Research ArticleCancer

Targeting therapy-resistant prostate cancer via a direct inhibitor of the human heat shock transcription factor 1

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Science Translational Medicine  16 Dec 2020:
Vol. 12, Issue 574, eabb5647
DOI: 10.1126/scitranslmed.abb5647

Putting the heat on cancer

Heat shock factor 1 is a transcription factor that helps protect healthy cells from cellular stress but can also protect tumor cells and interfere with anticancer treatment. In particular, the amount of heat shock factor 1 in the nucleus is a poor prognostic factor for patients, including those with neuroendocrine prostate cancer, an aggressive late-stage form of the disease. To target this protein, Dong et al. designed a selective small-molecule inhibitor, which directly binds nuclear heat shock factor 1 and promotes its degradation. The inhibitor was effective in multiple models of prostate cancer, including a neuroendocrine model indicating its potential for clinical testing.

Abstract

Heat shock factor 1 (HSF1) is a cellular stress-protective transcription factor exploited by a wide range of cancers to drive proliferation, survival, invasion, and metastasis. Nuclear HSF1 abundance is a prognostic indicator for cancer severity, therapy resistance, and shortened patient survival. The HSF1 gene was amplified, and nuclear HSF1 abundance was markedly increased in prostate cancers and particularly in neuroendocrine prostate cancer (NEPC), for which there are no available treatment options. Despite genetic validation of HSF1 as a therapeutic target in a range of cancers, a direct and selective small-molecule HSF1 inhibitor has not been validated or developed for use in the clinic. We described the identification of a direct HSF1 inhibitor, Direct Targeted HSF1 InhiBitor (DTHIB), which physically engages HSF1 and selectively stimulates degradation of nuclear HSF1. DTHIB robustly inhibited the HSF1 cancer gene signature and prostate cancer cell proliferation. In addition, it potently attenuated tumor progression in four therapy-resistant prostate cancer animal models, including an NEPC model, where it caused profound tumor regression. This study reports the identification and validation of a direct HSF1 inhibitor and provides a path for the development of a small-molecule HSF1-targeted therapy for prostate cancers and other therapy-resistant cancers.

INTRODUCTION

Cancer cells are chronically exposed to stressful conditions as they proliferate, invade, metastasize, and colonize distinct tissues and compartments (1). The malignant phenotype is associated with numerous stresses including the need to cope with increased demand for protein biosynthesis, the accumulation of mutated proteins, and the need to support conformationally labile oncogenic signaling proteins such as receptors, protein kinases, and transcription factors (1, 2).

In human cancers, heat shock factor 1 (HSF1) is a central stress-protective transcription factor driving a distinct oncogenic transcription program known as the HSF1 cancer gene signature (HSF1 CaSig), in which HSF1 activates expression of genes encoding protein-folding chaperones, DNA replication and repair factors, cell cycle drivers, metabolic enzymes, and prosurvival proteins, while repressing genes that promote immune surveillance, inflammation, cell adhesion, and cell death (15). In addition, HSF1 plays critical supportive roles in stroma and in the tumor microenvironment (13). Although HSF1 is dispensable for overall growth and development in mammals, pioneering studies demonstrated that a wide range of cancers exhibit a “non-oncogene addiction” to HSF1 (511). Increased HSF1 expression and activity are associated with disease severity, therapy resistance, and shortened disease-free survival in patients with cancer (2, 7, 12).

In normal cells, HSF1 is found as an inactive monomer in the cytoplasm, but it oligomerizes and accumulates in the nucleus in response to acute stress. However, in the chronically proteotoxic and broadly stressful environment of cancer cells, HSF1 is highly expressed, constitutively activated, oligomerized, and localized to the nucleus via diverse mechanisms (3, 810, 13). The constitutive activation of HSF1 in cancer and its prognostic value have prompted genetic experiments to validate the role of HSF1 in the initiation and progression of cancer (1, 2, 6). HSF1 knockout mice are protected from cancer driven by mutations in RAS, inactivation of the P53 tumor suppressor, and HER2-induced breast cancer (6, 1416). Genetic HSF1 knockout or knockdown studies have shown strong efficacy in a broad spectrum of animal cancer models (2, 9, 10, 15).

Prostate cancer (PCa) development and progression require androgen receptor (AR) signaling, which activates the expression of genes that underlie PCa biology (17, 18). AR is highly dependent on the HSF1-activated multichaperone complex composed of heat shock proteins 90, 70, and 40 (HSP90, HSP70, and HSP40) and other proteins for AR stability, ligand binding, nuclear translocation, dimerization, and target gene DNA binding (17, 19). Although AR antagonists and androgen deprivation are well-established therapies for PCa, virtually all patients eventually become therapy resistant and progress to castration-resistant prostate cancer (CRPC) or neuroendocrine prostate cancer (NEPC) (18). The traditional AR-targeted therapeutic approach has marked limitations due to multiple mechanisms of drug resistance including AR gene amplification (17), the emergence of drug-resistant AR mutations (18), the expression of constitutively active AR splice variants that lack the ligand-binding domain (LBD) (19, 20), and the development of AR-negative NEPC (21, 22). Hence, alternative, AR-independent therapies are urgently needed for patients with therapy-resistant PCa.

Despite compelling genetic data demonstrating the strict in vivo requirement for HSF1 in a broad spectrum of cancers, a direct and selective HSF1 inhibitor has not been comprehensively validated. Although “HSF1 pathway” inhibitors have been identified and evaluated in cellular and mouse xenograft cancer models, they either act in an indirect manner in the HSF1 pathway or have an unknown mechanism of action (2, 11). Here, we describe the identification and characterization of a Direct Targeted HSF1 InhiBitor (DTHIB), by screening for physical interactions between the structurally well-ordered HSF1 DNA binding domain (DBD) and small molecules. DTHIB selectively engaged with HSF1 and inhibited core HSF1 functions by accelerating the rate of nuclear HSF1 degradation. By exploiting the dependence of AR and AR splice variant 7 (AR-v7) on the multichaperone machine, DTHIB suppressed AR/AR-v7 signaling pathways. Moreover, DTHIB also acted independently of AR and potently attenuated PCa progression in mouse models, including highly aggressive NEPC. Together, these results described a direct HSF1 inhibitor with strong developmental potential for therapy-resistant PCa and other cancers.

RESULTS

Differential scanning fluorimetry identifies direct HSF1 inhibitors

Although genetic approaches have underscored the requirement for HSF1 in a variety of cancer models, current small-molecule HSF1 inhibitors suffer from a lack of evidence for a direct, high-affinity interaction with HSF1. Instead, most existing HSF1 inhibitors function indirectly through protein kinases, the general transcription/translation machinery, or via other proteins of unclear mechanistic relationship to HSF1 function (13, 11). To identify small molecules that directly target human HSF1, we established an in vitro screening platform based on differential scanning fluorimetry (DSF) (23, 24). DSF reveals direct protein-ligand interactions by monitoring changes in the melting temperature of the target protein as a function of ligand binding. Direct protein-ligand interactions could either increase or decrease the thermal stability of the target, leading to a change in the observed melting temperature (Tm) (23). We previously demonstrated that DSF serves as a robust assay to evaluate the binding of recombinant full-length HSF1 or the HSF1 DBD to its cognate DNA binding site, the heat shock element (HSE) (24).

Recombinant human 6xHis-tagged HSF1 DBD was expressed in Escherichia coli and purified via Ni-NTA (nitrilotriacetic acid) enrichment (Fig. 1A). The purified HSF1 DBD bound its DNA ligand HSE in vitro, as shown by DSF, indicating that it was properly folded and functional (Fig. 1B). A collection of structurally diverse small molecules was screened against the HSF1 DBD by DSF. To monitor the quality and interplate variability of HSF1 DBD–small molecule interactions, dimethyl sulfoxide (DMSO) and the HSE oligonucleotide were included as negative and positive controls, respectively. Whereas 98.87% of the ~5500 screened compounds were negative for HSF1 DBD binding, positive compounds either increased or decreased the thermal stability of the HSF1 DBD relative to DMSO (Fig. 1C). All positive compounds were subjected to secondary DSF validation screens to identify those molecules positive for HSF1 DBD binding but negative for binding to the heat shock cognate 70 kDa protein (HSC70) chaperone, as a preliminary measure of nonspecific interactions (fig. S1A).

Fig. 1 In vitro ligand binding screen identifies a direct small-molecule HSF1 inhibitor.

(A) Coomassie-stained SDS–polyacrylamide gel electrophoresis protein gel showing the purification of recombinant human HSF1 DBD. L, molecular weight ladder; WCE, whole-cell extract; FT, flow-through fraction; Wash, washing fractions; E, elution fraction. (B) DSF melting curves demonstrating the shift in thermal stability of the HSF1 DBD upon interaction with the HSE (n = 3, N = 1). RFU, relative fluorescence units. (C) A scatterplot summary of the primary DSF screen results. Each red dot represents the observed Tm of the HSF1 DBD in the presence of a screened compound. Blue line, Tm of HSF1 DBD with DMSO solvent; purple line, Tm of HSF1 DBD with HSE DNA ligand. (D) Chemical structure of compound DTHIB. (E) DTHIB physically binds to the HSF1 DBD and increased its melting temperature by 4°C (n = 3, N = 1). (F) SPR sensorgram of DTHIB binding to a surface coated with HSF1 DBD, with DTHIB concentrations and calculated Kd for binding shown (n = 3, N = 1). RU, response units. (G) HSF1 DBD fluorescence emission spectra obtained from the TFQ assay in response to increased concentrations of DTHIB (n = 3, N = 3). (H) Immunoblot showing that DTHIB dose-dependently inhibited heat shock–induced HSP25 and HSP70 expression in MEFs. GAPDH, glyceraldehyde-3-phosphate dehydrogenase. (I) qRT-PCR data showing impact of DTHIB on indicated transcript abundance in response to heat shock (n = 3, N = 3). ns, not significant.

Two structurally similar molecules identified in the screen, I06 and P06, selectively interacted with HSF1 DBD (fig. S1, B and C). Because these two compounds were not available from commercial sources at high purity, a structurally similar derivative, denoted DTHIB, was synthesized and purified in-house (Fig. 1D and fig. S1, D to F). As demonstrated with the DSF assay, DTHIB strongly interacted with the HSF1 DBD (Fig. 1E) and was further investigated to characterize its binding properties and potential impact on HSF1 function. On the basis of chemical structures, DTHIB, together with primary screen hits I06 and P06, represented a class of small molecules that has not been reported as interacting with HSF1 (2).

To further assess the specificity of DTHIB, DSF assays were performed using DTHIB against the purified recombinant replication protein A 1 (RPA1) single-stranded DBD (ssDBD) or the HSC70 chaperone. DTHIB exhibited no sign of nonspecific interaction with either protein (fig. S2A). A structurally related molecule, A01, was synthesized, which contained the 1,3-diphenylurea scaffold. A01 did not interact with the HSF1 DBD and was included as a negative control for downstream analyses (fig. S2, B to D). Surface plasmon resonance (SPR) was used to independently and quantitatively assess DTHIB binding to the HSF1 DBD, demonstrating a dissociation constant (Kd) for DTHIB binding to the HSF1 DBD of 160 nM (Fig. 1F). As elucidated in the crystal structure (25), the HSF1 DBD harbors two solvent-exposed tryptophan residues in proximity that could enable the detection of HSF1 DBD interactions based on intrinsic tryptophan fluorescence quenching (TFQ), an approach commonly used to monitor ligand binding (26). In the presence of solvent alone, the HSF1 DBD emitted an intrinsic fluorescence peak at λ = 330 nM after excitation at 280 nm. Titration of DTHIB into the in vitro reaction demonstrated dose-dependent quenching of the intrinsic tryptophan fluorescence of the HSF1 DBD (Fig. 1G), whereas DTHIB had no effect on tryptophan fluorescence for RPA1 ssDBD or HSC70 (fig. S2E). Moreover, consistent with DTHIB binding to the HSF1 DBD within the context of the full-length protein, incubation of DTHIB with full-length recombinant HSF1 conferred an altered limited proteolytic pattern for HSF1, detected by silver staining (fig. S3, A and B). Several polypeptide species with increased abundance upon addition of DTHIB were excised and analyzed by mass spectrometry (fig. S3B and data file S1). Among the altered species, a 15- to 20-kDa band was found to encompass the HSF1 DBD (amino acid residues 63 to 117) (fig. S3, C and D).

Unlike cell-based screens, the DSF screen is agnostic to in vivo activity. To ascertain the potential impact of DTHIB on HSF1, DTHIB was first evaluated in mouse embryonic fibroblasts (MEFs) under acute heat shock conditions. In MEFs, DTHIB attenuated the robust acute heat shock induction of the HSP70 and HSP25 molecular chaperones in a dose-dependent manner (Fig. 1H). Moreover, quantitative reverse transcription polymerase chain reaction (qRT-PCR) results demonstrated that DTHIB attenuated the heat shock response by reducing the steady-state transcript abundance of multiple molecular chaperones (Fig. 1I). The structurally related derivative A01, which did not bind the HSF1 DBD, had no detectable impact on heat shock induction of the HSP70 or HSP25 proteins (fig. S4A).

DTHIB is a direct HSF1 inhibitor and targets the HSF1 CaSig

A wide variety of cancer cell lines and tissues exhibit chronic HSF1 activation and strongly elevated abundance of protein chaperones and other HSF1-activated genes (15, 12). To investigate the impact of DTHIB in cells with constitutive HSF1 activity, the human CRPC cell line C4-2 was used (27). In C4-2 cells, DTHIB reduced steady-state protein abundance of the molecular chaperones P23, HSP27, HSP70, and HSP90—all bona fide HSF1 targets (Fig. 2A and quantitated in fig. S4B) (35). Moreover, analysis of transcript abundance demonstrated that DTHIB repressed the expression of genes that are directly bound and activated by HSF1, and derepressed expression of genes directly bound and repressed by HSF1 (Fig. 2B) (35). This latter observation suggested that DTHIB did not nonspecifically inhibit the transcription machinery at the concentrations used, because cells mounted a positive transcriptional response for specific genes. In the AR-negative human PCa cell line PC-3 (28), DTHIB also inhibited HSF1 (fig. S4C), suggesting that the activity of DTHIB was independent of AR status.

Fig. 2 HSF1 is highly expressed in human prostate cancer and is inhibited by DTHIB.

(A) DTHIB dose-dependently inhibited expression of molecular chaperones in C4-2 PCa cells after 48-hour treatment with the indicated concentrations of DTHIB. (B) Changes in HSF1 target gene transcript abundance after 48-hour DTHIB treatment with the indicated concentrations in C4-2 cells (n = 4, N = 4). Relative abundance of transcripts was first normalized internally to GAPDH and then compared to the DMSO control. (C) RNA-seq analysis of C4-2 cells after 48-hour treatment with 1 μM DTHIB or DMSO solvent (n = 4, N = 1). Shown is a heatmap for transcripts from direct HSF1 target genes grouped in functional categories. (D) HSF1 immunoblots and LI-COR quantitation (n = 4, N = 2) from CETSA with C4-2 lysates treated with DMSO or DTHIB at 10 μM. Shown are the temperatures used for lysate treatment. (E and F) Generation of scrambled shRNA (Scr) and shHSF1 cell lines from C4-2 and PC-3 cells. (E) For C4-2–derived cell lines, basal abundance of HSF1 and HSP70 was assessed by immunoblotting. (F) For PC-3–derived cell lines, both basal (37°C) and heat shock–induced HSF1 and HSP70 abundance were assessed by immunoblotting. (G) C4-2 Scr and shHSF1 cells exposed to 36-hour treatment with 2.5 μM DTHIB were evaluated for HSF1 target gene expression by qRT-PCR (n = 5, N = 2). HSPA4L is a target gene activated by HSF1, and STAT6 is a target gene repressed by HSF1. (H) Cell viability readouts of Scr and shHSF1 cell lines derived from C4-2 and PC-3 cells after 72-hour treatment with the indicated concentrations of DTHIB (n = 4, N = 1).

The impact of DTHIB administration in C4-2 cells was further validated by RNA sequencing (RNA-seq) experiments, cross-referenced to HSF1-ChIP (chromatin immunoprecipitation) sequencing databases, to investigate the impact of DTHIB on the broader landscape of the HSF1 CaSig (Fig. 2C, fig. S4D, and data file S2) (5, 29, 30). These studies demonstrated that DTHIB treatment broadly inhibited the HSF1 CaSig and HSF1-mediated transcriptional network (Fig. 2C). The observation that DTHIB administration derepressed multiple HSF1-repressed genes further supported the notion that DTHIB was not acting as an inhibitor of the core transcription machinery.

In addition to the direct binding of DTHIB to the HSF1 DBD demonstrated by multiple biochemical assays in vitro, a cellular thermal shift assay (CETSA) in C4-2 whole-cell lysate was performed. This experiment demonstrated that DTHIB increased the aggregation temperature (Tagg) of HSF1 by 5°C (Fig. 2D), validating the engagement of DTHIB with full-length HSF1 in cell extracts (31). In contrast, DTHIB elicited no impact on the Tagg of another nuclear transcription factor, SP1 (fig. S4E).

The activity of DTHIB is HSF1 dependent

Because the HSF1 inhibitor DTHIB was identified from an in vitro screen targeting recombinant human HSF1 DBD, it was critical to ascertain whether the activity of DTHIB in cell culture is HSF1 dependent. To address this question, scrambled short hairpin RNA (shRNA) (Scr) control and stable HSF1 knockdown (shHSF1) cell lines were generated from C4-2 and PC-3 cells, representing AR-positive and AR-negative PCa models, respectively (Fig. 2, E and F). The genetic depletion of HSF1 led to loss of HSP70 basal expression in C4-2 cells and diminished the heat shock response in PC-3 cells (Fig. 2, E and F).

The analysis of HSF1 target gene expression via qRT-PCR demonstrated that, in contrast to the C4-2 Scr line, the C4-2 shHSF1 cell line was not responsive to DTHIB for both the HSF1-activated gene HSPA4L and HSF1-repressed gene STAT6 (Fig. 2G). A similar result was observed in the isogenic pair of Scr and shHSF1 PC-3 cell lines when the HSF1 targets HSPA4L and HSP90AB1 were evaluated (fig. S4F). In cell viability assays, both the C4-2 and PC-3 shHSF1 cell lines showed reduced viability compared to control cells that was not further inhibited by DTHIB treatment (Fig. 2H). To further evaluate the dependency of DTHIB action on HSF1 in a distinct background, we used primary MEFs isolated from Hsf1+/+, Nf1−/− or Hsf1−/−, Nf1−/− mice. The neurofibromatosis type 1 (NF1) tumor suppressor protein represses HSF1 activity, and loss of NF1 leads to HSF1 activation and overexpression of several HSF1 target genes (8). In comparison to MEFs, Hsf1+/+, Nf1−/− MEFs, the Hsf1−/−, Nf1−/− double-knockout MEFs were resistant to DTHIB (fig. S4G).

Mechanism of action of HSF1 inhibition by DTHIB

DTHIB is a direct HSF1 inhibitor that physically engages with the HSF1 DBD with nanomolar affinity in vitro. In cell culture, DTHIB inhibited both the acute heat shock response and the activation or repression of HSF1 CaSig targets.

To evaluate whether DTHIB directly affects HSF1 DNA binding in vitro, a fluorescence anisotropy (FA) assay was used to quantitate the DNA binding affinity of recombinant HSF1 DBD or full-length HSF1 homotrimeric protein (24). As shown by FA, addition of DTHIB had no detectable impact on the in vitro DNA binding affinity of either HSF1 DBD or full-length HSF1 homotrimer (Fig. 3A and fig. S5, A and B). To determine whether DTHIB affected HSF1 DNA binding in cells, ChIP–quantitative PCR was used to measure HSF1 occupancy at both activated and repressed loci in the absence or presence of DTHIB. Unexpectedly, DTHIB reduced promoter occupancy at both HSF1-activated and HSF1-repressed target genes (Fig. 3B).

Fig. 3 DTHIB mechanism of action for HSF1 inhibition.

(A) FA titration curves showing the impact of DTHIB on the binding of full-length oligomeric HSF1 to a 6-FAM–conjugated HSE oligonucleotide in vitro (n = 3, N = 1). Shown are millipolarization units (y axis) and the concentration of HSF1 (nanomolars) on the x axis. (B) ChIP–quantitative PCR data from experiments quantitatively assessing promoter-HSF1 occupancy in C4-2 cells after 48-hour DTHIB treatment at the indicated concentrations (n = 3, N = 3). HSPA4L and CKS2 are HSF1-activated genes, whereas SPTN1 is an HSF1-repressed gene. The relative promoter-HSF1 occupancy for each gene was normalized to the DMSO control. (C) Immunoblots and LI-COR quantitation from subcellular fractionation experiments to assess cytosolic and nuclear HSF1 abundance from C4-2 cells treated with DTHIB for 48 hours at the indicated concentrations (n = 3, N = 3). (D) Immunoblots and LI-COR quantitation from cycloheximide (CHX) pause-chase and subcellular fractionation experiments with C4-2 cells to assess cytosolic and nuclear HSF1 half-life in response to 2.5 μM DTHIB (n = 3, N = 3). (E) C4-2 cells were treated without and with proteasome inhibitor MG132, in the absence or presence of the indicated concentration of DTHIB for 48 hours, and nuclear and cytosolic fractions were isolated. Proteins of interest were analyzed by immunoblotting and quantitated with LI-COR (n = 3, N = 3). −VE, negative control. (F) After 48-hour treatment with 5 μM DTHIB, Scr and shFBXW7 C4-2 cell lines were analyzed for DTHIB-induced nuclear HSF1 degradation after subcellular fractionation. Proteins of interest were analyzed by immunoblotting and quantitated with LI-COR (n = 3, N = 3).

To understand the seemingly disparate in vitro and in vivo HSF1 DNA binding results, we performed subcellular fractionation experiments to separate and quantitate the abundance of cytosolic and nuclear HSF1. DTHIB reduced nuclear HSF1 steady-state protein abundance in a dose-dependent manner, whereas cytosolic HSF1 was unaffected (Fig. 3C). Similar results were observed in AR-negative PC-3 cells (fig. S5C). To evaluate whether the reduction in nuclear HSF1 abundance was due to a change in HSF1 stability, nuclear HSF1 abundance in C4-2 cells was quantitated over time after treatment with cycloheximide to arrest new protein synthesis, in the absence or presence of DTHIB. Although there was no change in nuclear HSF1 stability in solvent-treated cells over 12 hours, DTHIB destabilized nuclear HSF1, resulting in a shortened HSF1 half-life of ~5 hours (Fig. 3D). In contrast, the stability of another nuclear transcription factor, SP1, was not affected by DTHIB (fig. S5D). The inactive analog A01 had no impact on either cytosolic or nuclear HSF1 stability (fig. S5E).

For both cancer and neurodegenerative diseases, nuclear HSF1 degradation is proteasome dependent and mediated by the FBXW7 (F-Box and WD repeat domain containing 7), the F-box component of SCF (Skp1, Cullin, and F-box proteins) E3 ligase complex, or the NEDD4 (Neural precursor cell expressed, developmentally down-regulated 4) E3 ligase (3). To ascertain whether DTHIB-induced degradation of nuclear HSF1 was proteasome dependent, C4-2 cells were treated with the proteasome inhibitor MG132 (N-carbobenzyloxy-l-leucyl-l-leucyl-l-leucinal) and the abundance of cytosolic and nuclear HSF1 assessed in the absence or presence of DTHIB. Although cytosolic HSF1 abundance was unchanged between solvent and DTHIB treatments, the loss of nuclear HSF1 in response to DTHIB was not observed when the proteasome was inhibited (Fig. 3E). In addition, cotreatment with DTHIB and MG132 led to the accumulation of polyubiquitinated HSF1 species, in comparison to the MG132-alone condition (fig. S5F). C4-2 cells with stable FBXW7 or NEDD4 knockdown were generated using shRNA (fig. S5G). When FBXW7 expression was knocked down, DTHIB was unable to accelerate nuclear HSF1 degradation (Fig. 3F), whereas NEDD4 knockdown had no impact on DTHIB activity (fig. S5H). FBXW7 has a number of physiological targets including C-MYC (9). To investigate whether DTHIB perturbs the normal function of FBXW7, we evaluated the steady-state abundance of nuclear C-MYC and observed no impact of DTHIB on nuclear C-MYC abundance (fig. S5I). Together, these studies demonstrated that DTHIB inhibited HSF1 by preferentially stimulating nuclear HSF1 degradation via a proteasome- and FBXW7-dependent pathway while sparing other targets of this degradation pathway. The loss of nuclear HSF1 led to depletion of promoter-bound HSF1 and inhibition of the HSF1 CaSig.

Targeting HSF1 in therapy-resistant prostate cancer

Since the initial discovery of the critical roles that HSF1 plays in supporting the malignant phenotype, this role has been validated in many cellular and animal cancer models. Accordingly, the high abundance of HSF1 in cancer cells and tissues and particularly high nuclear HSF1 abundance strongly correlate with poor clinical prognosis in a broad spectrum of human cancers (2, 7, 12). In PCa, elevated nuclear HSF1 abundance associated with higher Gleason grades and metastatic lymph node status, suggesting HSF1 as a critical factor for prostate cancer progression and metastasis (12).

Analysis of the National Cancer Institute The Cancer Genome Atlas database and published work (2, 13) demonstrated that the HSF1 gene was frequently amplified across a broad spectrum of human cancers. In PCa, including CRPC and NEPC, the HSF1 gene was highly amplified, and the abundance of nuclear HSF1 was associated with advanced disease, poor survival, and therapy resistance (Fig. 4A) (5, 7, 9, 12). Examination of a panel of human PCa cell lines demonstrated that HSF1 protein abundance was greatly increased in malignant cells, compared to the benign prostate cell line BPH-1 or RWPE-1 (Fig. 4B and fig. S6A). To independently ascertain the clinical relevance of high amounts of HSF1 in PCa, total HSF1 abundance was evaluated by immunohistochemistry staining in PCa patient tumor microarrays (TMAs). In comparison to benign samples (n = 9), prostate adenocarcinoma (n = 11), CRPC (n = 23), and NEPC (n = 21) samples all showed elevated HSF1 abundance. The most aggressive, AR-negative NEPC samples had the most profound elevation in HSF1 abundance compared to the other three groups, manifesting as strong nuclear staining (Fig. 4C and data file S3).

Fig. 4 Targeting HSF1 inhibits AR signaling.

(A) Genetic alteration frequencies of the HSF1 gene in prostate cancer (www.cbioportal.org/), amplified across a collection of independent prostate cancer studies (left) and in the three major clinical types of prostate cancer, in aggregate (right). TCGA, The Cancer Genome Atlas; MPC, metastatic prostate cancer. (B) Immunoblots of HSF1 in the malignant prostate cancer cell lines VCap, LNCap, C4-2, and PC-3, as compared to the benign prostate cell line BPH-1. (C) Representative images from HSF1 immunohistochemistry stains of human prostate cancer specimens from TMA, with scale bar equal to 20 μM and HSF1 quantitation shown below (N = 1). (D) Immunoblot showing the effect of 48-hour DTHIB treatment on the steady-state protein abundance of AR and PSA in C4-2 cells. (E) Effects of stable knockdown of HSF1 in C4-2 cells on HSF1 and AR activity, as ascertained for the HSF1 target gene HSPA4L, AR, and AR target genes (KLK3, FKBP5, and TMPRSS2), as compared to Scr control using qRT-PCR (n = 3, N = 3). (F) Forty-eight hours of DTHIB treatment of 22Rv1 cells dose-dependently inhibited expression of molecular chaperones HSP40 and HSP70 and led to reduction in full-length AR (AR-FL) and AR-v7 protein abundance and diminished PSA expression. (G) In comparison to enzalutamide (ENZA), 72-hour DTHIB treatment of 22Rv1 cells inhibited PSA expression and destabilized both full-length AR and AR-v7 protein, ascertained by immunoblotting. (H) Cell viability assay of 22Rv1 cells treated for 96 hours with enzalutamide or DTHIB at the indicated concentrations. Shown is the calculated median effective concentration (EC50) value for DTHIB. N/A indicates that a reliable EC50 value could not be estimated (n = 5, N = 1).

HSF1 inhibition suppresses AR activity

Among the many roles that HSF1 target genes play in cancer, the critical functions of chaperones in folding, maturing, stabilizing, and sustaining the activity of key signaling proteins such as nuclear hormone receptors are clear (17). This is particularly evident in hormonally driven cancers such as PCa, in which the AR plays a pivotal role in the regulation of genes that drive prostate cancer survival, proliferation, and progression (17, 19). Because DTHIB potently inhibited HSF1 and coordinately diminished abundance of the multichaperone machinery essential for AR stability and activity (Fig. 2A), we evaluated the impact of DTHIB on AR function. DTHIB treatment of C4-2 PCa cells depleted AR protein and potently dampened expression of the AR target gene KLK3, encoding prostate-specific antigen (PSA), a key serum biomarker for prostate cancer progression (Fig. 4D and quantitated in fig. S6B). As a genetic validation, in comparison to shScr cells, shHSF1 C4-2 cells showed strong depletion of HSF1 protein and diminished expression of PSA and other AR targets (Fig. 4E and fig. S6C).

After initial response to AR antagonists, patients with PCa eventually develop therapy resistance through several mechanisms including AR gene amplification, AR drug-resistant mutations, and the generation of AR splice variants (18). In particular, the constitutively active AR-v7 lacks both the LBD and the HSP90-binding motif, rendering AR-v7 resistant to AR antagonists such as enzalutamide, and independent of HSP90 function (20). However, the stability and activity of AR-v7 remain heavily reliant on other chaperones, such as HSP40 and HSP70 (19). Human PCa cell line 22Rv1, expressing both full-length AR and abundant AR-v7, was used to evaluate the efficacy of DTHIB. In 22Rv1 cells, DTHIB inhibited HSF1 and led to the coordinated depletion of molecular chaperones including HSP40 and HSP70. Simultaneously, the steady-state amounts of both full-length AR and AR-v7 were reduced in a dose-dependent manner, with a concomitant loss of PSA expression (Fig. 4F and quantitated in fig. S6D). Whereas the clinically used AR antagonist enzalutamide (17, 18) was ineffective against AR-v7, DTHIB inhibited AR signaling, PSA expression, and 22Rv1 cell growth (Fig. 4, G and H). Similarly, in C4-2 cells, DTHIB exhibited superior efficacy in the inhibition of AR signaling and cell growth, as compared to enzalutamide (fig. S6, E and F). These results demonstrated that DTHIB coordinately reduced expression of the multichaperone complex and attenuated AR and AR-v7 signaling in 22Rv1 cells, a human AR antagonist-resistant PCa cell line.

DTHIB inhibits therapy-resistant prostate cancer cell proliferation

Consistent with the functional importance of HSF1 in cancer (1, 2, 4, 11), pharmacological HSF1 inhibition via DTHIB decreased the viability of an array of PCa cells encompassing both the AR-positive cell lines LNCap, VCap, C4-2, TRAMP-C2, and 22Rv1 and the AR-negative cell line PC-3, as compared to the benign prostate cell line BPH-1 (Fig. 5A and fig. S7A) (27). The efficacy of DTHIB in C4-2 cell growth inhibition was further validated by trypan blue exclusion assays and automated viable cell counting (Fig. 5B). In clonogenic assays, DTHIB dose-dependently reduced the clonal expansion of both C4-2 and PC-3 PCa cells (Fig. 5C and fig. S7B). Moreover, the inactive analog A01 had no detectable impact on C4-2 cell viability (fig. S8A). In agreement with the broad dependency of cancers on HSF1 (1, 4), the efficacy of DTHIB was not limited to prostate cancers. Experiments demonstrated that DTHIB reduced cell viability in human breast cancer and melanoma cells and inhibited the HSF1 CaSig in the melanoma cell line SK-MEL-5 (fig. S8, B and C).

Fig. 5 DTHIB inhibits human prostate cancer cell proliferation.

(A) AlamarBlue assays from 96-hour DTHIB treatment assessed the viability of the malignant prostate cancer cell lines C4-2 and PC-3, as compared to the benign prostate cell line BPH-1 (n = 5, N = 1). Shown is the calculated EC50 for DTHIB in each cell line. (B) Validation of DTHIB efficacy in the C4-2 PCa cell model using the trypan blue exclusion assay (n = 3, N = 3). Shown is the EC50 for DTHIB in C4-2 cells calculated with this assay. (C) Clonogenic assay of C4-2 cells treated with the indicated concentrations of DTHIB for 10 days. A representative image of C4-2 colonies is shown. Shown is the EC50 for DTHIB in C4-2 cells calculated with the clonogenic assay (n = 3, N = 3). (D) Fluorescence-activated cell sorting analysis showing the effects of 48-hour 5 μM DTHIB treatment of C4-2 cells on cell cycle arrest and accumulation in the G1 phase. (E) DTHIB treatment drives C4-2 cells into senescence, indicated by the induction in SA-β-gal activity (n = 3, N = 1). (F) C4-2 cells were treated with 5 μM venetoclax, 5 μM DTHIB, or 5 μM each of venetoclax and DTHIB for 48 hours. Apoptosis in each sample was quantitated by measuring caspase 3/7 enzymatic activity (n = 3, N = 1). (G) C4-2 cells were treated as described in (F), and the percentage of dead cells was quantitated using the trypan blue exclusion assay (n = 3, N = 1). (H) C4-2 cells were treated as described in (F). Immunoblotting was used to detect changes in key molecular markers involved in the cooperation between DTHIB and venetoclax.

Acute genetic knockdown of HSF1 in a variety of cancer cells resulted in cell cycle arrest and cellular senescence, a condition in which cells retain viability without further proliferation (14, 32, 33). As analyzed by fluorescence-activated cell sorting, DTHIB treatment of C4-2 cells induced cell cycle arrest, with accumulation in the G1 phase (Fig. 5D). Moreover, DTHIB treatment increased senescence-associated β-galactosidase (SA-β-gal) activity, indicating that DTHIB stimulated C4-2 PCa cell entry into senescence (Fig. 5E). When DTHIB was removed from C4-2 cell culture medium after an initial 48-hour exposure, cell growth did not resume, and a robust induction of SA-β-gal activity occurred during the 72-hour washout period (Fig. 5E and fig. S9A). Together, these results demonstrated that DTHIB triggered PCa cell senescence and are consistent with the observed reprogramming of other cancer cell types into a senescence program after acute genetic HSF1 knockdown (15, 33).

Although senescent cells are arrested in the cell cycle and do not proliferate, they remain metabolically active (32). To evaluate downstream markers of HSF1 inhibition for the duration of the DTHIB response, the abundance of the HSP70, AR, and PSA proteins and their recovery rate after DTHIB removal were determined in C4-2 cells. After the initial 48-hour treatment, DTHIB resulted in strong depletion of HSP70, AR, and PSA (fig. S9, B and C). Treatment with venetoclax alone stimulated a low level of caspase 3/7 activity, an apoptotic marker, whereas 5 μM DTHIB only mildly perturbed the caspase 3/7 activity and was not statistically significant (Fig. 5F). Cotreatment with DTHIB and venetoclax strongly induced caspase 3/7 activity in a synergistic fashion (Fig. 5F and fig. S9D). This observation was corroborated using the trypan blue cell death assay (Fig. 5G). BAG3 is an HSF1-activated gene, and the BAG3 protein critically stabilizes the antiapoptotic protein BCL-2, thereby keeping apoptosis in check (34). Exposure of C4-2 cells to DTHIB reduced the expression of BAG3, HSP70, and HSP27, with a parallel reduction in BCL-2 protein abundance. Furthermore, cotreatment with DTHIB and venetoclax stimulated robust activation of caspase 3, as indicated by the enhanced cleavage of pro-caspase 3 (Fig. 5H).

Targeting HSF1 in animal models of therapy-resistant prostate cancer

The critical nature of HSF1 has been demonstrated in a variety of animal cancer models by genetic knockdown (2, 6, 15). However, no direct HSF1 inhibitor has been used to establish pharmacological proof of concept for HSF1 as a druggable target in animals. To test this, we evaluated a C4-2 xenograft mouse model after daily intraperitoneal administration of DTHIB at 5 mg/kg, as compared to vehicle alone. DTHIB attenuated tumor progression, with no visible tumor growth over a 3-week period and a 40% reduction in median tumor volume (Fig. 6A and data file S4). Longitudinal tumor measurements in situ were confirmed by postmortem quantitation of tumor weights (Fig. 6, A to C, and data file S4). During the 3-week course, DTHIB administration elicited no observable adverse effects on mouse behavior, weight, or organ histopathology (Fig. 6D and fig. S10A). At the experimental endpoint, circulating PSA concentrations, secreted from C4-2 tumor cells, showed a significant decrease (P < 0.01) in the DTHIB cohort, compared to vehicle-treated animals (Fig. 6E). Furthermore, qRT-PCR analysis of mRNA from the six largest tumors isolated from each cohort demonstrated that DTHIB inhibited the HSF1 CaSig in the tumor tissue (Fig. 6F), mirroring the impact of DTHIB in cultured C4-2 cells (Fig. 2C). DTHIB was also efficacious against the AR-negative PC-3 cell PCa model (fig. S10, B and C, and data file S4). This finding is consistent with the observation that the core activity of DTHIB was independent of AR (Fig. 5A and figs. S4C, S7B, and S8, B and C), likely as a result of perturbation of the critical HSF1 CaSig (5).

Fig. 6 Pharmacological inhibition of HSF1 attenuates prostate cancer progression in animal models.

(A) Median tumor volume plot over 21 days of C4-2 xenograft study in which vehicle (blue) or DTHIB (red) at 5 mg/kg were administered daily intraperitoneally to independent mouse cohorts (n = 12, N = 1). (B) Tumors from the C4-2 xenograft study (A) were excised from mice and weighed at the experimental endpoint (n = 12 per cohort). (C) Images of tumors from the C4-2 xenograft study in (A) harvested at the experimental endpoint. (D) Percent change in average body weight of mice from the C4-2 xenograft study in (A) (n = 12, N = 1). (E) Mouse serum samples from the C4-2 xenograft study in (A) obtained at the experimental endpoint were assayed for human PSA concentrations by enzyme-linked immunosorbent assay (ELISA) (n = 12, N = 1), with serum from nu/nu mice (no tumor implantation and no drug treatment) as negative control (n = 5, N = 1). (F) Pharmacodynamic analysis of transcript abundance corresponding to direct HSF1 target genes in C4-2 tumors (n = 6, N = 1) by qRT-PCR. (G) Median tumor volume plot from a 22Rv1 xenograft study in which vehicle, enzalutamide (15 mg/kg), or DTHIB (5 mg/kg) was administered daily intraperitoneally to three animal cohorts (n = 10, N = 1). (H) At the experimental endpoint, mouse serum samples (n = 10, N = 1) from 22Rv1 xenograft study in (G) were collected and assayed for human PSA concentrations by ELISA, using nu/nu mouse serum as blank (n = 3, N = 1). (I) In the TRAMP-C2 syngeneic mouse prostate cancer model, tumors from two cohorts of eight mice per cohort were allowed to grow to an average tumor volume of 100 mm3, and then mice were treated with vehicle or DTHIB at 5 mg/kg daily ip (n = 8, N = 1). Tumors from two other cohorts of four mice per cohort (late cohorts) were allowed to grow to an average tumor volume of ~200 mm3 and treated with DTHIB (5 or 1 mg/kg ip) daily (n = 4, N = 1). Arrow indicates the treatment start date of the “late” cohorts.

Human 22Rv1 cells are a widely used model to investigate AR pathway-targeted drug-resistant PCa, driven by the AR-v7 variant (27). The in vivo efficacy of DTHIB against AR-v7, as compared to the AR antagonist enzalutamide, was assessed in a 22Rv1 xenograft model. In agreement with a previous report (18) and our cell culture data (Fig. 4, G and H), 22Rv1-derived tumors were not responsive to enzalutamide in vivo. However, DTHIB administration at 5 mg/kg per day strongly attenuated the growth of 22Rv1 tumors (Fig. 6G and fig. S10D). Circulating PSA concentrations were evaluated at the experimental endpoint, where the DTHIB-treated cohort had significantly smaller tumors (P < 0.01) and lower serum PSA (P < 0.05) compared to either the vehicle- or enzalutamide-treated cohort (Fig. 6, G and H, fig. S10D, and data file S4). During the course of the experiment, DTHIB was well tolerated, and no weight loss was observed (fig. S10E and data file S4).

Although DTHIB imposed a strong suppression of tumor growth in nude mice, the host immune system can make profound contributions to tumor regression but cannot be evaluated in xenograft models (35, 36). To investigate whether pharmacological HSF1 inhibition shows improved efficacy in animals with a functional immune system, DTHIB was evaluated in a syngeneic mouse PCa model. This model of NEPC, which is highly refractory to current therapies, makes use of TRAMP-C2 cells implanted in syngeneic C57BL/6 mice (22, 27, 37). Implanted tumors were allowed to grow to ~100 mm3, after which either vehicle or DTHIB was administered at 5 mg/kg per day intraperitoneally (ip). In two additional cohorts, tumors were allowed to grow to ~200 mm3, and then mice received DTHIB (either 1 or 5 mg/kg per day). In cohorts administered with DTHIB (5 mg/kg), robust tumor regression was observed after initiating treatment (Fig. 6I, fig. S10F, and data file S4). Moreover, administration of DTHIB at 1 mg/kg per day also resulted in significant tumor regression (P < 0.001), demonstrating a dose-dependent activity for the HSF1 inhibitor in animals (Fig. 6I). In the syngeneic mouse PCa model, DTHIB was well tolerated, and neither weight loss nor organ damage was observed (fig. S10, G and H).

DISCUSSION

It was predicted over a decade ago that HSF1 inhibitors would soon be identified and in early preclinical development for use in oncology (6). Although the use of cell-based reporter screens has identified chemical probes to interrogate aspects of the heat shock response (2, 11), this approach has not yielded direct HSF1 inhibitors with a known mechanism of action. In contrast, the DSF screen detailed here provided a well-defined system to interrogate the human HSF1 DBD, which identified direct protein-ligand interactions. This approach was predicated on recent structural studies of HSF DBDs, which determined that the surface of the HSF1 DBD is highly solvent exposed (25, 38, 39). The HSF1 DBD, one of the few structured regions of mammalian HSF1, embodies the critical DNA binding activity and serves as a critical hub for protein-protein interactions that regulate the function of HSF1 in vivo (2, 39, 40). Here, we demonstrated that DTHIB directly binds to HSF1 by DSF, SPR, TFQ, and partial proteolysis in a purified system in vitro. Moreover, the ability of DTHIB to inhibit the HSF1 CaSig and cell viability in multiple cancer cell lines was dependent on the presence of HSF1. Thus, DTHIB binds with high affinity to a distinct domain on HSF1, but the precise amino acid residues and side chains with which DTHIB interacts have yet to be structurally elucidated.

DTHIB prevented tumor growth in multiple therapy-resistant PCa models and inhibited the HSF1 CaSig in human tumor cells implanted in mice. This indicates that DTHIB was distributed to tumor tissue and engaged HSF1 in vivo, consistent with DTHIB binding to full-length HSF1 in PCa cell extracts as shown by CETSA analysis. Furthermore, DTHIB was well tolerated in animals at doses of as high as 25 mg/kg per day. No overt weight loss, behavioral abnormalities, or histopathologic changes in multiple organs evaluated were observed in DTHIB-treated mice. Considering that Hsf1−/− mice have been generated in distinct genetic backgrounds, the tolerance to DTHIB may also, in part, reflect the nonessentiality of HSF1 in normal tissue (41, 42). Moreover, the derepression of HSF1-repressed genes by DTHIB indicated that DTHIB-treated cells were viable and that, in contrast to other HSF1 inhibitors, DTHIB did not inhibit the general transcription or translation machineries (2, 5). Given the time required for resynthesis and accumulation of downstream targets of HSF1, and their impact on AR abundance and activity, the degradation of nuclear HSF1 by DTHIB may result in prolonged therapeutic activity.

DTHIB selectively accelerated the degradation of nuclear HSF1 while sparing the inactive cytosolic fraction of the protein. This mechanism of action required the proteasome and depended on the E3 ligase F-box component FBXW7. Dysregulation of FBXW7 in cancer and Huntington’s disease greatly affects nuclear HSF1 degradation and HSF1 activity (9, 43). In both scenarios, HSF1 degradation requires FBXW7 binding to the same phosphodegron region in the HSF1 regulatory domain (3). Because DTHIB binds the HSF1 DBD, it is possible that DTHIB functions to directly recruit FBXW7 to the HSF1 DBD, and recent work suggests that FBXW7 can also interact with HSF1 through the C-terminal region of the DBD (44). Alternatively, DTHIB binding to the HSF1 DBD could alter the structure of full-length HSF1, enhancing the recruitment of FBXW7 to the phosphodegron. Further structural and biochemical experiments will be required to elucidate these molecular mechanisms.

The precise mechanism by which DTHIB drives nuclear compartment–specific HSF1 degradation remains to be elucidated. There are three splicing isoforms derived from the single FBXW7 gene, among which the FBXW7α isoform resides in the nucleus and is responsible for nuclear HSF1 degradation in melanoma (9, 45). Perhaps DTHIB preferentially recruited the FBXW7α isoform to nuclear HSF1 pools, thereby selectively driving nuclear HSF1 degradation. Furthermore, the inactive cytosolic HSF1 is bound and regulated by multichaperone complexes (3). HSF1 also binds and regulates the activity of adenosine 5′-monophosphate–activated protein kinase independent of its transcriptional function (46). The extensive protein-protein interactions between cytosolic HSF1 and other proteins, including molecular chaperones, could preclude DTHIB and/or cytosolic isoforms of FBXW7 from engaging cytosolic pools of HSF1. The large regions of HSF1 predicted to be intrinsically disordered (3, 47) also pose a challenge to structural biology approaches. As a result, little structural information is available for full-length HSF1, the HSF1-FBXW7 complex, or the HSF1-DTHIB complex. These molecular details will be important for a more thorough understanding of DTHIB mechanism of action. In contrast to the findings from melanoma models (9), genetic ablation of FBXW7 in C4-2 PCa cells failed to increase nuclear HSF1 abundance. Given the greatly elevated nuclear HSF1 abundance in both PCa cells and patient tumor samples, it is possible that key molecular processes needed for nuclear HSF1 degradation, such as the phosphorylation of HSF1 phosphodegron or the recruitment of FBXW7 to HSF1, are compromised in PCa. Alternatively, other mechanisms controlling HSF1 stability, such as lysine acetylation, could also function in regulating HSF1 abundance in PCa (48, 49).

Given the dependence of the estrogen receptor, progesterone receptor, and glucocorticoid receptor (GR) on protein chaperones (50), pharmacological HSF1 inhibition is likely to be effective against other hormone receptor–driven cancers and therapy resistance mechanisms driven by GR (51). It is important to emphasize that, although the HSF1 inhibitor DTHIB severely compromised AR and AR-v7 signaling, the efficacy of DTHIB in PCa cell and animal models was not limited by the AR status. This key feature could enable DTHIB to circumvent the challenges imposed by drug-resistant AR LBD mutations, therapy-resistant AR variants such as AR-v7, and AR-negative NEPC (18, 28). The critical functions of HSF1 in driving human malignancy are multifaceted, including but not limited to protein folding and maintaining nuclear hormone receptor activity, supporting a large constellation of proteins involved in oncogenic signaling, metabolism, DNA replication and repair, and preventing apoptosis. Thus, pharmacological HSF1 inhibition in PCa could have distinct advantages and a broader spectrum of action over traditional AR-targeted approaches by exploiting cancer cell addiction to HSF1 activity. Although the efficacy of DTHIB was demonstrated here in several animal xenograft PCa models, there was particular efficacy in immunocompetent mice, suggesting a potential role for tumor surveillance by the immune system or perhaps in vivo contributions to tumor regression via apoptosis. Pharmacodynamic markers and immune cell infiltration measurements early during DTHIB treatment could further clarify the mechanisms behind DTHIB efficacy. Additional validation of DTHIB in PCa models such as organoid cultures and patient-derived xenografts, which more closely resemble the complexity of human PCa, would also be of great value in validating HSF1 inhibitors for PCa (52, 53). The application of HSF1 inhibitors such as DTHIB could also extend beyond PCa as suggested by a cancer dependency map (DepMap) analysis, although the genetic and histological features of distinct cancers that affect responses to HSF1 inhibition remain to be elucidated (2). Apart from its potential for development as a therapeutic in oncology, a well-characterized, direct small-molecule inhibitor of HSF1 could serve as a useful tool to investigate mechanistic questions on the regulation and role of HSF1 in basic stress biology and in cancer.

MATERIALS AND METHODS

Study design

The objective of this study was to identify a direct small-molecule inhibitor of HSF1 and conduct validation, characterization, and animal model studies of the HSF1 inhibitor. For in vitro or cell culture experiments, at least three independent biological replicates per condition were used for statistical analysis. The precise number of biological replicates varied between different types of experiments because of technical limitations and was specified for each experiment. No outliers were excluded.

For animal experiments, power analyses were conducted on the basis of preliminary data, and selected cohort sizes used for all experiments were sufficient to have a power of 0.8 and P < 0.05. For all animal experiments, tumor-bearing animals were randomized before any treatment. No outliers were excluded unless explicitly mentioned, in which case the extreme outliers were identified and removed using 3xIQR as determining criteria. Detailed descriptions of experimental methods described in this manuscript are provided in Supplementary Materials and Methods.

Animal studies

For C4-2, PC-3, and 22Rv1 PCa xenograft experiments, nude (nu/nu) mice were acquired from the Duke University Department of Laboratory Animal Research Breeding Core at 6 weeks of age. Before any procedures, animals were acclimated for 1 week in the vivarium. Prostate cancer cells used in xenograft experiments passed the IMPACT III rodent pathogen test at IDEXX BioAnalytics (www.idexxbioanalytics.eu/impact). Cells were cultured in complete growth medium to 70% confluency, detached by trypsinization [Gibco Trypsin-EDTA (0.25%), catalog no. 25200056] at 37°C for 2 min, and trypsin was inactivated by adding prewarmed complete growth medium to the cell suspension. Cells were washed twice with phosphate-buffered saline (PBS) at room temperature and resuspended in ice-cold RPMI 1640 (no phenol red, Gibco 32404-014) medium. Before administration to nu/nu mice, the cell suspension was prechilled on ice at 4°C, mixed well with Matrigel (Corning 354234) at a 1:1 ratio, and maintained on ice. One hundred microliters of final cell suspension was injected subcutaneously into the right flank of each mouse using 26-G × 1.6 cm SubQ needles (BD 305115). A total of 1 × 106 cells were administered per animal.

For TRAMP-C2 syngeneic PCa model experiments, C57BL/6 mice were acquired from the Jackson laboratory at 6 weeks of age and allowed to acclimate in the vivarium for 2 weeks. Fur on the flank and belly was removed using electric clippers. TRAMP-C2 cells used in the syngeneic experiments passed the IMPACT III rodent pathogen test. TRAMP-C2 cells were cultured in complete growth medium to 70% confluency, detached using trypsinization as for xenograft experiments, washed twice with PBS, and resuspended in Dulbecco’s minimum essential medium–high glucose (HG) (no phenol red, Gibco 21063029) medium. The cell suspension was prechilled on ice at 4°C, mixed well with Matrigel (Corning 354234) at a 1:1 ratio, and maintained on ice. One hundred microliters of final cell suspension was administered subcutaneously into the right flank of each mouse using 26-G × 1.6 cm SubQ needles (BD 305115). A total of 1 × 106 cells were administered per animal.

For intraperitoneal administration of small molecules, DTHIB or enzalutamide was dissolved in DMSO. For DTHIB, polyethylene glycol (PEG) 400 was then added to DMSO in a 4:1 volume ratio. Then, 40% Captisol (RC-0C7-100) in saline was added to the PEG 400/DMSO mixture in a 3:1 volume ratio to achieve the final intraperitoneal formulation: 30% Captisol + 45% saline + 20% PEG 400 + 5% DMSO. One hundred microliters of the intraperitoneal solution was administered to animals intraperitoneally using 27-G × 1/2-in needles (BD 305109). For enzalutamide, PEG 400 was added to DMSO in a 4:1 volume ratio. Then, 40% Captisol in saline was added to the PEG 400/DMSO mixture in a 3:2 volume ratio to achieve the final intraperitoneal formulation of 24% Captisol + 36% saline + 32% PEG 400 + 8% DMSO. One hundred microliters of the enzalutamide solution was administered to animals intraperitoneally using 27-G × 1/2-in needles (BD 305109). Tumors were measured using a digital caliper (Thermo Fisher Scientific 06-664-16) and tumor volumes calculated using the formula V = ((W^2) × L)/2. Animals’ general health was noted daily. Tumor and body weight measurements were conducted every Monday, Wednesday, and Friday.

Statistical analysis

Unless specifically indicated, statistical analyses of data described in this manuscript were conducted using the unpaired Student’s t test assuming equal variance for comparison between two groups and standard one-way analysis of variance (ANOVA) for comparison between multiple groups. The SEM is depicted by the error bars. P values are *P < 0.05, **P < 0.01, and ***P < 0.001, and P > 0.05 is indicated as not (ns) significant. Sample sizes (n) were incorporated into the text and figure legends, and individual data points from each replicate were depicted as small circles in all figures. The number of times each experiment was independently repeated (N) is incorporated into the figure legends.

SUPPLEMENTARY MATERIALS

stm.sciencemag.org/cgi/content/full/12/574/eabb5647/DC1

Materials and Methods

Fig. S1. DSF screen validation and controls.

Fig. S2. In vitro experiments assess DTHIB binding specificity.

Fig. S3. Evaluation of DTHIB binding to HSF1 by partial proteolysis.

Fig. S4. DTHIB impact on HSF1 target gene expression.

Fig. S5. DTHIB mechanism of action and degradation specificity.

Fig. S6. HSF1 abundance and DTHIB activity in PCa cell lines.

Fig. S7. Assays of DTHIB impact on prostate cancer cell lines from mouse and human.

Fig. S8. DTHIB efficacy in breast cancer and melanoma cell lines.

Fig. S9. DTHIB washout and drug synergy analysis.

Fig. S10. Animal data on DTHIB efficacy and tolerability.

Table S1. List of DNA oligonucleotide primers used and their sequences.

Table S2. List of antibodies used and their technical information.

Data file S1. Raw data on mass spectrometry analysis from the HSF1 partial proteolysis assays (Excel format).

Data file S2. Raw information on RNA-seq analysis in C4-2 human prostate cancer cells (Excel format).

Data file S3. Raw images and information from the TMAs (ZIP format).

Data file S4. Raw measurements and records of all animal experiments (Excel format).

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REFERENCES AND NOTES

Acknowledgments: We gratefully acknowledge advice from E. Burchfiel, M. Mollapour, D. Stokoe, P. Zhou, J. Hong, and S. O’Brien. We thank S. Wardell for guidance on xenograft models; J. Everitt for veterinary pathology analysis; M. Schumacher for guidance on FA; J. Gestwicki for providing the human HSC70 expression vector; S. Lindquist and T. Jacks for providing the NF1-related MEFs; S. Cutler for providing the LATCA small-molecule library; and J. Liu, C. Chong, and D. Sullivan for providing the Johns Hopkins Clinical Compound Library. Funding: This work was partially supported by an award from the Duke Cancer Institute as part of the P30 Cancer Center Support Grant (grant ID: NIH CA014236) to B.D. and D.J.T., a grant from the Prostate Cancer Foundation to J.H., and a Damon Runyon Cancer Research Foundation Postdoctoral Fellowship to A.M.J. Author contributions: All authors contributed to the conception and design of experiments. B.D., A.M.J., P.F.H., D.R.L., J.S.H., and Y.F. conducted experiments. All authors contributed to the interpretation of experimental data. B.D. and D.J.T. wrote the manuscript, and all authors made editorial contributions to the final manuscript. Competing interests: B.D., A.M.J., P.F.H., T.A.H., J.H., and D.J.T. are co-inventors on a technology related to HSF1 inhibitors (U.S. Provisional Patent Application #62/777,831: Compositions and methods for the treatment of cancer). D.J.T. and J.H. have equity and a board relationship with Sisu Pharma Inc. All other authors declare that they have no competing interests. Data and materials availability: All data associated with this study are present in the paper or the Supplementary Materials. Both raw and processed RNA-seq data were deposited to NCBI-GEO (GES155248). The Johns Hopkins Clinical Compound Library (JHCCL) version 1.3 is available from D. Sullivan, J. Liu, and C. Chong under a material agreement with Johns Hopkins University.

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