Clinical management of castration-resistant prostate cancer (CRPC) resulting from androgen deprivation therapy remains challenging. CRPC is driven by aberrant activation of androgen receptor (AR) through mechanisms ranging from its amplification, mutation, post-translational modification, and expression of splice variants (e.g., AR-V7). Herein, we present experimental evidence for therapeutic vulnerability of CRPC to a novel phytochemical, leelamine (LLM), derived from pine tree bark. Exposure of human prostate cancer cell lines LNCaP (an androgen-responsive cell line with mutant AR), C4-2B (an androgen-insensitive variant of LNCaP), and 22Rv1 (a CRPC cell line with expression of AR-Vs), and a murine prostate cancer cell line Myc-CaP to plasma achievable concentrations of LLM resulted in ligand-dependent (LNCaP) and ligand-independent (22Rv1) growth inhibition in vitro that was accompanied by downregulation of mRNA and/or protein levels of full-length AR as well as its splice variants, including AR-V7. LLM treatment resulted in apoptosis induction in the absence and presence of R1881. In silico modeling followed by luciferase reporter assay revealed a critical role for noncovalent interaction of LLM with Y739 in AR activity inhibition. Substitution of the amine group with an isothiocyanate functional moiety abolished AR and cell viability inhibition by LLM. Administration of LLM resulted in 22Rv1 xenograft growth suppression that was statistically insignificant but was associated with a significant decrease in Ki-67 expression, mitotic activity, expression of full-length AR and AR-V7 proteins, and secretion of PSA. This study identifies a novel chemical scaffold for the treatment of CRPC. Mol Cancer Ther; 17(10); 2079–90. ©2018 AACR.
This article is featured in Highlights of This Issue, p. 2077
Prostate cancer continues to be a leading cause of cancer-related deaths among men in western countries despite rigorous screening efforts for early detection of the disease (1). The American Cancer Society estimates diagnosis of about 165,000 new cases of prostate cancer and over 29,000 deaths from this malignancy in the United States alone in 2018. Androgen-receptor (AR) plays an important role in prostate cancer pathogenesis (2–4). AR signaling axis, which is essential for normal male reproductive function, is activated after binding of androgens (e.g., dihydrotestosterone) to the receptor, leading to its nuclear trafficking for subsequent regulation of transcriptional targets, including PSA and transmembrane protease, serine 2 (TMPRSS2; refs. 2–4). Androgen deprivation therapy (ADT) is the standard of care for initial systemic treatment of localized and advanced prostate cancer (5, 6). Unfortunately, a great majority of patients on ADT eventually progresses to castration-resistant prostate cancer (CRPC) within 2 to 3 years (5, 6). Clinically available therapeutic options for advanced and metastatic prostate cancer include anti-androgens such as abiraterone acetate (an irreversible inhibitor of CYP17A1) or enzalutamide, a nonsteroidal antiandrogen (7, 8). However, a subset of patients is inherently resistant to both abiraterone (Zytiga) and enzalutamide (Xtandi) due to expression of constitutively active splice variants of AR like AR-V7 (9). Therefore, identification of novel agents effective against CRPC-expressing splice variants of AR is still desirable.
The molecular understanding of the changeover from androgen dependence to CRPC continues to evolve but AR occupies a central place in this transition (10–13). AR is a 110-kDa transcription factor belonging to the steroid hormone receptor superfamily (2–4). Full-length AR comprises of four major functional domains including an N-terminal domain, a DNA-binding domain, a hinge region containing the nuclear localization sequence, and the C-terminal ligand-binding domain (LBD; ref. 4). Continued dependence on the AR is partly accountable for CRPC development, which may be driven by mechanisms ranging from increased amplification or gain-of-function mutations to ligand-independent activation and expression of C-terminally truncated and constitutively active splice variants (14–18).
The search for novel small-molecule inhibitors of AR continues because of the mechanistic complexity of its aberrant activation in CRPC. Naturally occurring phytochemicals abundant in edible or medicinal plants remain attractive for treatment of cancer (19, 20). This study identifies a novel chemical scaffold, leelamine (LLM, also known as dehydroabietylamine), with activity in human prostate cancer cells. LLM is derived from the bark of pine tree. Growth inhibitory effects of LLM have been studied previously in melanoma cell lines in vitro and in vivo (21, 22). However, this study is the first to demonstrate inhibition of AR expression and activity in prostate cancer cells (LNCaP, C4-2B, 22Rv1, and Myc-CaP), including a cell line that is resistant to enzalutamide (22Rv1). We also provide in vivo evidence for LLM-mediated inhibition of AR expression and its downstream targets (PSA) using 22Rv1 xenograft model. A functionally important noncovalent interaction between LLM and LBD of AR is also shown.
Materials and Methods
Use of mice for this study was approved by the University of Pittsburgh Animal Care and Use Committee.
LLM (purity ≥98%) was purchased from Cayman Chemical Company. Dehydroabietyl isothiocyanate (LLM-ITC) was purchased from American Custom Chemicals Corp., whereas enzalutamide was purchased from Sigma-Aldrich. Reagents for cell culture were purchased from Life Technologies-Thermo Fisher Scientific, and charcoal-dextran–stripped FBS (cFBS) was purchased from HyClone. The synthetic androgen R1881 was purchased from PerkinElmer. The antibody against AR was purchased from Santa Cruz Biotechnology. The anti-PSA antibody was from Dako-Agilent Technologies. An antibody against phospho-AR (Ser210/213) was purchased from Imgenex-Novus Biologicals. FuGENE 6, Dual-Luciferase Reporter Assay Kit, and pRL-CMV were purchased from Promega. AR mutant plasmid pCMV-AR-Y739A was kindly provided by Dr. Elizabeth M. Wilson (University of North Carolina, Chapel Hill, NC). The rat probasin promoter plasmid p159pPr-luc was a gift from Jeffery Green (National Cancer Institute, Bethesda, MD) (Addgene plasmid #8392).
Cell lines and culture conditions
The 22Rv1 and LNCaP cells were obtained from the ATCC. These cell lines were last authenticated by us in March of 2017 and found to be of human origin. The C4-2B cell line was obtained from UroCor, and last authenticated by us in January of 2015. The 22Rv1, LNCaP, and C4-2B cells were maintained in RPMI1640 supplemented with 10% FBS, antibiotic mixture, sodium pyruvate, HEPES, and 2.5 g/L glucose. Myc-CaP cells were kindly provided by Dr. Charles L. Sawyers (Department of Medicine, University of California, CA). This cell line was not authenticated by us. The Myc-CaP cells were maintained in DMEM supplemented with 10% FBS, and antibiotic mixture. Normal human prostate cells (PrSC) were purchased from Lonza, and cultured in growth medium supplied by the provider. The PC3 cells with stable overexpression of GFP-AR (PC3-AR) were a generous gift by Dr. Zhou Wang (Department of Urology, University of Pittsburgh, PA). The PC3-AR cells were not authenticated by us. PC3-AR and corresponding empty vector transfected cells (PC3-EV) were maintained in RPMI-1640 medium supplemented with 10% FBS and G418 (600 μg/mL). For the experiments that required androgen-depleted condition, cells were maintained in phenol red-free media supplemented with 10% cFBS.
Cell viability assay
The effects of LLM, enzalutamide, and LLM-ITC on cell viability were determined by trypan blue dye exclusion assay as described by us previously (23). Briefly, prostate cancer cells or PrSC cells were seeded in 12-well plates, and allowed to attach by overnight incubation. The cells were then treated with the specified concentrations of the test agents for 24 or 48 hours. Cells were trypsinized and stained with trypan blue. The live cells were counted under an inverted microscope.
Cell proliferation assay
LNCaP cells or C4-2B cells (750 cells/well) were seeded in 96-well plates. After 16 hours of incubation, cells were treated with ethanol (control) or the indicated doses of LLM or synthetic androgen R1881 for 24, 48, and 72 hours. Subsequently, 20 μL of the manufacturer's supplied color development reagent (MTS, Promega) was added to each well and the plates were incubated at 37°C for 2 hours. Absorbance was measured at 492 nm.
Cells (500 cells/well) were seeded in 6-well plates. After overnight incubation to allow attachment of the cells, they were treated with different concentrations of LLM. The medium containing ethanol (control) or LLM was replaced every third day. After 10 days of treatment, cells were rinsed with PBS, fixed with 100% methanol for 5 minutes, and stained with 0.5% crystal violet solution in 20% methanol for 30 minutes at room temperature. The colonies were counted using GelCount (Oxford Optronix).
Details of lysate preparation and immunoblotting have been described by us previously (24). The cells were treated with desired concentrations of LLM or its analogue for different time points. Western blot analysis was performed as described previously by us (24). In some experiments, cells were pretreated with 1.5 μmol/L of MG132 for 1 hour followed by treatment with LLM for an additional 12 hours.
Microscopy for nuclear translocation of AR
The 22Rv1 (7 × 104) or LNCaP (5 × 104) cells were plated in triplicate on coverslips in 12-well plates in phenol red-free medium supplemented with 10% cFBS and allowed to attach by overnight incubation. Cells were treated with ethanol or LLM for 3 hours followed by addition of R1881. The plates were incubated for an additional 9 hours. The cells were washed with PBS and fixed in 2% paraformaldehyde for 1 hour followed by blocking with a solution containing 0.5% bovine serum albumin and 0.15% glycine in PBS. After blocking, cells were incubated with AR antibody (4°C; overnight) followed by treatment with Alexa Fluor 488-conjugated secondary antibody for 1 hour at room temperature. The cells were counterstained with 4′,6-diamidino-2-phenylindole (DAPI; 50 ng/mL) and examined under a Leica DC 300F fluorescence microscope.
RNA isolation and real-time RT-PCR
The expression of AR mRNA and its target genes (PSA and TMPRSS2) were determined by RT-PCR. Total RNA from ethanol- and LLM-treated cells was isolated using RNeasy Kit. One μg RNA was used for cDNA synthesis with the use of SuperScript III reverse transcriptase and oligo (dT)20 primer. Quantitative PCR was performed using 2× SYBR Green master mix for 40 cycles. The expression of AR and its target genes were normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH). The primers for human AR, PSA, TPMRSS2, and GAPDH were as follows: Forward (AR): 5′-ATGGTGAGCAGAGTGCCCTA-3′; reverse (AR) 5′-GTGGTGCTGGAAGCCTCTCCT-3′; forward (PSA): 5′-AAAAGCGTGATCTTGCTGGG-3′; reverse (PSA): 5′-CATGACCTTCACAGCATCCG-3′; forward (TMPRSS2): TCTAACTGGTGTGATGGCGT-3′; reverse (TMPRSS2): 5′-GGATCCGCTGTCATCCACTA-3′; 5′- forward (GAPDH): 5′-GGACCTGACCTGCCGTCTAGAA-3′; reverse (GAPDH): 5′-GGTGTCGCTGTTGAAGTCAGAG-3′. The PCR conditions were as follows: 95°C for 10 minutes followed by 40 cycles of 95°C for 15 seconds, 60°C (AR, TMPRSS2, and GAPDH) and 63°C (PSA) for 1 minute, and 72°C for 30 seconds.
Quantitation of PSA in cell culture medium
Desired cells (22Rv1 and LNCaP-5-7 × 104) were plated in triplicate in 12-well plates in phenol red-free medium containing 10% cFBS. After attachment, cells were treated with ethanol or LLM for 24 hours. Media were collected and centrifuged at 3,500 rpm for 15 minutes. Equal volume of supernatant was used to determine PSA levels using Quantikine Human KLK3/PSA Immunoassay Kit from R&D Systems.
The 22Rv1 or LNCaP cells were plated in triplicate in phenol red-free media containing cFBS and allowed to attach by overnight incubation. The cells were then treated with ethanol, 1 nmol/L R1881, and/or indicated doses of LLM for 24 hours. Apoptosis was quantified by analysis of histone-associated DNA fragment release into the cytosol or by flow cytometry after staining the cells with Annexin V/propidium iodide as described by us previously (25).
Molecular docking for the LBD of AR with LLM or LLM-ITC was performed using HEX 8.0 software. The coordinates for LLM and LLM-ITC were taken from the SDF files and converted into pdb format using Discovery Studio 4.1 software. The crystal structure of AR (PDB ID: 2PIW) was retrieved from the Protein Data Bank (http://www.rcsb.org./pdb). Visualization of the model was performed using Discovery Studio 4.0 software. A “by default” parameter was used for the docking calculation with correlation type shape only, FFT mode at 3D level, grid dimension of 6 with receptor range 180 and ligand range 180 with twist range 360 and distance range 40. The resulting models were visually inspected, during which one minor adjustment was made to eliminate a steric conflict between LLM and an amino acid side chain on the surface of AR LBD.
Transient transfection and luciferase reporter assay
PC-3 cells (4 × 104) were plated in triplicate in phenol red-free Opti-MEM medium containing 10% cFBS and 10 nmol/L R1881. The cells were cotransfected with 2 μg of mutant AR or 2 μg of rat probasin luciferase (pPr-luc) and 0.5 μg pCMV-RL plasmid for 24 hours. After transfection, cells were treated with ethanol or LLM for 12 hours. The transient transfection was achieved using FuGENE6. Luciferase activity was measured using Dual-Luciferase Reporter Assay System (Promega) following the manufacturer's protocol.
Twelve male SCID (NOD.CB17-PRkdcscid/J) mice at 4 to 5 weeks of age were purchased from The Jackson Laboratory. After a 5-day acclimation, fur was removed from the torso of each animal in the area directly above each hind limb using scissors. Both sides of each mouse in that area were injected with 2 × 106 22Rv1 cells suspended in 200 μL of serum-free medium diluted 1:1 with Matrigel (BD Biosciences). Cells were grown to approximately 60% confluency to ensure that the cells were in active growth phase. One week postimplantation, the mice were divided into two groups. Mice of group 1 were treated with 100 μL vehicle, whereas group 2 mice received 9.1 mg LLM/kg by intraperitoneal injection 5 times per week. The vehicle consisted of 10% ethanol, 10% DMSO, 30% Kolliphor EL (Sigma-Aldrich), and 50% PBS. At the onset of the study, mice were weighed and this measurement continued on a weekly basis. Tumor volume measurements were taken using Vernier calipers as soon as tumors became measurable and continued 3 times each week until the conclusion of the study. Treatment continued until the tumor burden exceeded 2,000 mm3 at which time the animals were euthanized by CO2 overdose (supplied via compressed gas cylinder) and blood, tumor tissue, and vital organs were harvested. A portion of each tumor and all vital organs were fixed in 10% neutral buffered formalin for hematoxylin and eosin (H&E) or IHC. The other portion of each tumor was placed on dry ice and later stored at −80°C. Blood was collected using a heparinized needle then placed on ice and later centrifuged at 3,000 RPM for 5 minutes. Plasma was removed and stored at −20°C.
IHC was performed as described by us previously (26, 27) with some modifications. Briefly, 4- to 5-μm-thick tumor sections were de-paraffinized, hydrated in graded alcohol, and then washed with PBS. Sections were treated with 0.3% H2O2 in 100% methanol for 20 minutes at room temperature and then incubated with the blocking buffer for 1 hour. Subsequently, the tumor sections were treated with the anti–Ki-67 antibody overnight in humid chambers at room temperature. After washing, sections were incubated with horseradish peroxidase-conjugated secondary antibody for 1 hour at room temperature. A characteristic brown stain was developed with 3,3′-diaminobenzidine. Stained sections were examined under a Leica DC300F microscope. At least five non-overlapping representative images were captured from each section, and analyzed with the Aperio ImageScope v9.1 software (Aperio) using nuclear algorithm.
Statistical analyses were carried out using GraphPad Prism (version 6.07). Statistical significance of difference was determined by the one-way analysis of variance (ANOVA) followed by Dunnett or Bonferroni test or unpaired Student t test.
LLM treatment inhibited viability of prostate cancer cells in vitro in association with downregulation of AR protein
Two-well characterized human prostate cancer cell lines (22Rv1 and LNCaP), an androgen-insensitive variant of LNCaP cells, a murine prostate cancer cell line (Myc-CaP), and a normal prostate cell line (PrSC) were used to determine the growth inhibitory effect of LLM (structure of LLM is shown in Fig. 1A). Pharmacokinetics of LLM has been determined in male ICR mice after a single oral administration at 10 mg/kg body weight (28). Peak plasma concentration (Cmax) of LLM was about 2.8 μmol/L with a Tmax (time to reach Cmax) of 4.7 hours and plasma half-life of 5.7 hours (28). Therefore, LLM concentrations of 0.5, 1, 2.5, and/or 5 μmol/L were used to determine its effect on viability of prostate cancer cells. LLM treatment inhibited viability of 22Rv1 and LNCaP cells in a concentration-dependent manner (Fig. 1B). Viability of Myc-CaP cell line was also inhibited significantly upon LLM treatment (Supplementary Fig. S1A). However, the inhibitory effect of LLM treatment on viability of Myc-CaP cells was relatively less pronounced compared with 22Rv1 or LNCaP (Supplementary Fig. S1A), which may be attributable to very high expression of the Myc oncoprotein. Further work is necessary to test this possibility, but PrSC cells were relatively more resistant to cell viability inhibition by LLM compared with prostate cancer cells (Fig. 1B). For example, viability of 22Rv1 cells was decreased by >90% after 24-hour treatment with 5 μmol/L LLM. The viability of PrSC was not affected at all after 24-hour treatment with 5 μmol/L LLM (Fig. 1B). We also found that the 22Rv1 cell line was completely resistant to cell viability inhibition by enzalutamide concentrations of 2.5 and 5 μmol/L (Fig. 1C). As expected, the LNCaP cell line, but not 22Rv1, was sensitive to growth stimulation by a synthetic androgen (R1881; Fig. 1D). The R1881-stimulated growth of LNCaP cell line was also suppressed significantly in the presence of LLM (Fig. 1D). Furthermore, LNCaP and C4-2B cells were more or less equally sensitive to cell proliferation inhibition by LLM regardless of R1881 treatment (Supplementary Fig. S2). Clonogenic assay confirmed cell survival inhibition by LLM (Fig. 1E and F). These results indicated anticancer effect of LLM in prostate cancer cells regardless of the androgen responsiveness.
Treatment of 22Rv1, LNCaP, and C4-2B human prostate cancer cells (Fig. 2A) and the Myc-CaP cell line (Supplementary Fig. S1B–S1C) with LLM resulted in a dose-dependent suppression of protein levels of full-length AR as well as its splice variants, including AR-V7, and phosphorylated AR. LLM-mediated suppression of full-length AR was evident at both 12- and 24-hour time points. Densitometric quantitation of the full-length AR (22Rv1, LNCaP, and C4-2B) and AR-V7 proteins (22Rv1) in LLM-treated cells normalized to corresponding solvent-treated control are shown in Supplementary Fig. S3. An antibody specific for AR-V7 was used to determine the effect of LLM on AR-V7 expression in 22Rv1 cells. Similar to the full-length AR, LLM treatment caused a decrease in protein level of AR-V7 in 22Rv1 cells (LNCaP or C4-2B cells do not express AR-V7). As can be seen in Fig. 2B, the AR protein was predominantly nuclear in 22Rv1 cells. Nuclear level of AR protein was decreased markedly in the presence of LLM irrespective of the R1881 treatment (Fig. 2B). On the other hand, R1881-stimulated nuclear translocation of AR in LNCaP cells (Supplementary Fig. S4). Similar to the 22Rv1 cells, however, nuclear level of AR protein was reduced following LLM exposure with or without R1881 treatment (Supplementary Fig. S4). Downregulation of AR by LLM treatment was accompanied by suppression of AR target genes PSA and TMPRSS2 (Fig. 2C and D). Consistent with these results, protein levels (Fig. 2E) and/or secretion of PSA (Fig. 2F) were decreased markedly upon treatment of 22Rv1, LNCaP, and C4-2B cells with LLM. Densitometric quantitation of PSA levels in LLM-treated 22Rv1, LNCaP, and C4-2B cell lysates normalized for corresponding solvent control are shown in Supplementary Fig. S5. Collectively, these results indicated inhibition of AR expression and activity by LLM treatment in prostate cancer cells.
Transcriptional suppression of AR by LLM treatment
Expression of AR mRNA was also decreased following LLM treatment in both 22Rv1 and LNCaP cells (Fig. 3A). We used a proteasomal inhibitor (MG132) to determine the role of posttranscriptional mechanisms in AR downregulation by LLM treatment. LLM-mediated downregulation of AR protein expression was not reversed in the presence of MG132 in either cell line (Fig. 3B; Supplementary Fig. S6A). These results indicated transcriptional suppression of AR following LLM treatment.
LLM treatment resulted in apoptotic cell death in prostate cancer cells
Treatment of 22Rv1 and LNCaP cells with LLM resulted in increased release of histone-associated DNA fragments into the cytosol, a measure of apoptotic cell death, in comparison with vehicle-treated control, which was not affected by the presence of R1881 (Fig. 3C). The 22Rv1 cell line was relatively more sensitive than LNCaP to apoptosis induction by LLM (Fig. 3C), which was consistent with cell viability inhibition data (Fig. 1B). Annexin V methods confirmed apoptosis induction by LLM treatment regardless of R1881 exposure (Fig. 3D and E).
Overexpression of AR resulted in sensitization of PC-3 cells to growth inhibition by LLM
In comparison with 22Rv1 or LNCaP cells (Fig. 2A), AR protein level was decreased to a lesser extent by LLM treatment in PC-3 cells stably transfected with GFP-AR plasmid (Fig. 3F; Supplementary Fig. S6B). This finding, and the observation that LLM decreases GFP-AR level to a lesser extent in PC-3 cells than in other cell lines, makes it more likely that LLM acts to suppress the activity of the endogenous AR promoter (vs. the strong promoter driving GFP-AR expression). Interestingly, overexpression of AR resulted in sensitization of PC-3 cells to growth inhibition by LLM especially at the higher concentrations as revealed by trypan blue dye exclusion assay (Fig. 3G). These results were confirmed by clonogenic assay (Fig. 3H and I). These results indicated that AR is a valid therapeutic target of LLM.
Interaction of LLM with amino acid residues in the LBD of AR
AR is a modular protein consisting of an N-terminal domain, a central zinc-finger DNA-binding domain, a hinge region, and a highly structured LBD (4). LLM is not electrophilic and hence a covalent interaction is not expected. However, molecular docking identified a binding-pocket for LLM in AR (Fig. 4A). The bottom of the LLM-binding pocket is formed by the side chains of A735, Y739, P817, and V821. The hydroxyl group of Y739 is positioned to form an on-face hydrogen bond with the π-electron cloud of the LLM phenyl ring system. The wall of the pocket is formed by the K822, K731, M734, K905, and Q902 side chains, among which the K905 and Q902 are in close contact with LLM. One side of the LLM-binding pocket is open. On the left-hand side of the opening as shown in Fig. 4A is located the D819 side chain. Potentially, the D819 side chain carboxyl group is able to form a salt bridge with the LLM amine group. A salt bridge is the combination of hydrogen bonding and electrostatic interactions. It may contribute significantly to the stability of the LLM:AR-LBD complex.
We tested the functional significance of one of these potential interactions using the PC-3 cell line, which lacks AR expression. As can be seen in Fig. 4B, overexpression of the wild-type (WT) AR in PC-3 cells resulted in an increase in probasin luciferase activity in the presence of R1881, which was decreased significantly by LLM treatment. The Y739A mutation significantly attenuated LLM-mediated suppression of probasin luciferase reporter activity (Fig. 4B). The LLM-mediated suppression of AR protein was also abolished by Y739A mutation. LLM treatment decreased AR protein level in PC-3 cells overexpressing WT AR (Fig. 4C).
The role of amine group in AR inhibition by LLM
We used an analogue of LLM, dehydroabietyl isothiocyanate (LLM-ITC; structure is shown in Fig. 5A), to further understand the significance of amine group in AR suppression by LLM. Molecular docking revealed a different mode of interaction between LLM-ITC and AR LBD (Fig. 5B). The protein levels of full-length AR, splice variants of AR, or PSA were not decreased by LLM-ITC treatment in 22Rv1, LNCaP, or AR-overexpressing PC-3 cells (Fig. 5C; Supplementary Fig. S7). In addition, both 22Rv1 and LNCaP cells were significantly more resistant to growth inhibition by LLM-ITC treatment (Fig. 5D) in comparison with LLM (Fig. 1B). Consistent with these results, probasin luciferase activity was not affected by LLM-ITC treatment (Fig. 5E). Collectively, these results indicated a critical role for the amine group in suppression of cell growth and AR by LLM.
In vivo downregulation of AR, AR-V7, and PSA in 22Rv1 xenografts
We used the 22Rv1 xenograft model for the in vivo studies because: (i) this cell line rapidly develops tumor upon implantation as compared with LNCaP cells, and (ii) unlike LNCaP, the 22Rv1 cell line allows determination of the effect of LLM treatment on protein levels of both full-length AR and its splice variants. Figure 6A shows tumor volume in individual mouse of the control and the LLM treatment group. The mean tumor volume was lower by 34% in the LLM treatment group compared with control but the difference was insignificant due to large data scatter and a small sample size. Figure 6B depicts representative IHC for Ki-67 and H&E staining for tumor of a control mouse and a tumor of LLM-treated mouse. The Ki-67 expression as well as mitotic count was significantly lower in the tumors of LLM-treated mice compared with control (Fig. 6C). Figure 6D shows Western blots for AR, AR-V7, and PSA using tumor supernatants of control and LLM-treated mice. Expression of both full-length AR and AR-V7 was significantly lower in the tumors of LLM-treated mice in comparison with controls (Fig. 6E). In addition, tumor expression of PSA protein (Fig. 6F) as well as its circulating level (Fig. 6G) was lower in LLM treatment group compared with control. LLM treatment did not cause weight loss or any other side effects (Supplementary Fig. S8). These results indicated downregulation of AR and its target PSA in vivo upon LLM administration to 22Rv1 xenograft bearing mice.
The current study is the first to show inhibitory effect of LLM on AR in prostate cancer cells. We also found that LLM inhibits growth of prostate cancer cells that are resistant to clinically used antiandrogen enzalutamide due to expression of AR-V7. The LLM-mediated inhibition of prostate cancer cell growth is accompanied by downregulation of AR and its target PSA both in vitro and in vivo. It is important to point out that growth inhibition and AR downregulation by LLM treatment is observed at pharmacologic doses.
The current study suggests a critical role for noncovalent interactions of LLM with A735, Y739, P817, V821, K822, D731, M734, K905, and Q902 of AR LBD. The fact that AR downregulation and transcriptional activity inhibition by LLM is abolished by Y739A mutation of AR provides experimental evidence for functional significance of one of the interactions. However, other interactions may also be important. For example, the nuclear/cytoplasmic shuttling of AR is regulated by a nuclear localization signal (residues 617-633) at the junction of DNA-binding domain and the hinge region (29, 30) and a ligand-regulated nuclear export signal (residues 742-817; ref. 31). Because LLM interacts with P817, the possibility that this interaction is responsible for LLM-mediated inhibition of nuclear localization of the AR protein cannot be excluded. However, further studies are needed to explore this possibility. LLM decreases protein level of AR-V7 that lacks LBD domain of AR. It is possible that abrogation of AR activation of the probasin target promoter by overexpression of AR-Y739A is due to increased abundance of the mutant protein and probably its half-life. Further work is also necessary to test this possibility.
On one hand, the current study suggests that noncovalent interaction with residues in AR LBD, including P817, may contribute to AR inhibition by LLM as the ITC derivative, which does not interact with this AR LBD domain, loses its efficacy. On the other hand, LLM is effective against AR-V7, which lacks the LBD and hence the amino acids for noncovalent interactions, indicating that the primary mechanism of its action is likely through inhibition of AR promoter activity. LLM may even work against an overlapping transcriptional mechanism as evidenced by its ability to inhibit AR-dependent anchorage-independent growth of PC3-AR cells.
Even though LLM administration to 22Rv1 xenograft bearing SCID mice resulted in statistically significant decreases in expression of AR and PSA, the tumor volume was not significantly different between the control and LLM treatment groups possibly due to a few outliers. The LLM dose used in the present is about 40% of the MTD of LLM (21). Thus, the possibility that the higher concentrations of LLM are effective for growth inhibition cannot be ruled out without further experimentation.
The mechanistic understanding of the antitumor effect of LLM is restricted to a few publications using melanoma cell lines (21, 22). Antitumor effect of LLM in melanoma cell lines in vitro and in vivo was associated with inhibition of Akt, Stat3, and Erk1/2 activation (reduced phosphorylation; refs. 21, 22). The inhibition of these prosurvival and oncogenic pathways upon LLM treatment was observed as early as 3 hours after treatment at 3 to 5 μmol/L concentrations (21, 22). However, the precise role and contribution of these pathways in growth inhibition and cell death induction by LLM is still unclear. Studies have also shown that ectopic expression of AR in PC-3 renders them more susceptible to apoptosis (32). This study does reveal proapoptotic effect of LLM. However, it is possible that LLM-induced apoptosis in prostate cancer cells is mediated by modulation of Akt, Stat3, and/or Erk1/2. Further studies are needed to explore the role of Akt, Stat3, and Erk1/2 in apoptosis induction by LLM in prostate cancer cells.
Despite exciting mechanistic insights presented in this study, further preclinical studies are needed in preparation for the clinical development of LLM, including (i) determination of the dose–response effect of LLM treatment on in vivo growth of prostate cancer cells other than 22Rv1 (LNCaP and C4-2B) as well as patient-derived xenografts, (ii) determination of the oral bioavailability of LLM and a careful analysis of the clinical pharmacology and metabolism of LLM, (iii) determination of an appropriate dosing schedule of LLM that could then be taken in to the clinic, and (iv) determination of the toxicity profile of LLM administration by analyzing a wide range of normal host tissues. If LLM is found to be well tolerated in the mouse model, additional toxicology studies in a larger sized animal model to determine the safety of this agent will be required. The findings from these preclinical studies should then provide the rational basis for designing the first-in-man phase I clinical studies of LLM.
In summary, the results of the current study indicate that AR is a novel mechanistic target of prostate cancer cell growth inhibition by LLM. We also provide in vitro (human and murine prostate cancer cell lines) and in vivo (22Rv1 xenografts in SCID mice) evidence for inhibition of AR expression and activity following LLM treatment. Finally, the current study reveals that noncovalent interactions of LLM with AR LBD are functionally important.
Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
Conception and design: K.B. Singh, S.V. Singh
Development of methodology: K.B. Singh, S.V. Singh
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): K.B. Singh
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): K.B. Singh, X. Ji, S.V. Singh
Writing, review, and/or revision of the manuscript: K.B. Singh, X. Ji, S.V. Singh
Study supervision: S.V. Singh
This work was supported by the grant RO1 CA101753 awarded by the National Cancer Institute (to S.V. Singh) and the Intramural Research Program of the NIH, NCI, Center for Cancer Research (to X. Ji). This research used the Animal Facility and the Tissue and Research Pathology Facility supported in part by Cancer Center Support Grant from the NCI (P30 CA047904; to Robert L. Ferris-principal investigator).
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.