and survival. We first knocked down endogenous Sp1 by RNA interference in U266 MM cells. Western blot and ELISA-based analysis confirmed reduction in both cytoplasmic and nuclear levels of Sp1 protein following transient transfection of MM cells with Sp1 siRNA compared with cells transfected with control scrambled siRNA (Fig. 2A and B). Interestingly, we also found that reduction of Sp1 levels was associated with inhibition of MM cell proliferation (Fig. 2C) and significant changes in the cell cycle with increase in G2–M and decrease in S-phases (Fig. 2D). We have confirmed these data using OPM2 cells (Fig. 2E). In addition, using 5 different Sp1-specific shRNA constructs, we have confirmed the inhibitory effect of Sp1 knockdown on MM1S cell proliferation and Sp1 activity (Fig. 2F). The cell populations with the largest reduction in Sp1 protein (shRNA #2, #3, #5) showed the greatest cell growth inhibition compared with the vector control cell lines. Two of the Sp1 shRNAs that do not efficiently knockdown Sp1 have less or no effect on phenotype, indirectly but reliably, confirming the specificity of the effect. We used these 3 constructs to confirm shRNA-related results with U266 cell line (Fig. 2G). Together, these results show the role of Sp1 in MM cell growth and survival.

Figure 2.

Sp1 knockdown decreases MM cell proliferation. A, U266 cells were transfected with TranSilent Human Sp1 siRNA. Cell lysates were obtained at indicated time and subjected to Western blot analysis to assess decrease in the Sp1 protein expression posttransfection using anti-Sp1 and GAPDH antibodies. B, the effect of Sp1 knockdown on Sp1 binding activity in MM cells transfected with Sp1 or control siRNA was assessed by Sp1 TF ELISA and presented as proportional change from control cells. C, the effect of Sp1 knockdown on cell proliferation in MM cells transfected with Sp1 or control siRNA was assessed by [3H]thymidine uptake and presented as the percentage of control cells. Data represent mean ± SD of 3 independent experiments carried out in triplicate. D, 48 hours posttransfection with Sp1 or control siRNA, the measurement of cell-incorporated BrdUrd (with FITC anti-BrdUrd) and total DNA content (with 7-AAD) in U266 cells allowed for the discrimination of cell subsets that resided in G0–G1, S, or G2–M phases of the cell cycle. E, OPM2 cells were transfected with TranSilent Human Sp1 siRNA. Cell lysates were obtained at indicated time and subjected to Western blot analysis (top), and cell proliferation was assessed by [3H]thymidine uptake at the indicated posttransfection time. F, MM1S cells were infected with either scrambled (Scr) or 5 different Sp1 shRNAs (sh #1, #2, #3, #4, and #5). Cell lysates were subjected to Western blotting with anti-Sp1 and GAPDH antibodies (top). The transfected cells were analyzed for Sp1 binding activity (line) and cell growth (columns) 24 hours after the second transfection by Sp1 TF ELISA and [3H]thymidine uptake, respectively. The results are presented as change from cells infected with scramble shRNA (bottom). G, U266 cells were infected with either scrambled (Scr) or 3 different Sp1 shRNAs (sh #2, #3, and #5). Cell lysates were subjected to Western blotting with anti-Sp1 and GAPDH antibodies (top), and cell proliferation was assessed 24 hours after the second transfection by [3H]thymidine uptake. The results are presented as change from cells infected with scramble shRNA (bottom).

Figure 2.

Sp1 knockdown decreases MM cell proliferation. A, U266 cells were transfected with TranSilent Human Sp1 siRNA. Cell lysates were obtained at indicated time and subjected to Western blot analysis to assess decrease in the Sp1 protein expression posttransfection using anti-Sp1 and GAPDH antibodies. B, the effect of Sp1 knockdown on Sp1 binding activity in MM cells transfected with Sp1 or control siRNA was assessed by Sp1 TF ELISA and presented as proportional change from control cells. C, the effect of Sp1 knockdown on cell proliferation in MM cells transfected with Sp1 or control siRNA was assessed by [3H]thymidine uptake and presented as the percentage of control cells. Data represent mean ± SD of 3 independent experiments carried out in triplicate. D, 48 hours posttransfection with Sp1 or control siRNA, the measurement of cell-incorporated BrdUrd (with FITC anti-BrdUrd) and total DNA content (with 7-AAD) in U266 cells allowed for the discrimination of cell subsets that resided in G0–G1, S, or G2–M phases of the cell cycle. E, OPM2 cells were transfected with TranSilent Human Sp1 siRNA. Cell lysates were obtained at indicated time and subjected to Western blot analysis (top), and cell proliferation was assessed by [3H]thymidine uptake at the indicated posttransfection time. F, MM1S cells were infected with either scrambled (Scr) or 5 different Sp1 shRNAs (sh #1, #2, #3, #4, and #5). Cell lysates were subjected to Western blotting with anti-Sp1 and GAPDH antibodies (top). The transfected cells were analyzed for Sp1 binding activity (line) and cell growth (columns) 24 hours after the second transfection by Sp1 TF ELISA and [3H]thymidine uptake, respectively. The results are presented as change from cells infected with scramble shRNA (bottom). G, U266 cells were infected with either scrambled (Scr) or 3 different Sp1 shRNAs (sh #2, #3, and #5). Cell lysates were subjected to Western blotting with anti-Sp1 and GAPDH antibodies (top), and cell proliferation was assessed 24 hours after the second transfection by [3H]thymidine uptake. The results are presented as change from cells infected with scramble shRNA (bottom).

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Inhibition of Sp1 activity by a chemical inhibitor induces MM cell growth arrest and apoptosis via caspase activation and reduction in survivin protein level

Our next approach was based on selective interference of Sp1-mediated transactivation of genes with TMP, a lignan tetra-O-methyl nordihydroguaiaretic acid derivative, which has been shown to specifically bind to Sp1-specific DNA binding domains within the responsive gene promoter regions and interfere with the transcription of these Sp1-controlled genes (33–36). We have confirmed decreased DNA-binding activity of Sp1 by TMP as assessed by ELISA-based Sp1 binding assay (Fig. 3A), along with decreased basal and BMSC-induced Sp1 transcriptional activity (Fig. 3B). Importantly, Sp1 protein level in MM cells was not affected by 24-hour TMP treatment (Fig. 3C), suggesting that the modulation of Sp1 binding activity by TMP was not due to change of Sp1 protein level. We have confirmed these data using an additional MM cell line, observing that longer exposure to TMP led to decrease Sp1 protein expression (Supplementary Fig. S1).

Figure 3.

Inhibition of Sp1 binding and transcriptional activity correlates with MM cell growth arrest. A, MM1S and U266 cells were cultured in the presence of 20 μmol/L TMP for 24 hours and nuclear extracts were subjected to Sp1 ELISA assay to assess Sp1 binding to its consensus sequence on the plate-bound oligonucleotide. B, MM cells were transiently transfected with Sp1-Luc plasmid, and 48 hours posttransfection, MM cells were cultured in absence (−) or presence (+) of BMSCs and treated with placebo (C) or 20 μmol/L of TMP (T) for an additional 6 hours. Luciferase activity was measured. Results are reported as mean of fold change from control (untreated cells). Mean values were calculated from 5 independent experiments and are shown as mean ± SD. C, U266 cells were treated with placebo or different concentrations of TMP (1–20 μmol/L) for 24 hours. Nuclear extracts were subjected to Western blot analysis using anti-Sp1 and p84 antibodies to assess Sp1 protein levels. The ratio of Sp1 to p84 for each sample as assessed by densitometric quantitation of band intensity from the Western blot is denoted. D, several MM cell lines were treated with various concentrations of TMP (1–20 μmol/L) for 24 hours and MM cell growth was assessed by [3H]thymidine uptake. Data are presented as the percentage of vehicle-treated cell proliferation. E, primary CD138+ MM cells were cultured in the absence (−) or presence (+) of BMSCs at different concentrations of TMP for 24 hours. Cell proliferation was assessed by [3H]thymidine uptake and expressed as counts per minute (cpm). F, BMSCs from MM patients were treated with different concentrations of TMP for 48 hours, and cell proliferation was assessed by [3H]thymidine uptake and expressed as counts per minute.

Figure 3.

Inhibition of Sp1 binding and transcriptional activity correlates with MM cell growth arrest. A, MM1S and U266 cells were cultured in the presence of 20 μmol/L TMP for 24 hours and nuclear extracts were subjected to Sp1 ELISA assay to assess Sp1 binding to its consensus sequence on the plate-bound oligonucleotide. B, MM cells were transiently transfected with Sp1-Luc plasmid, and 48 hours posttransfection, MM cells were cultured in absence (−) or presence (+) of BMSCs and treated with placebo (C) or 20 μmol/L of TMP (T) for an additional 6 hours. Luciferase activity was measured. Results are reported as mean of fold change from control (untreated cells). Mean values were calculated from 5 independent experiments and are shown as mean ± SD. C, U266 cells were treated with placebo or different concentrations of TMP (1–20 μmol/L) for 24 hours. Nuclear extracts were subjected to Western blot analysis using anti-Sp1 and p84 antibodies to assess Sp1 protein levels. The ratio of Sp1 to p84 for each sample as assessed by densitometric quantitation of band intensity from the Western blot is denoted. D, several MM cell lines were treated with various concentrations of TMP (1–20 μmol/L) for 24 hours and MM cell growth was assessed by [3H]thymidine uptake. Data are presented as the percentage of vehicle-treated cell proliferation. E, primary CD138+ MM cells were cultured in the absence (−) or presence (+) of BMSCs at different concentrations of TMP for 24 hours. Cell proliferation was assessed by [3H]thymidine uptake and expressed as counts per minute (cpm). F, BMSCs from MM patients were treated with different concentrations of TMP for 48 hours, and cell proliferation was assessed by [3H]thymidine uptake and expressed as counts per minute.

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We next assessed the effect of the TMP-mediated inhibition of Sp1 binding activity on MM cell growth. We have examined the effect of the Sp1 inhibitor in MM cell lines with constitutive activation of the canonical and/or noncanonical NF-κB pathways. TMP significantly inhibited DNA synthesis in all MM cells lines tested in a dose-dependent fashion (Fig 3D). In the most sensitive cell line, the IC50 value is in the range of 1 to 10 μmol/L whereas in the less sensitive cell lines, the IC50 value is in the range of 10 to 20 μmol/L for a 24-hour period of treatment. Importantly, TMP inhibited proliferation of primary patient MM cells overcoming the growth-promoting effect of BMSCs in MM (Fig. 3E), without affecting the viability of the normal BMSCs (Fig. 3F).

Following exposure to TMP, we have also observed increase in G2–M and decrease in S-phases of the cell cycle (Fig. 4A), as well as late induction of apoptotic cell death (Fig. 4B) in MM cells. We have further observed activation of the mitochondrial apoptotic pathway by TMP via activation of caspase-9, -3, and -7 and PARP cleavage whereas caspase-8 was not activated (Fig. 4C). Change in the protein expression of survivin, a known antiapoptotic gene transcriptionally regulated by Sp1 (37), was confirmed following TMP treatment (Fig. 4C). Because caspase activation is a relatively late event, the reduction in survivin protein level during TMP-induced apoptosis seems to be an early event not mediated by caspase-dependent pathway. We have also observed decrease in cyclin-dependent kinase 1 (cdk1), an Sp1-regulated and cell-cycle–controlling gene following exposure to TMP (data not shown). Finally, because Sp1 is an important regulator of the expression of important angiogenic factors, including VEGF, which is believed to play a critical role in myeloma angiogenesis, we aimed to evaluate whether inhibition of Sp1 binding activity by TMP impairs VEGF production by MM cells. As shown in Supplementary Figure S2, 6-hour exposure to TMP was sufficient to decrease VEGF levels in the culture supernatant of both MM1S and U266.

Figure 4.

TMP induces apoptosis via caspase activation. A, flow cytometric cell-cycle analysis of BrdUrd incorporation was conducted after treatment of cells with the inhibitor for 24 hours. Data shown are the percentage of cells in the different phases of the cell cycle. B, MM1S cells were cultured in the absence or presence of TMP and apoptotic cell death was assessed by flow cytometric analysis following Annexin-V and PI staining. Top, the percentage of Annexin-V+/PI (early apoptosis) and Annexin-V+/PI+ (late apoptosis) cells at the indicated time. Bottom, a representative experiment (48 hours posttreatment) is shown. C, whole-cell lysate from MM1S cells treated with TMP (10 μmol/L) for the indicated time periods was subjected to Western blot analysis and probed with antibodies against caspase-3, -8, -9, -7, PARP, and survivin, with GAPDH as a loading control.

Figure 4.

TMP induces apoptosis via caspase activation. A, flow cytometric cell-cycle analysis of BrdUrd incorporation was conducted after treatment of cells with the inhibitor for 24 hours. Data shown are the percentage of cells in the different phases of the cell cycle. B, MM1S cells were cultured in the absence or presence of TMP and apoptotic cell death was assessed by flow cytometric analysis following Annexin-V and PI staining. Top, the percentage of Annexin-V+/PI (early apoptosis) and Annexin-V+/PI+ (late apoptosis) cells at the indicated time. Bottom, a representative experiment (48 hours posttreatment) is shown. C, whole-cell lysate from MM1S cells treated with TMP (10 μmol/L) for the indicated time periods was subjected to Western blot analysis and probed with antibodies against caspase-3, -8, -9, -7, PARP, and survivin, with GAPDH as a loading control.

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Thus, the antitumor activity of TMP may result, at least in part, from suppression of Sp1 activity and the consequent downregulation of downstream targets that are key to cell growth, apoptosis, and angiogenesis.

TMP inhibits MM cell growth and prolongs survival in vivo in a xenograft murine model of MM

Next, we investigated the anti-MM effect of Sp1 inhibition by TMP in vivo in 3 different murine xenograft models of human myeloma and in the SCID-hu model of human myeloma. In the xenograft models, we injected subcutaneously 3 different MM cell lines, U266, OPM2, and MM1S, in SCID mice. Following detection of tumor, mice were treated with either 50 mg/kg TMP or placebo subcutaneously daily for 3 weeks. Tumors were measured in 2 perpendicular dimensions once every 3 days. Treatment with TMP, compared with vehicle alone, significantly inhibited MM cell tumor growth in all 3 murine models of MM (Fig. 5A). As seen in Figure 5B, treatment with TMP also significantly prolonged survival in treated animals compared with control (P = 0.017); the median overall survival was 18 days in the control group and 28 days in the TMP-treated group. TMP-related toxicity was not observed in mice, as determined by daily evaluation of activity and overall body weight change during the course of treatment. Histologic examinations of tumor retrieved from MM1S-bearing mice confirmed decreased proliferation (as highlighted by Ki67 staining), significant tumor cell apoptosis (caspase-3 and TUNEL staining; ref. Fig. 5C), and decreased expression of survivin (Fig. 5D) following TMP treatment in vivo.

Figure 5.

TMP inhibits MM cell growth and prolongs survival in vivo. A, U266, OPM2, and MM1S cells were injected subcutaneously in 3 different cohorts of SCID mice. Following detection of tumor, mice were treated with either 2 mg TMP or placebo subcutaneously daily for 3 weeks. Tumors were measured in 2 perpendicular dimensions once every 3 days. B, survival was evaluated from the first day of treatment until death using the GraphPad analysis software. C, tumors were isolated from TMP-treated and control mice and sections were evaluated by histologic examinations following Ki67 staining showing decreased proliferation; caspase-3 and TUNEL stains showing significant tumor cell apoptosis. D, evaluation of survivin levels was done in the paraffin-embedded tumor sections stained with a combination of primary antibodies specific for CD138 and survivin. The tissue sections were imaged and the relative amount of survivin localized within tumor-cell nuclear compartment was determined using AQUA analysis. The data reflects the results of 12 fields imaged at 200× magnification per tumor. E, in the SCID-hu model, mice at the first detection of tumor were treated with vehicle (n = 3) or TMP (n = 3). Serum samples were collected weekly and level of shuIL-6R was measured by ELISA as a marker of tumor growth. Baseline values before treatment were not significantly different among groups.

Figure 5.

TMP inhibits MM cell growth and prolongs survival in vivo. A, U266, OPM2, and MM1S cells were injected subcutaneously in 3 different cohorts of SCID mice. Following detection of tumor, mice were treated with either 2 mg TMP or placebo subcutaneously daily for 3 weeks. Tumors were measured in 2 perpendicular dimensions once every 3 days. B, survival was evaluated from the first day of treatment until death using the GraphPad analysis software. C, tumors were isolated from TMP-treated and control mice and sections were evaluated by histologic examinations following Ki67 staining showing decreased proliferation; caspase-3 and TUNEL stains showing significant tumor cell apoptosis. D, evaluation of survivin levels was done in the paraffin-embedded tumor sections stained with a combination of primary antibodies specific for CD138 and survivin. The tissue sections were imaged and the relative amount of survivin localized within tumor-cell nuclear compartment was determined using AQUA analysis. The data reflects the results of 12 fields imaged at 200× magnification per tumor. E, in the SCID-hu model, mice at the first detection of tumor were treated with vehicle (n = 3) or TMP (n = 3). Serum samples were collected weekly and level of shuIL-6R was measured by ELISA as a marker of tumor growth. Baseline values before treatment were not significantly different among groups.

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In the SCID-hu model, 4 weeks after implanting human fetal bone in mice, human myeloma cells are injected in the bone and sIL-6R levels are measured in murine blood as a marker of myeloma tumor growth. SCID-hu mice were injected with TMP 5 times a week for 3 weeks after first detection of shuIL-6R in mice. We observed significant antitumor activity of TMP as measured by shuIL-6R levels in murine blood, suggesting MM cell growth inhibitory effects of TMP (Fig. 5E).

Alteration in expression and function of transcription factors has been frequently associated with neoplastic transformation. In this study, we provide experimental evidence that Sp1, a transcription factor that controls the number of cellular processes, plays an important regulatory role in MM cell growth and survival.

Although Sp1 is ubiquitously expressed, its nuclear localization observed in MM is functionally important. We have confirmed high Sp1 activity in MM cells by showing both increased DNA binding and increased Sp1-responsive promoter activity measured by luciferase reporter assay. We here further confirmed the effect of Sp1 on MM cell growth by both siRNA- and shRNA-mediated Sp1 knockdown, using multiple constructs.

MM cell–BMSC interaction induces transcription and secretion of cytokines and growth factors, which, in turn, confer proliferation and survival of MM cells. We here observed that this interaction leads to Sp1 activation; and inhibition of Sp1 activity by TMP led to suppression of MM cell–BMSC interaction–mediated growth of MM cells. Moreover, MM cell–BMSC interaction induces the activation of several signaling pathways, which, in turn, lead to Sp1 phosphorylation and transactivation of target genes. In line with these observations, we report that the MM cell–BMSC interaction–induced increase in both DNA binding and Sp1 transcriptional activity in MM cells was completely abrogated by inhibition of ERK pathway.

Compounds, such as TMP, able to disrupt the interaction between Sp1 and GC-rich motifs inhibit Sp1 activity. We have confirmed specific inhibition of both Sp1 binding and transcriptional activity in MM cells by TMP, including in the context of MM cell–BMSC interaction, without direct effect on Sp1 protein expression. Along with inhibition of Sp1 activity, we observed both in vitro and in vivo antimyeloma effects of TMP. Importantly, there was no significant synergistic effect when MM cells transfected with Sp1 siRNA were treated with TMP (data not shown), confirming specificity of mechanism of action of TMP. These results provide the rationale to evaluate efficacy of TMP in MM. TMP is currently in phase I/II clinical development for the treatment of glioma, treatment-refractory solid tumors, and cervical dysplasia (38).

Altered survivin expression may be one of the mechanisms by which Sp1 may affect MM cell survival. Survivin is an inhibitor of apoptosis and a possible modulator of the terminal effector phase of cell death/survival and is highly expressed in the number of human cancers including MM but not in normal adult human tissue. Transcription of survivin is modulated by Sp1 (39–43) and in pancreatic cancer cells, inhibition of Sp1 activity has been shown to decrease survivin expression and subsequently sensitize the cells to radiotherapy (43, 44).

The importance of Sp1 in myeloma is also supported by the recent observation that conventional and novel anti-MM drugs have direct effect on Sp1 activity. For example, it has been shown that HDAC1 could interact with Sp1 to regulate its activity (45) and HDAC inhibitors induce Sp1 activity (46), suggesting potential for synergism by using HDAC and Sp1 inhibitors together. Interestingly, bortezomib has been shown to inhibit DNA-binding activity of Sp1 and disrupt the physical interaction of Sp1/NF-κB (47, 48). In MM, bortezomib specifically downregulates the expression of class I HDACs through caspase-8–dependent degradation of Sp1 protein (49). Having defined the cellular, signaling, and the molecular mechanisms of sensitivity of MM to TMP, rationally designed combinations of conventional and novel agents to enhance cytotoxicity, to avoid or overcome drug resistance and to minimize adverse side effect profiles could be developed. More recently, it has been reported that both lenalidomide and pomalidomide upregulate Sp1 providing a rationale for their preclinical evaluation to increase cytotoxicity and overcome drug resistance (50).

In conclusion, we report significant role of Sp1 in myeloma cell growth and survival with its influence on clinical outcome in MM. Our preclinical in vitro and in vivo results suggest that specific inhibition of Sp1 activity may be an interesting potential therapeutic target alone and in combination with other agents in MM.

No potential conflicts of interest were disclosed.

K.C. Anderson, the Editor-in-Chief for Clinical Cancer Research, is a coauthor of this article. In keeping with the AACR's Editorial Policy, the article was peer reviewed and a member of the AACR's Publications Committee rendered the decision concerning acceptability.

M. Fulciniti and N.C. Munshi designed research; M. Fulciniti, P. Nanjappa, and S. Amin carried out research; S. Rodig carried out pathology; Y. Tai, R. Prabhala, S. Minvielle, P. Tassone, T. Hideshima, C. Li, and H. Avet-Loiseau contributed reagents/analytic tools; M. Fulciniti, K.C. Anderson, and N.C. Munshi analyzed data; and M. Fulciniti and N.C. Munshi wrote the manuscript.

This work was supported in part by DF/HCC myeloma SPORE Career Development Award to M. Fulciniti, Manton Foundation to N.C. Munshi, grants from the Dept. of Veterans Affairs Merit Review Awards, and grants from the NIH (RO1-124929 and PO1-155258 to N.C. Munshi and P50-100007 and PO1-78378 to N.C. Munshi and K.C. Anderson).

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.

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