Improved treatment strategies are required for bladder cancer due to frequent recurrence of low-grade tumors and poor survival rate from high-grade tumors with current therapies. Histone deacetylase inhibitors (HDACi), approved as single agents for specific lymphomas, have shown promising preclinical results in solid tumors but could benefit from identification of biomarkers for response. Loss of activating transcription factor 3 (ATF3) expression is a feature of bladder tumor progression and correlates with poor survival. We investigated the utility of measuring ATF3 expression as a marker of response to the HDACi pracinostat in bladder cancer models. Pracinostat treatment of bladder cancer cell lines reactivated the expression of ATF3, correlating with significant alteration in proliferative, migratory, and anchorage-dependent growth capacities. Pracinostat also induced growth arrest at the G0–G1 cell-cycle phase, coincident with the activation of tumor suppressor genes. In mouse xenograft bladder cancer models, pracinostat treatment significantly reduced tumor volumes compared with controls, accompanied by reexpression of ATF3 in nonproliferating cells from early to late stage of therapy and in parallel induced antiangiogenesis and apoptosis. Importantly, cells in which ATF3 expression was depleted were less sensitive to pracinostat treatment in vitro, exhibiting significantly higher proliferative and migratory properties. In vivo, control xenograft tumors were significantly more responsive to treatment than ATF3 knockdown xenografts. Thus, reactivation of ATF3 is an important factor in determining sensitivity to pracinostat treatment, both in vitro and in vivo, and could serve as a potential biomarker of response and provide a rationale for therapeutic utility in HDACi-mediated treatments for bladder cancer. Mol Cancer Ther; 15(7); 1726–39. ©2016 AACR.

This article is featured in Highlights of This Issue, p. 1425

Activating transcription factor 3 (ATF3) plays an indispensable role in different biologic processes ranging from regulating the immune response and coordinating gene expression responses to stress signals affecting metabolism and homeostasis. It plays a crucial role in diseases resulting in nerve injury and in cancer (1). All evidence suggests that ATF3 plays a dichotomous role in various stages of cancer progression. ATF3 has been shown to function as either an oncogenic suppressor or activator, depending on the degree of malignancy, context, and cell type (2, 3). ATF3 has a basic region leucine zipper (bZIP) DNA-binding domain, and by forming homodimers and heterodimers with other bZIP proteins, such as c-JUN and Jun B/D, ATF3 can function as either a transcriptional activator or repressor. ATF3 is composed of 181 amino acids, of which 21 serine or threonine, 1 tyrosine, and 17 lysine residues are potential sites for posttranslational modification, namely phosphorylation, acetylation, methylation, or ubiquitination (4, 5). Studies in breast, squamous skin carcinomas, and Hodgkin lymphoma suggest that ATF3 has an oncogenic role, both in tumor initiation and progression, whereas evidence from colorectal and esophageal cancers indicates that ATF3 suppresses colon and esophageal squamous cell tumorigenesis (6–10). Previously, we have shown that decreased ATF3 expression in bladder cancer is associated with cancer progression and is correlated with a reduced survival rate of patients with high-grade and metastatic tumors (11). Hence, there is an interest in determining whether reactivating ATF3 expression in bladder cancer has any potential therapeutic or biomarker utility.

Bladder cancer is the fifth most common cancer affecting men in Western countries, and the most common cancer of the urinary tract, with approximately 380,000 new cases and approximately 150,000 deaths reported each year, worldwide (12). More than 90% of bladder cancers are urothelial carcinomas; at diagnosis, approximately 60% of bladder cancers are classified as low-grade non–muscle invasive (NMIBC), whereas the remaining 40% are classified as high-grade muscle-invasive tumors (MIBC; refs. 13, 14). Treatment for NMIBC tumors involves transurethral resection, followed by the instillation of chemotherapeutic agents (mitomycin) or an immunostimulatory agent, Bacillus Calmette Guerin, to delay recurrence, and periodical cystoscopic surveillance (15, 16). Bladder-confined muscle-invasive tumors are treated by radical cystectomy with platinum-based chemotherapy, which results in approximately 50% survival for 5 years; the remaining 50% of patients develop metastatic disease and only approximately 5% of these survive for 5 years (17, 18). Because of the poor survival rate from high-grade tumors and the economic burden of the management of low-grade tumors, there is a greater demand for better alternative therapies, accompanied by biomarkers to predict treatment outcomes and improve the therapeutic management of bladder cancer.

In addition to genetic alterations, epigenetic changes are widely implicated in the development and progression of solid tumors, including bladder cancer (19, 20). Among the various posttranslational modifications associated with epigenetic changes, acetylation and deacetylation are well characterized and are regulated by two crucial enzymes: histone acetyltransferases (HAT) and histone deacetylases (HDAC), respectively. The transfer of acetyl groups by HATs follows acetylation of the amino tails of lysine residues in histones, with a subsequent reduction in affinity to the negatively charged DNA, thus resulting in enhanced transcriptional activity (21). In contrast, HDACs remove the acetyl group from lysine tails, resulting in a highly condensed chromatin and thus altered gene transcription. Aberrant acetylation of histones and nonhistone proteins (transcription factors, chaperone proteins, and nuclear receptors) plays a key role in regulating malignant traits, such as proliferation, growth arrest, angiogenesis, and apoptosis (22). In bladder cancer, there are reduced levels of global histone modifications compared with normal urothelium. Histone methylation levels decrease with disease severity and are correlated with advanced pathologic stages in NMIBC and MIBC in a large cohort of clinical samples (23, 24). Altered histone modifications are also evident in bladder cancer cell lines, and EZH2, a histone methyl transferase, is found to be frequently overexpressed in high-grade aggressive tumors and associated with transcriptional silencing of tumor suppressor gene E-cadherin (25, 26). The overwhelming evidence of histone modifications associated with disease progression provides the rationale to target HDACs using HDAC inhibitors (HDACi) in bladder cancer.

At least 15 different HDACis are currently in clinical trials, including representatives from four major chemical classes: hydroxamic acids, cyclic peptides, benzamides, and aliphatic acids. These HDACis have been shown to be effective in inhibiting the growth of various cancer cell lines and reducing malignancy in xenograft models (27). Pracinostat (SB939) is an orally available hydroxamic acid–based HDACi that is well tolerated, with highly favorable pharmacologic properties (28). Pracinostat has been shown to be superior to SAHA (an FDA-approved HDACi for cancer therapy), with double the potency against HDAC enzymes, and >3-fold increase in half-life and bioavailability, without affecting the activity of other zinc-binding enzymes in vitro. In in vivo studies, pracinostat has been shown to accumulate in the tumor bed and provide a sustained inhibition of tumor growth (94%, vs. 48% for SAHA). A large number of studies using HDACi have identified nonhistone proteins as substrates of HDACs, including transcription factors and signaling cytokines (29). However, little work has been done on potential markers that can be utilized to indicate how tumors respond to HDACi-mediated therapies to enable better therapeutic management.

In this study, we provide evidence that reactivation of ATF3 by pracinostat can be predictive of the outcome of HDACi-mediated treatment. We show a concentration- and time-dependent reexpression of ATF3 in vitro in a series of bladder cancer cells, with the subsequent restoration of nonmalignant traits in these cell lines, including a reduced proliferation rate, low colony formation capacity, decreased migration, and cell-cycle arrest. Reactivation of ATF3 also occurs in tumor cells that are in a nonproliferative state in mouse xenograft samples in the treatment-responding core of the tumors. Furthermore, depleted ATF3 expression in a stable cell line correlates with decreased sensitivity to pracinostat treatments in vitro and in vivo. Taken together, these data suggest that reactivation of ATF3 expression by pracinostat could serve as a potential biomarker of treatment response, enhancing therapeutic utility in HDACi-mediated treatment for advanced bladder cancer.

Cell culture

Human bladder cancer cell lines 5637 (grade 2), T24 (grade 3), J82 (grade 3), TCC-SUP (grade 4), and the human normal uroepithelial cell line SV-HUC1 were purchased from ATCC and were cultured in the recommended media following ATCC instructions. Upon receiving, lines were expanded and frozen aliquots were made in the first 2 months, and fresh aliquots were thawed before the lines reached 6 months in culture. TSU-Pr1 cells (grade 3) were sourced as described previously (30). The identity of all cell lines was authenticated with STR analysis using AmpFLSTR Identifiler Kit from Applied Biosystems at our MHTP Medical Genomics Facility and confirmed that no cross-contamination was present during this study.

Reagents and antibodies

For in vitro studies, pracinostat (SB939, MEI Pharma) stock solution was made by dissolving pracinostat at a concentration of 10 mmol/L in DMSO and stored in small aliquots at −20°C. For in vivo studies, pracinostat was dissolved in 0.5% methylcellulose (w/v) and 0.1% Tween 80 in water for oral dosing. The following primary antibodies were used in this study: ATF3 (sc-188), VEGF-A (sc-152) (Santa Cruz Biotechnology); acetyl-histone H3 (9675), acetyl-histone H4 (2591), acetyl-α-tubulin (5335), phospho RB (8180), RB1 (9303), cleaved caspase-3 (9661) (Cell Signaling Technology); p21 (ab7960), α-SMA (ab5694) (Abcam); and CD31 (DIA-310M) (Dianova).

Western blot analysis

Cells were washed twice with cold PBS, followed by lysis in modified RIPA buffer. Following centrifugation at 14,000 × g for 15 minutes, the supernatant was collected as total cell lysate. Lysates were quantified using a Pierce Protein Assay Kit (Thermo Scientific) according to the manufacturer's instructions. A total of 25 μg of protein lysate was separated on 12% or 15% SDS-PAGE gels and transferred onto nitrocellulose membranes (PerkinElmer) using standard Western blotting protocol. Targeted proteins were detected using IRDye conjugated (Rockland) or Alexa Fluor–labeled (Life Technologies) secondary antibodies against mouse 800 # 610-431-020, rabbit 800 # 611-131-122, mouse 680 # A21057, and rabbit 680 # A21076 and quantified using a Licor-Odyssey infrared imaging system (LI-COR Biosciences).

Cell viability assay

Cell viability was measured over a period of 6 days using a ViaLight Plus Kit (Lonza). Cells in the log growth phase were seeded in a 96-well plate at a predetermined density and allowed to adhere overnight before pracinostat treatment. Treatment media were replenished daily. Following the manufacturer's instructions, each day, cells were lysed and ATP-monitoring reagent was added to the lysate and luminescence read on a FLUOstar Optima from BMG LABTECH.

Colony formation assay

The minimum numbers of cells required for colony formation by bladder cancer cell lines T24 and TSU-Pr1 were optimized. Cells were treated for 3 days with varying concentrations of pracinostat or DMSO control. Cells (2 × 103) were plated with 0.3% Noble agar (Becton, Dickinson and Company) in DMEM supplemented with 10% FCS on top of a 0.6% agar layer in a 6-well plate and incubated for 21 days under regular culture conditions. At the completion of the culture period, colonies >500 μm were counted and photographed.

2D cell migration assay

Bladder cancer cells were grown to confluence and treated with pracinostat for 24 hours before a uniform scratch was made using a sterile 200-μL pipette tip. Floating cells were washed off with PBS, and new growth medium containing pracinostat or DMSO control was added. The migration of these cells was digitally photographed at 0, 6, 12, 16, 20, and 24 hours using a Nikon fluorescent digital imaging system. The area of the uncovered wound gap was quantified by ImageJ software. All experiments were done in triplicate.

Cell cycle and apoptosis analysis

Following treatment with pracinostat or in DMSO control for 24 hours, cells were harvested, washed twice in ice-cold PBS, and fixed in 70% ethanol at −20°C for >2 hours. Fixed cells were centrifuged and washed with PBS to remove fixative. Cells were stained with 20 μg/mL propidium iodide/0.1% Triton X-100 staining solution with 2.5 μg/mL RNase A for 15 minutes in the dark. Cell-cycle distribution was determined using BD Biosciences FACSCanto II Analyzer by collecting a minimum of 20,000 cells for each sample. The cell-cycle profile was defined by using FlowJo software (version 7.6.3). For apoptosis analysis, adherent and floating cells were collected and washed twice in PBS and resuspended in 1× binding buffer at a density of 1 × 106 cells/mL. FITC Annexin V (BD Pharmingen)–positive cells were analyzed using the BD Biosciences FACSCanto II Analyzer within 1 hour.

Human bladder cancer xenograft model

Six-week-old female BALB/c nude mice (Animal Resources Centre, Murdoch, Australia) received right flank injections of 1 × 106 TSU-Pr1 cells/mouse in 200 μL of a 1:1 mixed cell suspension and Matrigel (BD Biosciences). Tumor sizes were measured using digital calipers, and volumes were calculated according to the following formula: tumor volume (mm3) = (width2 × length)/2. Once tumor volume reached 200 mm3, the mice were randomized to receive pracinostat (100 or 50 mg/kg) as a single agent or vehicle (0.5% methyl cellulose and 1% Tween). The percent tumor growth inhibition (% TGI) was calculated according to the following formula: % TGI = (Cday aTday a)/(Cday aCday 1) × 100, in which Cday a is the median tumor volume for vehicle control group on day a, Tday a is the median tumor volume for treatment group on day a, and Cday 1 is the median tumor volume for the vehicle control group on day 1. All experiments involving animals were approved in advance by an Animal Ethics Committee at Monash University (Melbourne, Victoria, Australia) and were carried out in accordance with the “Australian code of Practice for the Care and Use of Animals for Scientific Purpose.”

IHC and immunofluorescence analysis

Tumors were harvested at the end of the 20-day treatment, fixed in formalin, and embedded in paraffin. Sections of 4 μm were made, followed by deparaffinization, rehydration, and heat-induced antigen retrieval in citrate buffer (pH 7.8) to uncover the epitope, and then primary antibodies or concentration-matched isotype controls were applied. Representative sections from both pracinostat-treated mice xenografts and vehicle control mice xenografts were analyzed for each marker. Images were taken for analysis using a Leica bright-field microscope. For correlation studies between ATF3 and Ki67, cells were identified as ATF3 positive via IHC on pracinostat-treated xenograft samples by the nuclear localization of ATF3 on the next immediately adjacent serial section and measured for Ki67 staining. Minimum staining thresholds used to categorize positive from negative staining were established using matching isotype control samples of the pracinostat-treated xenografts. ImageJ (version 1.49k) was utilized to measure and assess the staining intensities. For dual immunofluorescence analysis of CD31 and α smooth muscle actin (α-SMA), the specimens were digitized via whole-slide scanning with a Nikon C1 confocal microscope. Image analysis was performed using Imaris software (Bitplane AG). Fluorescent images were thresholded and masked for fluorescence intensity to remove autofluorescence and nonspecific signals. The total number of green or red positive blood vessels per image was counted as number of connected areas in the respective masked images. Subsequently, the red channel mask was applied to the green channel to obtain the total number of red fluorescent vessels also showing green fluorescence and vice versa.

Statistical analysis

Statistical analysis was performed using GraphPad Prism software version (version 6.0c) and displayed as the mean and SEM. Significant difference between two groups was analyzed using unpaired Student t test, and one-way ANOVA with Tukey post-test conducted for multiple comparisons, and P < 0.05 was considered as statistically significant. Correlation between variables was evaluated by Pearson rank correlation coefficient. Survival analysis was performed using Kaplan–Meier method and log-rank test.

HDACi pracinostat reactivates the expression of ATF3 in bladder cancer cell lines

Decreased expression of ATF3 is associated with bladder cancer progression and reduced patient survival rates (11). Loss of ATF3 expression as cancer progresses suggests that chromatin remodeling and the concomitant loss of histone acetylation may lead to epigenetic silencing of ATF3. To reestablish the normal acetylation pattern and thus to reactivate ATF3 expression, we treated a series of bladder cancer cell lines of different tumor grade with the HDACi pracinostat. The results (Fig. 1) show that pracinostat treatment reactivated ATF3 expression in a concentration- and time-dependent manner in all five cell lines tested (5637, T24, TSU-Pr1, J82, TCC-SUP). This was observed (data shown for three cell lines only) at both the mRNA (Fig. 1A) and protein level (Fig. 1B). A significant increase in ATF3 expression was observed in 5637 (P < 0.001) and TSU-Pr1 cells (P < 0.01), as early as 6 hours after pracinostat treatment, whereas significant changes were observed in J82 after 12 hours. In parallel, reexpression of ATF3 protein was also observed in the above cell lines. This coincided with progressive increases in the acetylation of the histone proteins H3 and H4. Densitometric analysis (Fig. 1B, ii–iv) confirmed the effect of concentration- and time-dependent activation of ATF3 in bladder cancer cells. Our results showed an increase in the acetylation of the nonhistone protein α-tubulin at 500 nmol/L pracinostat within 24 hours of treatment (Supplementary Fig. S1). No cell death was observed in any of the cell lines, even at the higher concentration of pracinostat (500 nmol/L) that restored ATF3 expression as evidenced by the absence of cleaved PARP (Supplementary Fig. S1, iii and iv). Analysis of apoptosis by Annexin V staining in TSU-Pr1 cells revealed that a similar proportion of cells (7%–9%) stained for early and late apoptosis, both in the control and in cells treated with 500 nmol/L pracinostat for 24 hours (Fig. 1C). Together, these results demonstrate that pracinostat treatment reestablishes the acetylation pattern of histone proteins and restores the expression of ATF3, and this reactivation does not induce cell death following short-term exposure to the drug.

Figure 1.

Pracinostat reactivates ATF3 mRNA and protein expression in vitro in a dose- and time-dependent manner. A, qPCR for detecting ATF3 reactivation in bladder cancer cell lines of different tumor grades: 5637 (i), TSU-Pr1 (ii), J82 (iii) at 6 and 12 hours of treatment with pracinostat. Pracinostat treatment significantly increased the relative expression of ATF3 in 5637 (***, P ≤ 0.001) for 500 nmol/L and in TSU-Pr1 (**, P ≤ 0.01) for ≥200 nmol/L cells by 6 hours of treatment and by 12 hours in J82 cells (*, P ≤ 0.05). Data shown are representative of three separate experiments (****, P ≤ 0.0001). B, Western blot analysis of the above cell lines treated with increasing concentration (20–500 nmol/L) of pracinostat for 12 and 24 hours (i). Reactivation of ATF3 in a dose- and time-dependent manner also coincides with progressive increase in acetylation of H3 and H4 (i). Data shown are representative of three separate experiments. Actin is used as an internal control to standardize the relative expression of ATF3 both at the mRNA and protein level. ii–iv, densitometric analysis corresponding to the immunoblot also quantifying a dose- and time-dependent ATF3 reactivation in the bladder cancer cell lines upon treatment with pracinostat (n = 3; *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001). C, scatter plot showing the apoptotic rate in TSU-Pr1 cells by double staining Annexin V and propidium iodide. The top two quadrants, which account for dead and late apoptosis cells, showed the percentage of cells was similar in control (i) and at the highest concentration (ii) of pracinostat (500 nmol/L) after 24 hours of treatment. The percentage of live cells in the Annexin and PI remained the same in both the cases (∼86.9%; n = 2).

Figure 1.

Pracinostat reactivates ATF3 mRNA and protein expression in vitro in a dose- and time-dependent manner. A, qPCR for detecting ATF3 reactivation in bladder cancer cell lines of different tumor grades: 5637 (i), TSU-Pr1 (ii), J82 (iii) at 6 and 12 hours of treatment with pracinostat. Pracinostat treatment significantly increased the relative expression of ATF3 in 5637 (***, P ≤ 0.001) for 500 nmol/L and in TSU-Pr1 (**, P ≤ 0.01) for ≥200 nmol/L cells by 6 hours of treatment and by 12 hours in J82 cells (*, P ≤ 0.05). Data shown are representative of three separate experiments (****, P ≤ 0.0001). B, Western blot analysis of the above cell lines treated with increasing concentration (20–500 nmol/L) of pracinostat for 12 and 24 hours (i). Reactivation of ATF3 in a dose- and time-dependent manner also coincides with progressive increase in acetylation of H3 and H4 (i). Data shown are representative of three separate experiments. Actin is used as an internal control to standardize the relative expression of ATF3 both at the mRNA and protein level. ii–iv, densitometric analysis corresponding to the immunoblot also quantifying a dose- and time-dependent ATF3 reactivation in the bladder cancer cell lines upon treatment with pracinostat (n = 3; *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001; ****, P ≤ 0.0001). C, scatter plot showing the apoptotic rate in TSU-Pr1 cells by double staining Annexin V and propidium iodide. The top two quadrants, which account for dead and late apoptosis cells, showed the percentage of cells was similar in control (i) and at the highest concentration (ii) of pracinostat (500 nmol/L) after 24 hours of treatment. The percentage of live cells in the Annexin and PI remained the same in both the cases (∼86.9%; n = 2).

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Pracinostat treatment and reactivation of ATF3 alters malignant behavior in vitro—viability, colony formation, and 2D migration

Next, we explored the functional significance of reactivation of ATF3 and reestablishment of histone acetylation in bladder cancer cell lines. To test phenotypic changes associated with the pracinostat treatment and reactivation of ATF3, different functional assays were performed. To analyze the viability of bladder cancer cell lines under pracinostat treatment, the cells were cultured with pracinostat (50, 100, or 500 nmol/L) for 6 days and cell viability assessed. Pracinostat significantly reduced the in vitro proliferation rate of all bladder cancer cells in a dose-dependent manner (Fig. 2A, i and ii). At a concentration as low as 100 nmol/L, pracinostat demonstrated a significant effect on the viability of four (5637, T24, TCC-SUP, and TSU-Pr1; P < 0.05) of the five cell lines we tested, whereas 500 nmol/L pracinostat resulted in a significant growth reduction in all five cell lines.

As anchorage-independent growth on a soft agar surface to produce colonies is characteristic of malignant transformation, we investigated whether treatment with pracinostat had any impact on this capacity of bladder cancer cells. Accordingly, cells were treated for 3 days, plated on agar at a low density, and incubated at 37°C for 21 days to allow the formation of spheres (Fig. 2B and Supplementary Fig. S2). The number of spheres formed at both 200 and 500 nmol/L of pracinostat was significantly reduced (P < 0.004 and P < 0.002, respectively) when compared with the DMSO control.

Figure 2.

Pracinostat treatment alters malignant properties of bladder cancer cells. A, assessment of viability by ViaLight assay on TSU-Pr1 cells treated with pracinostat demonstrated a dose-dependent growth inhibition of these cells over a period of 6 days, and fold change in relative luminescence unit was plotted in the graph (i and ii). All three doses tested showed a significant reduction in proliferation rate in these cells (P < 0.05; n = 4). ii, percentage of viable cells in four bladder cancer cell lines after 6 days of treatment with pracinostat at different concentrations. Data represent mean ± SEM; n = 4 per cell line. B, representative images of triplicate experiments of soft agar colony formation assay on TSU-Pr1 cells treated with pracinostat (200 and 500 nmol/L) for 3 days compared with the control cells (i). ii, quantification of spheres from soft agar assay. Colony number was reduced by approximately 75% (P = 0.008) in cells treated with 200 nmol/L pracinostat and approximately 85% (P = 0.005) in cells treated with 500 nmol/L pracinostat. Data shown are representative of at least three independent experiments. C, scratch wound assay in T24 (i) and 5637 (iii) cells treated with pracinostat. Representative images of different time points (0, 12, and 20 hours) are shown. ii and iv, cell migration was assessed by measuring the relative wound closure. Data represent mean ± SEM (n = 3; **, P ≤ 0.05; ***, P ≤ 0.001; ****, P ≤ 0.0001).

Figure 2.

Pracinostat treatment alters malignant properties of bladder cancer cells. A, assessment of viability by ViaLight assay on TSU-Pr1 cells treated with pracinostat demonstrated a dose-dependent growth inhibition of these cells over a period of 6 days, and fold change in relative luminescence unit was plotted in the graph (i and ii). All three doses tested showed a significant reduction in proliferation rate in these cells (P < 0.05; n = 4). ii, percentage of viable cells in four bladder cancer cell lines after 6 days of treatment with pracinostat at different concentrations. Data represent mean ± SEM; n = 4 per cell line. B, representative images of triplicate experiments of soft agar colony formation assay on TSU-Pr1 cells treated with pracinostat (200 and 500 nmol/L) for 3 days compared with the control cells (i). ii, quantification of spheres from soft agar assay. Colony number was reduced by approximately 75% (P = 0.008) in cells treated with 200 nmol/L pracinostat and approximately 85% (P = 0.005) in cells treated with 500 nmol/L pracinostat. Data shown are representative of at least three independent experiments. C, scratch wound assay in T24 (i) and 5637 (iii) cells treated with pracinostat. Representative images of different time points (0, 12, and 20 hours) are shown. ii and iv, cell migration was assessed by measuring the relative wound closure. Data represent mean ± SEM (n = 3; **, P ≤ 0.05; ***, P ≤ 0.001; ****, P ≤ 0.0001).

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We also tested the nature of the cell-to-cell interaction in 2D cultures of the cell lines, with and without pracinostat treatment, by measuring their scratch wound closure capacity. Wounds made in cells treated with 50 to 500 nmol/L pracinostat for 24 hours remained open at the time point when a wound made in DMSO control cells was closed. There was a significant delay in the wound closure process in T24 cells treated with pracinostat at a concentration of 200 nmol/L (Fig. 2C, i and ii) from 6 hours (P < 0.001, percentage of wound closure), and 40% of the wound remained open. Similarly, for 5637 cells, 25% of the wound remained open after treatment with 200 nmol/L pracinostat compared with the control. Taken together, the data suggest that pracinostat treatment in these bladder cancer cell lines restores a more nonmalignant phenotype.

Pracinostat treatment induces cell-cycle arrest at G0–G1 phase and activates tumor suppressor genes in bladder cancer cells, but not in normal urothelial cells

We have shown that pracinostat treatment over a period of 6 days in culture affects the viability of bladder cancer cell lines. To investigate whether this effect is due to impairment in their cycling capacity, we compared the cell-cycle pattern of bladder cancer cells, with and without pracinostat treatment. Treatment with pracinostat induced G0–G1 phase arrest in all bladder cancer cells tested (Fig. 3A, i, and B). The percentage of cells entering into the S-phase is strikingly reduced, with a subsequent accumulation of cells in G0–G1 phase in all cancer lines (Fig. 3B). In contrast, in normal urothelial cells SV-HUC1 (Fig. 3A, ii), the percentage of cells in the S-phase remained similar to the untreated controls even at the highest concentration (500 nmol/L) of pracinostat, with an increase in G2–M phase suggesting that the normal cells are relatively resistant to pracinostat-mediated treatment. This is consistent with previously published observations indicating that the proliferation inhibitory concentration of pracinostat is >100 μmol/L in normal human dermal fibroblast and ≤1 μmol/L in another 29 cancer cell lines (28).

Figure 3.

Pracinostat treatment accelerates cell-cycle arrest in cancer cells and activates tumor suppressor genes; normal urothelial cells demonstrate resistance. A, representative images of cell-cycle analysis by flow cytometry in TCC-SUP bladder cancer cells treated with 100 and 500 nmol/L pracinostat for 24 hours compared with the untreated control (i). The percentage of cells in S-phase in untreated TCC-SUP control cells was 22.19%, which was gradually reduced to 9.13% and 4.65% with 100 and 500 nmol/L pracinostat with a gradual increase in G0–G1 phase. Data shown are representative of three individual experiments. ii, representative images of cell-cycle distribution of normal urothelial cells, SV-HUC1 untreated, compared with 100 and 500 nmol/L pracinostat treated for 24 hours. The percentage of cells in S-phase remained similar in untreated and pracinostat-treated normal cells (∼20%). B, cell-cycle distribution of three different bladder cancer cells; the quantitation is presented as mean ± SEM (n = 3). C, immunoblot analysis of tumor suppressor genes associated with bladder cancer (RB1 and p21). Expression analysis in four different bladder cell lines indicated a dose-dependent transition from hyperphosphorylated (inactive) to hypophosphorylated (active; 106 kDa) form of RB1. Representative of a replicated experiment (n = 2).

Figure 3.

Pracinostat treatment accelerates cell-cycle arrest in cancer cells and activates tumor suppressor genes; normal urothelial cells demonstrate resistance. A, representative images of cell-cycle analysis by flow cytometry in TCC-SUP bladder cancer cells treated with 100 and 500 nmol/L pracinostat for 24 hours compared with the untreated control (i). The percentage of cells in S-phase in untreated TCC-SUP control cells was 22.19%, which was gradually reduced to 9.13% and 4.65% with 100 and 500 nmol/L pracinostat with a gradual increase in G0–G1 phase. Data shown are representative of three individual experiments. ii, representative images of cell-cycle distribution of normal urothelial cells, SV-HUC1 untreated, compared with 100 and 500 nmol/L pracinostat treated for 24 hours. The percentage of cells in S-phase remained similar in untreated and pracinostat-treated normal cells (∼20%). B, cell-cycle distribution of three different bladder cancer cells; the quantitation is presented as mean ± SEM (n = 3). C, immunoblot analysis of tumor suppressor genes associated with bladder cancer (RB1 and p21). Expression analysis in four different bladder cell lines indicated a dose-dependent transition from hyperphosphorylated (inactive) to hypophosphorylated (active; 106 kDa) form of RB1. Representative of a replicated experiment (n = 2).

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We further investigated the changes in the expression of tumor suppressor genes implicated in bladder cancer, such as retinoblastoma (RB1) and p21. RB1 has been identified as a predominant growth and tumor suppressor in bladder cancer (31). Hyperphosphorylated RB1 is inactive, with phosphorylated serine residues, and is unable to bind E2F and control the cell cycle. When RB1 is hypophosphorylated (active RB1), it binds to E2F and prevents transcription of target genes. Treatment with pracinostat resulted in an increase in the hypophosphorylated (active) RB1 in a dose-dependent manner in three of the bladder cancer cell lines we tested (Fig. 3C), suggesting that an increase in expression of hypophosphorylated RB1 may have an impact on cell-cycle arrest in bladder cancer cells. In J82, a grade 3 tumor cell line, RB1 expression is lost in control cells but pracinostat treatment restored hypophosphorylated RB1 (active form) expression. Furthermore, overexpression of ATF3 in J82 cells induced the expression of RB1, and a concomitant increase in G0–G1 phase cell-cycle arrest (Supplementary Fig. S3). The tumor suppressor gene, cyclin-dependent kinase inhibitor P21WAF1/CIP1, was also reactivated in three of the four cell lines (5637, T24, and TSU-Pr1) treated with 500 nmol/L pracinostat (Fig. 3C).

Pracinostat inhibits bladder cancer growth in vivo

To investigate the biomarker potential of measuring the reactivation of ATF3 expression by pracinostat in vivo, bladder cancer cell lines TSU-Pr1 (Fig. 4, i) and T24 (Supplementary Fig. S4) were injected into the flanks of nude mice. Full engraftment of TSU-Pr1 cells was observed 2 to 3 weeks after injection. Mice were initially randomized to receive vehicle control or 100 mg/kg pracinostat in a cycle of 5 days on 2 days off, orally for 21 days. The dosage was chosen based on published observations where 97% of tumor growth inhibition (TGI) was recorded for colon cancer xenografts with a maximum of 16.6% body weight loss in mice over a period of 3 weeks (28). The results (Fig. 4) show tumor volumes in pracinostat-treated mice were significantly smaller when compared with the vehicle control group, exhibiting greater tolerability to the drug over 20 days as indicated by body weight loss (Fig. 4A, i, ii, and B, ii). In pracinostat-treated mice, the median tumor volume was 190.04 mm3 on day 20 of treatment, compared with 529.08 mm3 for the vehicle control group (Fig. 4A, ii). This is equivalent to 98% (P < 0.0001) TGI in pracinostat-treated mice (100 mg/kg) compared with the vehicle control group. Histologic changes between the vehicle control and pracinostat-treated mice were analyzed by hematoxylin and eosin staining (Fig. 4B, i). Control tumors exhibited a uniform distribution of viable cells stained with hematoxylin, whereas the sections from pracinostat-treated samples demonstrated large areas of necrotic or late apoptotic cells characteristically stained by eosin only.

Figure 4.

Pracinostat treatment inhibits in vivo tumor growth and promotes ATF3 reexpression correlating with tumor response. A, efficacy of pracinostat (100 mg/kg) as a single agent for treatment of mice bearing bladder xenografts for 20 days: percentage body weight (i; mean weight ± SD, n = 8) and tumor volume (ii). The tumor volumes were significantly different in pracinostat-treated groups compared with the vehicle groups (****, P ≤ 0.0001). Data points represent the mean tumor volume for each group (n = 8 mice/group, error bars ± SD). B, representative images demonstrating histologic changes in vehicle control mice compared with pracinostat (100 mg/kg) treated mice for 3 weeks (i); hematoxylin and eosin staining. Image magnification, 40×; scale bar, 100 μm. ii, representative macroscopic images of tumors from vehicle-treated mice and pracinostat-treated mice at the end of 20 days. C, immunohistochemical analysis of ATF3 and Ki67 in mouse xenograft samples after treatment (i). Serial sections were stained for ATF3 and Ki67, respectively, after 5 days (top) and 20 days (middle) of pracinostat treatment. Cells expressing ATF3 (thin arrowhead) are mainly in the treatment-responding edge of the tumor, and cells expressing Ki67 (proliferating; thick arrowhead) are away from the responding edge. Representative images for vehicle control xenograft samples for ATF3 and Ki67. Image magnification, 40×; scale bar, 100 μm. ii, scatter plot and linear regression analysis of staining intensity of the cells in the treatment-responding edge of the xenograft samples (n = 10). Cells identified as ATF3 positive were located, and mean intensity was determined using ImageJ and measured for Ki67 staining.

Figure 4.

Pracinostat treatment inhibits in vivo tumor growth and promotes ATF3 reexpression correlating with tumor response. A, efficacy of pracinostat (100 mg/kg) as a single agent for treatment of mice bearing bladder xenografts for 20 days: percentage body weight (i; mean weight ± SD, n = 8) and tumor volume (ii). The tumor volumes were significantly different in pracinostat-treated groups compared with the vehicle groups (****, P ≤ 0.0001). Data points represent the mean tumor volume for each group (n = 8 mice/group, error bars ± SD). B, representative images demonstrating histologic changes in vehicle control mice compared with pracinostat (100 mg/kg) treated mice for 3 weeks (i); hematoxylin and eosin staining. Image magnification, 40×; scale bar, 100 μm. ii, representative macroscopic images of tumors from vehicle-treated mice and pracinostat-treated mice at the end of 20 days. C, immunohistochemical analysis of ATF3 and Ki67 in mouse xenograft samples after treatment (i). Serial sections were stained for ATF3 and Ki67, respectively, after 5 days (top) and 20 days (middle) of pracinostat treatment. Cells expressing ATF3 (thin arrowhead) are mainly in the treatment-responding edge of the tumor, and cells expressing Ki67 (proliferating; thick arrowhead) are away from the responding edge. Representative images for vehicle control xenograft samples for ATF3 and Ki67. Image magnification, 40×; scale bar, 100 μm. ii, scatter plot and linear regression analysis of staining intensity of the cells in the treatment-responding edge of the xenograft samples (n = 10). Cells identified as ATF3 positive were located, and mean intensity was determined using ImageJ and measured for Ki67 staining.

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ATF3 expression correlates with tumor response to pracinostat in vivo

Consistent with the observation made in vitro, oral administration of pracinostat reactivated ATF3 expression in xenograft samples. IHC analysis on tissue samples collected from early to late stage of therapy clearly demonstrated nuclear reexpression of ATF3 in the treatment-responding core of the tumor tissue in pracinostat-treated samples (Fig. 4C, i). Intriguingly, the majority of cells expressing ATF3 were in a nonproliferative state, indicated by loss of expression of the proliferative nuclear marker Ki67. Cells expressing ATF3 were detected only in the treatment-responding core of the tumor, which is largely devoid of Ki67 expression, and actively proliferating cells (Ki67-positive cells) were mainly located toward the outer edge of the xenograft sample. The staining intensity of the cells expressing ATF3 in the treatment-responding edge of the tumor samples was scored and data analyzed to evaluate whether ATF3-expressing cells were in a nonproliferating state. The mean intensity of cells expressing ATF3 was negatively correlated to Ki67 expression (Fig. 4C, ii; r = −0.66, P < 0.0001). As expected, tissue samples from the vehicle control group stained negatively for ATF3 and demonstrated abundant expression of Ki67.

To extend our findings on tumor response to treatment with pracinostat, we examined markers of apoptosis and angiogenesis in xenograft samples. Caspase-3 has been reported to play an important role in apoptosis, especially the active form of the enzyme, cleaved caspase-3 (32). Immunohistochemical analysis of xenograft samples demonstrated that both cytoplasmic and perinuclear cleaved caspase-3 were expressed by the remaining tumor cells of pracinostat-treated samples compared with the vehicle control, suggesting that there is a marked induction of apoptosis signals in the treatment group (Fig. 5A).

Figure 5.

Pracinostat treatment induces apoptosis and antiangiogenesis in xenografts. A, representative images of IHC sections of cleaved caspase-3 staining in vehicle- and pracinostat-treated mice. Image magnification, 40×; scale bar, 100 μm. B, representative images of immunohistochemical analysis of endothelial cell marker CD31 on vehicle-treated and pracinostat-treated mice tumor samples. ii, microvessel density was quantified by identifying neovascular “hot spots” and unbiased scoring of the number of vessels stained for CD31 in representative sections of vehicle control and pracinostat-treated mice for 3 weeks (**, P ≤ 0.01). Quantification of data from n = 8 per group; error bars ± SEM. C, dual immunofluorescence detection of CD31 (red) and α-SMA (green) in vehicle- and pracinostat-treated mouse xenograft samples for vascular integrity (i). ii, quantification of CD31 and SMA scoring was determined as described in Materials and Methods (***, P ≤ 0.001; ****, P ≤ 0.0001). D, representative images of IHC sections of VEGF-A staining in vehicle- and pracinostat-treated mice. Image magnification, 40×; scale bar, 100 μm.

Figure 5.

Pracinostat treatment induces apoptosis and antiangiogenesis in xenografts. A, representative images of IHC sections of cleaved caspase-3 staining in vehicle- and pracinostat-treated mice. Image magnification, 40×; scale bar, 100 μm. B, representative images of immunohistochemical analysis of endothelial cell marker CD31 on vehicle-treated and pracinostat-treated mice tumor samples. ii, microvessel density was quantified by identifying neovascular “hot spots” and unbiased scoring of the number of vessels stained for CD31 in representative sections of vehicle control and pracinostat-treated mice for 3 weeks (**, P ≤ 0.01). Quantification of data from n = 8 per group; error bars ± SEM. C, dual immunofluorescence detection of CD31 (red) and α-SMA (green) in vehicle- and pracinostat-treated mouse xenograft samples for vascular integrity (i). ii, quantification of CD31 and SMA scoring was determined as described in Materials and Methods (***, P ≤ 0.001; ****, P ≤ 0.0001). D, representative images of IHC sections of VEGF-A staining in vehicle- and pracinostat-treated mice. Image magnification, 40×; scale bar, 100 μm.

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Neovascularization is vital for in vivo tumor growth, and reduction of microvessel density is a measure of antiangiogenic activity. Endothelial cell marker CD31 or platelet endothelial cell adhesion molecule-1 (PECAM-1) is commonly used to quantify angiogenesis in xenograft models (33). Immunohistochemical analysis of CD31 in xenograft samples indicated a profound increase in the number of blood vessels in the vehicle control group compared with the pracinostat-treated samples (Fig. 5B, i). Quantification of microvessel density confirmed that there was a significant reduction in the number of microvessels in pracinostat-treated mice compared with the vehicle control mice (Fig. 5B, ii; P < 0.0012). To further support the finding of an antiangiogenic effect for pracinostat, we examined xenograft samples using dual immunofluorescence staining for CD31 (red) and α-SMA (green), thereby depicting vascular integrity in these samples (Fig. 5C, i). Association of α-SMA–expressing pericytes with vascular endothelial cells indicates vascular maturation (34). Immunofluorescence analysis of tumor samples demonstrated that the numbers of newly formed blood vessels without pericytes and vessels with pericytes are significantly higher in the vehicle control group (P < 0.0001), indicating actively growing tumors in the untreated group (Fig. 5C, ii). In the pracinostat-treated group, the endothelial cells and pericytes were strikingly reduced, indicating a treatment-associated reduction in angiogenic activity. Interestingly, the number of pericytes was slightly higher than the number of endothelial cells in pracinostat-treated tumor samples, suggesting the presence of collapsed vessels following treatment, although this was not significant (Fig. 5C, ii). The expression of VEGF-A, a potent angiogenic agent that stimulates the full cascade of events required for angiogenesis, was also analyzed (35). In vitro pracinostat treatment for 24 hours significantly reduced the expression of VEGF-A expression in all three cell lines tested (Supplementary Fig. S5). Histology analysis of xenograft tumor samples demonstrated areas with abundant expression of VEGF-A in controls, which was undetectable in pracinostat-treated samples (Fig. 5D). Taken together, our data clearly demonstrate that ATF3 reexpression in vivo is a marker of tumor response to pracinostat treatment, indicated by the activation of antiangiogenesis and apoptosis.

Reactivation of ATF3 is integral to the pracinostat-mediated tumor response in vivo and in vitro

To investigate whether reexpression of ATF3 is central to determining tumor response in HDACi-mediated anticancer therapy, we generated stable knockdown TSU-Pr1cell lines by transfection with short hairpin RNA (shRNA) targeting human ATF3 or a nontargeting control sequence to generate a control cell line (Supplementary Fig. S6). To understand the significance of the treatment in the presence and absence of ATF3, we performed in vitro functional assays, viability, wound healing, and colony formation on the stable cell lines (Fig. 6A and B and Supplementary Fig. S6). To compare the viability of ATF3-depleted cells and control cells in the presence of pracinostat, the cells were cultured in varying concentrations of pracinostat (50, 100, 200, and 500 nmol/L) for a period of 6 days and analyzed. Although pracinostat treatment had an antiproliferative effect on both lines, the viability of ATF3-depleted cells was significantly higher than control cells over time (Fig. 6A, i), particularly at lower concentrations of drug (50, 100, and 200 nmol/L; data shown only for 100 nmol/L). Pracinostat-treated (100 nmol/L) ATF3 knockdown cells exhibited significantly higher viability from day 4 compared with the shControl cells (P < 0.004 for day 4, P < 0.0001 for days 5 and 6). Protein lysates were analyzed by Western blot to monitor the reactivation of ATF3 in these cells over 6 days with pracinostat treatment and clearly showed less reactivation of ATF3 in the knockdown cells compared with the control (Fig. 6A, ii).

Figure 6.

ATF3 reactivation is integral in determining the sensitivity to HDACi treatment. A, assessment of viability by ViaLight assay on stable control cells and ATF3-depleted cells treated with pracinostat over a period of 6 days at a concentration of 100 nmol/L (i). From day 4, the proliferation rate was significantly higher in knockdown cells compared with control cells (**, P ≤ 0.01; ***, P ≤ 0.001). Data represent results of one of three individual experiments. ii, immunoblot analysis confirmed the reactivation of ATF3 in control cells compared with knockdown cells over 6 days in treatment with pracinostat (n = 3). B, representative pictures of scratch wound assay on cells: stable control and ATF3 knockdown cells treated with 100 nmol/L pracinostat or untreated. Scratches were made on the confluent cells, and wound closure was monitored over 16 hours. Data represent the results of one of three separate experiments. C, in vivo tumor growth and survival in mice with bladder xenografts with control cells and knockdown cells and treated with 50 mg/kg or vehicle control. i, tumor volume was significantly smaller in pracinostat-treated shControl (shctrl) cells (***, P ≤ 0.001). Data points represent the mean tumor volume for each group (n = 8 mice/group, error bars ± SD). ii, percentage body weight, showing no significant difference in the weights between treatment and control groups (n = 8 mice/group; error bars ± SD). iii, Kaplan–Meier survival curves showing a significant difference in the survival between the shcontrol and shATF3 treated with pracinostat (***, P ≤ 0.001; n = 8 mice per group). D, representative images of immunohistochemical analysis of xenografts samples of shATF3 and shcontrol for acetyl histone 3 (i) and Ki67 (ii). Image magnification, 40×; scale bar, 100 μm.

Figure 6.

ATF3 reactivation is integral in determining the sensitivity to HDACi treatment. A, assessment of viability by ViaLight assay on stable control cells and ATF3-depleted cells treated with pracinostat over a period of 6 days at a concentration of 100 nmol/L (i). From day 4, the proliferation rate was significantly higher in knockdown cells compared with control cells (**, P ≤ 0.01; ***, P ≤ 0.001). Data represent results of one of three individual experiments. ii, immunoblot analysis confirmed the reactivation of ATF3 in control cells compared with knockdown cells over 6 days in treatment with pracinostat (n = 3). B, representative pictures of scratch wound assay on cells: stable control and ATF3 knockdown cells treated with 100 nmol/L pracinostat or untreated. Scratches were made on the confluent cells, and wound closure was monitored over 16 hours. Data represent the results of one of three separate experiments. C, in vivo tumor growth and survival in mice with bladder xenografts with control cells and knockdown cells and treated with 50 mg/kg or vehicle control. i, tumor volume was significantly smaller in pracinostat-treated shControl (shctrl) cells (***, P ≤ 0.001). Data points represent the mean tumor volume for each group (n = 8 mice/group, error bars ± SD). ii, percentage body weight, showing no significant difference in the weights between treatment and control groups (n = 8 mice/group; error bars ± SD). iii, Kaplan–Meier survival curves showing a significant difference in the survival between the shcontrol and shATF3 treated with pracinostat (***, P ≤ 0.001; n = 8 mice per group). D, representative images of immunohistochemical analysis of xenografts samples of shATF3 and shcontrol for acetyl histone 3 (i) and Ki67 (ii). Image magnification, 40×; scale bar, 100 μm.

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Two-dimensional cell migration and cell–cell interaction was also analyzed by measuring wound closure in control and knockdown cells (Fig. 6B). The rate of wound closure was faster in untreated and pracinostat-treated knockdown cells compared with the control cells. The cells in which ATF3 expression was silenced underwent wound closure at a much earlier time point (12 hours), compared with control cells in the presence of pracinostat. In control cells, 50% of the wound remained open at the time it was closed in knockdown cells. Together, the in vitro data strongly suggest that the cells in which the expression of ATF3 was silenced fail to respond to pracinostat treatment.

To extend our findings, we explored the functional significance of the loss of ATF3 expression with pracinostat treatment in vivo. Control cells and ATF3 knockdown cells were injected into the flank of the nude mice and pracinostat administrated orally daily for up to 30 days at 50 mg/kg, a dose that did not result in body weight loss (Fig. 6C, ii). In pracinostat treatment groups, the tumor volumes of ATF3-depleted xenografts were significantly larger than in the shControl group (<400 mm3 at the end of the experiment; P < 0.0001) and reached the endpoint volume (≥800 mm3) earlier, at 25 days after treatment (Fig. 6C, i). The tumor volume data showed that pracinostat treatment delayed the rate of tumor growth in mice with ATF3-depleted xenografts compared with matching vehicle controls (although this was not significant; P = 0.6470). In the shControl xenografts, tumor volumes were significantly different between pracinostat-treated mice with the vehicle control (P < 0.0001). Importantly, the overall tumor growth rates in ATF3-depleted xenografts were higher than in the shControl xenografts. The association between ATF3 expression and survival rate was also evaluated in the pracinostat-treated groups. Pracinostat treatment induced ATF3 expression in control xenografts, which responded well to treatment with 100% of mice in this group surviving to the end of the experiment, whereas the median survival in the ATF3-depleted group was 22.5 days. Kaplan–Meier survival analysis showed a significant difference in the survival curves for these two groups (Fig. 6C, iii; P < 0.0001). The activity of pracinostat in delaying the rate of tumor growth in the treatment group was further examined by assessing the acetylation pattern of H3 in the xenograft samples. Immunohistochemical analysis of ATF3 knockdown xenograft samples demonstrated increased expression of acetylated H3 in pracinostat-treated samples compared with matching vehicle control samples (Fig. 6D, i). We then tested whether the increased acetylation pattern was reflected in the proliferation status of pracinostat-treated ATF3-depleted xenografts samples. However, immunohistochemical analysis of proliferation marker Ki67 demonstrated that the number of cells in actively proliferating status is similar in vehicle control and the pracinostat-treated ATF3-depleted group (Fig. 6D, ii). These findings reaffirm the importance of reexpression of ATF3 in bladder xenografts to make them sensitive to the HDACi treatment. Taken together, the data from in vitro and in vivo studies clearly demonstrate that reactivation of ATF3 by pracinostat is a determining factor in the tumor response to this HDACi therapy and confirm ATF3 as a biomarker of tumor response. Furthermore, our data strongly support the proposition that regulation of ATF3 expression by pracinostat may be of therapeutic utility for bladder cancer patients.

Identification of biomarkers with the potential to predict tumor response to therapy is a critical and important step toward better clinical planning and therapeutic management, particularly considering the heterogeneity of solid tumors. Here, we show that the transcription factor ATF3 can be utilized as a potential marker of the response to HDACi-mediated therapy in bladder cancer. Using the pan-HDACi pracinostat, we demonstrate that ATF3 expression was reactivated in vitro in a series of bladder cancer cells of different grade and origin. Similarly, histology data from an in vivo mouse model also reaffirmed the reexpression of ATF3 in the responding core of the tumor from early- to late-stage therapy. Furthermore, stable cells with depleted ATF3 expression were significantly more proliferative in vitro and in vivo and significantly less sensitive to pracinostat treatment. Our findings show that regulation of ATF3 expression by the HDACi pracinostat has a therapeutic potential and that reexpression of ATF3 is important in determining treatment outcome, particularly in advanced cancers. Previous reports (36) have identified a direct association of ATF3 with HDAC1 in regulating acetylation patterns and chromatin structures in innate immune systems. Earlier studies in colorectal cancer using HDACi as cytotoxic agents show that ATF3 contributes in enhancing the synergistic effect of HDACi, in combination with cisplatin (37). In addition, in p53-deficient colorectal cancers, the synergistic anticancer effect of ATF3 and HDACi has been previously shown, where ATF3 is essential in inducing death receptor 5 (DR5) and a proapoptotic effect (38). We show that treatment with pracinostat reactivates the expression of ATF3 without inducing cellular death in bladder cancer cells, and that this is essential in defining the subsequent alteration of their malignant traits. The in vivo data clearly show reexpression of ATF3 in xenograft samples in the responding core of the tumor during pracinostat treatment, indicating that this is essential for predicting treatment outcome. Moreover, these results are the first observations of the reactivation of ATF3 in real time with HDACi treatment in nonproliferating cells in the responding core of the tumor.

Pracinostat has been previously shown to be on average 2-fold more potent than SAHA as an inhibitor of HDAC enzymes and to have a 3-fold longer half-life (28). It is noteworthy that as we show in Fig. 1C, the dosage of pracinostat we used in vitro for a short period (24 hours) was sufficient to reactivate the expression of ATF3 in parallel with acetylation of H3 and H4 without inducing cellular death at the same time point in vitro, suggesting the treatment facilitates reestablishment of a global acetylation pattern in malignant cells, which subsequently results in the formation of euchromatin and the activation of transcription factors, including ATF3. Following this reactivation of ATF3, prolonged treatment with pracinostat triggers a cascade of events including antiangiogenesis (Fig. 5 and Supplementary Fig. S5). Previous studies have identified the antiangiogenic properties of other HDACis, including trichostatin A, SAHA, and valproic acid (39–41). The in vitro data we present in cancer cell lines and in vivo results in mouse xenograft samples show that the proangiogenic factor VEGF-A (one of the essential factors shown to facilitate angiogenesis) secreted by tumor cells is inhibited by pracinostat treatment. Furthermore, staining for blood vessels and pericytes was also significantly reduced with pracinostat treatment in vivo, compared with controls. Perivascular smooth muscle cells are essential for vascular integrity, by providing physiologic and mechanical support for blood vessels (42). Untreated bladder xenografts showed a significantly higher number of blood vessels and pericytes than pracinostat-treated tumor samples (Fig. 5). A combination of blood vessels with and without a lining of pericytes suggests the presence of an actively growing tumor aided by a reactive stroma in control xenografts. Interestingly, we observed that pracinostat treatment induced damage to the structure of blood vessels, with a slight increase in the number of pericytes, which may have formed to support the endothelial lining before the start of treatment. Although we clearly demonstrate inhibition of angiogenesis associated with pracinostat treatment, this antiangiogenesis effect could be a consequence of ATF3 reactivation or independent of ATF3, mediated by pracinostat.

HDACis have been shown to impose the G1 phase arrest and subsequent activation of P21 in malignant cells (43, 44). In accord with this, cell-cycle analysis of pracinostat-treated cells indicates G0–G1 phase arrest and restricted entry into the S-phase, coincident with dephosphorylation of RB1 and induction of p21 (Fig. 3). Overexpression of ATF3 in J82 cells correspondingly induced the expression of RB1 and subsequently resulted in G0–G1 phase arrest in these cells, suggesting that ATF3 has a direct impact on RB1 expression. In silico analysis of the RB1 promoter reveals an ATF-binding site, suggesting this possibility (Supplementary Fig. S3). Pracinostat treatment reactivated the expression of ATF3 and consequently resulted in cell-cycle arrest in vitro. Similarly, in vivo results demonstrated that the ATF3-expressing cells are localized to the treatment-responding core and are negatively correlated to Ki67-expressing proliferating cells. Together, these observations suggest that the expression of RB1 is regulated by ATF3, and consequently, the ATF3-expressing cells are less proliferative in vitro and in vivo. Although the treatment with pracinostat induces cell-cycle arrest in all bladder cancer cells, the percentage of normal cells (HUC1, normal urothelial cells) in S-phase remains unaffected by pracinostat treatment; normal cells progress through the cell cycle and there was an increase of cells in G2–M phase. G2–M arrest and the resistance of noncancerous cells to HDACi have been described previously (45) and reflect the presence of a G2 checkpoint in normal cells, which is absent in the majority of cancer cells; abolishing the G2 checkpoint using checkpoint kinase 1 abrogates this resistance (46). Our findings in bladder cancer that the reactivation of p21 is in parallel to the reactivation of ATF3 are in contrast to the observation made in epidermoid carcinoma, where ATF3 acts as an oncogenic activator (47).

In vitro functional assays with pracinostat in our panel of bladder cancer cells, including viability, 2D migration, and cell-cycle analysis, showed that there was a concentration-dependent sensitivity in every cell line. However, the degree of sensitivity to pracinostat varied in the different cell lines. Three of the five lines showed significantly reduced viability at a concentration of 100 nmol/L, whereas the other two cell lines were less sensitive, independent of their tumor grades from which they derived. TCC-SUP and TSU-Pr1 cells showed a significant delay in wound closure at concentrations of 50 and 100 nmol/L pracinostat, whereas in other cell lines, a delay in wound closure occurred at concentrations >100 nmol/L (Fig. 2 and Supplementary Fig. S2). Heterogeneity within cancer cell populations is a common survival advantage against anticancer therapy, including HDACi. In addition, previous studies have proposed different mechanisms of resistance to HDACi-mediated therapy, including differing levels of HDAC enzymes in different cells (48), HDACi efflux mechanism, and the level of P-glycoprotein (49, 50), and changes in signaling pathways, including antiapoptotic factors, such as BCL-2 and NF-κB (51, 52). Although the detailed mechanisms underlying differences in sensitivity were not explored, an increase in mRNA expression of class 1, 2B, and 4 HDAC enzymes (Supplementary Fig. S7) may contribute toward cell line resistance to pracinostat.

In summary, we have identified ATF3 as a potential marker of response to pracinostat (HDACi)-mediated bladder cancer therapy. We have demonstrated that the reactivation of ATF3 by pracinostat is a decisive factor in determining treatment outcome in vitro and in in vivo mouse xenograft models, thus supporting a novel therapeutic strategy for the treatment of patients with advanced bladder tumors where ATF3 expression is lost.

D.P. Gold has ownership interest (including patents) in MEI Pharma. No potential conflicts of interest were disclosed by the other authors.

Conception and design: D. Sooraj, D. Xu, D.P. Gold, B.R.G. Williams

Development of methodology: D. Sooraj, D. Xu, B.R.G. Williams

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): D. Sooraj, B.R.G. Williams

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): D. Sooraj, D. Xu, J.E. Cain, B.R.G. Williams

Writing, review, and/or revision of the manuscript: D. Sooraj, D. Xu, J.E. Cain, D.P. Gold, B.R.G. Williams

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): B.R.G. Williams

Study supervision: D. Xu, B.R.G. Williams

The authors thank Drs. Jacqueline Donoghue and Howard Yim (Hudson Institute of Medical Research) for advice and helpful discussion, Samantha Jayasekara for technical support for in vivo studies, and Camden Lo and Kirstin Elgass (Monash Micro Imaging) for help with image acquisition and analysis. The authors also thank Dr. Frances Cribbin for assistance with the preparation of this article.

This work was supported by grants from the National Health and Medical Research Council of Australia (APP1066665; to B.R.G. Williams and D. Xu) and the Victorian Government's Operational Infrastructure Support Program. D. Sooraj was supported by an Australian Postgraduate Award. This work was also supported by MEI Pharma.

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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