Abscisic acid signaling through LANCL2 and PPARγ induces activation of p38MAPK resulting in dormancy of prostate cancer metastatic cells
- Authors:
- Published online on: January 8, 2024 https://doi.org/10.3892/or.2024.8698
- Article Number: 39
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Copyright: © Parajuli et al. This is an open access article distributed under the terms of Creative Commons Attribution License.
Abstract
Introduction
Prostate cancer (PCa) is the second leading cause of death from cancer in men in the United States (1). The majority of men who are diagnosed with localized PCa and who elect to undergo therapy are treated with surgery or radiation, by which they expect to be cured (2). A number of patients experience a prolonged period without evidence of biochemical recurrence or obvious disease, but some relapse with single or multiple metastases frequently detected in bone (3,4). Notably, ~90% of patients who succumb to advanced-stage PCa have bone metastases, whereas only ~10% of these individuals had bone metastases at the time of diagnosis (5–7). These observations suggest that disseminated tumor cells (DTCs) may have left the primary tumor site in the prostate and took up residence in distant sites. For PCa, it has been shown that DTCs may be present in the bone marrow at the time of initial diagnosis (8–11). In the bone marrow, DTCs maintain the ability to become reactivated and generate metastases by poorly understood processes (8–11). Further studies are needed in the context of PCa DTC dormancy to identify and develop improved therapeutic strategies to treat metastatic diseases.
One well-delineated pathway in the understanding of cellular dormancy is p38/ERK signaling. High levels of p38MAPK activity function as an inhibitory regulator of ERK, thus preventing cell proliferation by inducing G0/G1 phase arrest, or by initiating senescence and apoptosis (12–14). Transforming growth factor-β2 (TGF-β2), which is secreted from bone marrow-derived cells, is known to induce expression of a p38high/ERKlow phenotype in cancer (15). The subsequent activation of Smad1/5 and increased expression of p27 results in the downregulation of cyclin-dependent kinase (CDK)4, which collectively facilitates the transition into cellular quiescence (15,16). Additionally, p38high/ERKlow facilitates G0/G1 phase arrest via the regulation of various factors, including NR2F1, and CDK inhibitors p27 and p21. Therefore, the combined regulation of transcription factors by a p38high/ERKlow phenotype is responsible for quiescence (17). A second well-described pathway that regulates PCa dormancy in the bone marrow is activated by growth arrest-6 (GAS6) signaling. In bone marrow, GAS6 secreted by osteoblasts activates the TAM family of receptors (Tyro3, Axl and MerTK) on PCa cells, which serve as molecular switches to turn on (Tyro3, Axl) and off (MerTK) the dormancy programs (18). Similarly, bone morphogenetic protein (BMP)7, which is produced by bone stromal cells, induces dormancy in PCa cells by activating p38 signaling (19). These studies have suggested that cells in the bone marrow microenvironment, including osteoblasts, serve key roles in regulating PCa dormancy.
The present study explored the signaling pathways activated by the dormancy-inducing phytohormone ABA in PCa cells. ABA has long been known as a regulator of plant growth and survival under stress conditions, but has more recently been detected in rat brain tissues (20). Since its discovery in mammals, ABA has been identified in blood plasma (21), and has been demonstrated to be produced by pancreatic β cells (22), adipocytes (21), keratinocytes (23), hematopoietic immune cells, such as granulocytes (24), monocytes (25) and macrophages (26), and human mesenchymal stem cells (27). ABA has also been shown to be linked to several human diseases, including type-2 diabetes and colitis (28,29). ABA signaling is mediated by two known receptors, lanthionine synthetase C-like protein 2 (LANCL2) and peroxisome proliferator activated receptor γ (PPARγ) (28–30). Numerous studies have demonstrated that LANCL2 serves a role in the regulation of stress responses, inflammation, and in metabolic and immune-related diseases (31,32). It has also been demonstrated that LANCL2 is expressed by DTCs in the bone marrow of patients with breast cancer (30). PPARγ is a nuclear hormone receptor, the signaling of which results in anti-proliferative activities by decreasing the levels of cyclin D1 and E, and increasing the expression of p21 and p27 in PCa (33–35). The present study further explored the role of ABA in dormancy and the molecular pathways activated by its signaling in PCa.
Materials and methods
Reagents
A number of the reagents used, their source and catalogue numbers are presented in Table I; short hairpin RNA (shRNA) targeting sequences are presented in Table II; primer sequences are presented in Table III.
Cell culture
As the present study aimed to study bone marrow DTCs, the following metastatic cell lines were used. The human androgen-independent bone metastatic PCa cell lines PC3 and LnCaP subline C4-2B were predominantly used in the present study, as they metastasize to bone (36–38). When injected into bone, PC3 cells result in predominantly lytic lesions, whereas C4-2B cells may produce mixed osteoblastic and osteolytic lesions (36–38). In some experiments, the human dura metastasis cell line DU145 and murine Myc-CaP cells were used. We recently demonstrated that Myc-CaP cells can also metastasize to bone (39). The human PCa cells were routinely grown in RPMI 1640, whereas the murine Myc-CaP cells were grown in DMEM. All cultures were supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin, and were maintained in an incubator set to 5% CO2 atmosphere at 37°C (Marshall Scientific).
Cell viability assays
To evaluate viability, 3,000 PCa cells/well (PC3, C4-2B, Du145 and Myc-CaP) in 100 µl medium were added to 96-well plates in quintuplicate, and allowed to adhere overnight. The next day, the cells were washed in PBS (Thermo Fisher Scientific, Inc.) and fresh medium was added. ABA was diluted in dimethyl sulfoxide (DMSO), and was added at the following doses: 0 µM (DMSO only), 25, 50, 100 and 200 µM for 72 h at 37°C. To determine viability, cells were either recovered from culture with trypsin (MilliporeSigma) and counted using a hemocytometer under a Nikon Eclipse Ts2 inverted phase light microscope (Nikon Corporation) following trypan blue staining (at 23°C for 3 min), or were assessed using a colorimetric assay. For the colorimetric assays, a total of 70 h after the addition of ABA or control, 20 µl CellTiter 96® Aqueous Non-Radioactive Cell Proliferation Reagent (cat. no. G5421; Promega Corporation) was added to the cultures for 2 h at 37°C. The optical density (O.D.) of the cell culture plates was then measured using a plate reader (DU530 UV/Vis scanning spectrophotometer; BD Biosciences) at a wavelength of 490 nm. To determine if ABA permanently affected viability, cells were treated with ABA for 72 h at 37°C, washed, trypsinized and counted. An equal number of cells (1×105 cells/well in 12-well plates, 1 ml of culture media) were then added back to culture for an additional 72 h at 37°C and cell viability was evaluated by cell counting using a hemocytometer. Cell viability (%) was calculated as follows: (treatment group-background)/(control group-background)x100. The data are presented as the mean ± SEM.
Cytotoxicity assays
The cytotoxicity of ABA was assessed using a CytoTox96 Non-Radioactive Cytotoxicity Assay kit (Promega Corporation) according to the manufacturer's instructions. Briefly, PCa cell lines (PC3, C4-2B, DU145 and Myc-CaP) were seeded on 96-well plates (1×104 cells/well) in quintuplicate per group in 100 µl complete culture medium. After overnight incubation, the cells were treated with 25, 50, 100 and 200 µM ABA for 4 h at 37°C; the control group was treated with DMSO. To measure lactate dehydrogenase (LDH) release, 50 µl supernatant was mixed with 50 µl reconstituted substrate mix solution in a new 96-well plate and was incubated for 30 min in the dark at room temperature. The reaction was stopped with 50 µl stop buffer and the O.D. was then measured at a wavelength of 490 nm. Each cytotoxicity assay was independently repeated two times.
shRNA and lentivirus preparation
pLKO.1 lentiviral vector-based shRNAs targeting specific candidate genes and the non-specific (NS) control shRNA were obtained from Horizon Discovery (NS control and shRNA LANCL2) and MilliporeSigma (shRNA PPARγ) (Table II). Lentivirus particles were prepared by transfecting 293T cells in 12-well plates with 0.5 µg either gene-specific shRNA plasmids (LANCL2, clone ID no. TRCN0000045403; and PPARγ, clone ID no. TRCN0000001670) or NS shRNA plasmids along with lentiviral packaging plasmids [2nd generation psPAX2 packaging (plasmid #12260; Addgene, Inc.) and pMD2.G envelope (plasmid #12259; Addgene, Inc.) in a 1:1 ratio for 48 h at 37°C. All lentiviral transfections were performed using Effectene Transfection Reagent. Stable cell lines were generated by infecting PCa cells (PC3 and C4-2B) with a multiplicity of infection of ~250 viral particles per cell in 200 µl (total collection volume, 1 ml) in 12-well plates for 24 h, followed by selection in puromycin (1 µg/ml) at 37°C for 1 week. Thereafter reverse transcription-quantitative PCR (RT-qPCR) was used to assess the mRNA expression changes and western blotting was used to validate protein expression changes. After the initial assessment of the efficiency of shRNA knockdown (KD) of the target genes, clone 1 was selected for both LANCL2 and PPARγ for further studies. Thereafter, frozen stocks were established and cells were used for experimentation after validation of the targeted KD within 2 weeks of transfection or thaw.
RNA extraction, cDNA preparation and RT-qPCR
Total RNA was extracted from target cells using TRIzol® Reagent (Invitrogen; Thermo Fisher Scientific, Inc.) and was purified using the RNeasy Mini Kit (Qiagen, Inc.). cDNA was synthesized using the ProtoScript First Strand cDNA Synthesis Kit (New England Biolabs, Inc.) according to the manufacturer's instructions. qPCR was performed with gene-specific primers using the iTaq Universal SYBR® Green Supermix (Bio-Rad Laboratories, Inc.) on an Applied Biosystems 7500 thermocycler system (Applied Biosystems; Thermo Fisher Scientific, Inc.). The thermocycling conditions were as follows: Initial denaturation at 95°C for 15 min; 40 cycles at 95°C for 15 sec, annealing at 60°C for 30 sec and 72°C for 30 sec; and a final extension step at 72°C for 7 min. Results were normalized against β-actin levels using the formula ΔCq=Cq of target gene-Cq of β-actin. The mRNA expression levels of the control group were used to establish the baseline; therefore, ΔΔCq was calculated using the formula ΔΔCq=ΔCq of target gene-ΔCq of the baseline. The fold change of mRNA expression was calculated as fold=2−ΔΔCq (40). The primer sequences for all the genes analyzed in the present study are provided in Table II.
Migration and invasion assays
Migration assays were performed using a Transwell with 8.0 µm pore polycarbonate membrane inserts (cat. no. 3422; Corning, Inc.), and invasion assays were performed using BioCoat Growth Factor Reduced Matrigel Invasion Chambers (cat. no. 354483; Corning, Inc.) in Transwell inserts. For these studies, PC3 and C4-2B cells infected with NS, LANCL2 and PPARγ shRNAs were serum-starved for 6 h, and were then recovered from culture and seeded in triplicate in the upper chamber of Transwell inserts (5×104 cells/well). The lower chambers were filled with medium containing 10% FBS as a chemoattractant. The cells in the upper chambers were treated with 100 µM ABA or DMSO as a control at 37°C. After 24 h for PC3 cells and 48 h for C4-2B cells, the cells remaining in the top chambers (those cells that had not migrated or invaded) were removed with cotton-tipped applicators. The number of cells that had migrated or invaded into the lower chambers were quantified following DAPI staining at 23°C for 3 min; 5–8 fields/membrane were evaluated under an All-in-one Fluorescence Microscope (BZ-X800/BZ-X810; Keyence Corporation; ×10 magnification) and nuclei quantification was performed using ImageJ 1.53K software (National Institutes of Health). The percentage of migration or invasion was calculated as follows: (number of cells that migrated or invaded in the treated group)/(number of cells that migrated or invaded in the control group) ×100.
Soft agar assays
PC3 and C4-2B cells (5×103/well) infected with LANCL2, PPARγ or NS shRNAs were seeded into 6-well plates on 0.4% low gelling point agarose (cat. no. A9045; MilliporeSigma) and layered on top of 0.8% agarose. The cultures were maintained for 15–18 days, with fresh medium replenished every third day containing DMSO (control) or 100 µM ABA. At the end of the experiment, the images of the cell colonies (>30 cells) in soft agar were captured using an inverted light microscope (Olympus Corporation; ×10 magnification). Colony size was measured using ImageJ 1.53K software (National Institutes of Health) plotted as relative colony size (%) when compared with control cells.
Western blot analysis
Whole cell protein extracts were prepared using RIPA lysis buffer (Pierce; Thermo Fisher Scientific, Inc.) containing a Protease Inhibitor Cocktail (Roche Diagnostics) and Phosphatase Inhibitor (Thermo Fisher Scientific, Inc.). Lysed samples were centrifuged at 18,000 RCF for 15 min at 4°C, and the clarified supernatants were collected and stored at −80°C. Protein concentrations were determined using the Bradford Protein Assay Reagent (Bio-Rad Laboratories Inc.). Equal amounts of protein samples (25 µg) were separated by 10 or 12% SDS-PAGE and transferred onto polyvinylidene difluoride membranes (Thermo Fisher Scientific, Inc.) using wet-transfer apparatus (Bio-Rad Laboratories, Inc.). The membranes were blocked with 5% skim milk (AmericanBio) and probed with primary antibodies (1:1,000 dilution) overnight at 4°C. After washing with TBST buffer (25 mM Tris-HCl, 125 mM NaCl, 0.1% Tween 20), the membranes were incubated with IRDye®800CW- or IRDye®680-conjugated secondary antibodies (cat. nos. 926-32211 and 926-68071; 1:5,000 dilution; LI-COR Biosciences) for 1 h at room temperature. The results were visualized using an Odyssey Infrared Imager (LI-COR Biosciences). For loading controls, the membranes were stripped and reprobed for β-actin and/or tubulin. All primary antibodies used for western blotting are listed in Table I.
Ethics approval and consent to participate
Animal studiesThe University of Alabama at Birmingham (UAB) Institutional Animal Care and Use Committee (IACUC) (Birmingham, AL, USA), approved the work (approval no. IACUC-21928). Male C57B/6J mice (age, 4–6 weeks; weight, 22 g) were housed for 2–3 weeks to permit acclimatization and were monitored daily. The animals were housed at 25°C, under a 12-h light/dark cycle, with ad libitum access to food and water at 50% relative humidity. The ~22 g animals (n3) were exposed to 5% isoflurane for induction of anesthesia. Thereafter, the animals were maintained in 3% isoflurane until cardiac puncture and removal of the required blood volume (1.0 ml) was performed. To ensure euthanasia, the heart was removed. The long bone marrow of three animals was isolated by sterile dissection, and the cells from three animals were pooled.
Human studies
The studies were evaluated by the UAB Human Subjects Committee under IRB-300004457 ‘The Biology of Prostate Cancer Skeletal Metastases’. Given that no subject interactions were planned, and cell lines were to be purchased from commercial vendors, the studies were not deemed Human Subjects investigations and were therefore considered exempt.
Co-cultures of PC3-GFP cells with bone marrow cells and fluorescence-activated cell sorting (FACS) analyses
Murine primary bone marrow stromal cells were isolated by crushing the long bones from ~2-month-old male C57BL/6J mice. The cell clumps and debris were removed by filtering the cells using a 70-µm cell strainer (Thermo Fisher Scientific, Inc.). PC3 cells infected with GFP-labeled LANCL2 and PPARγ shRNA lentiviral vectors were placed into co-culture with confluent mouse primary bone marrow cells. Co-culture of PC3-GFP and bone marrow cells, with and without 100 µM ABA treatments were carried out for 3 days in an atmosphere containing 5% CO2 at 37°C. The proliferating PC3-GFP cells in the co-cultured (PC3-GFP/bone marrow) system were recovered using trypsin in PBS, and sorted using a BD FACS Melody™ Cell Sorter (BD Biosciences) to distinguish PC3 cells from murine bone marrow cells based upon GFP expression. To distinguish live versus dead cells or cellular debris, parallel cultures were used to establish gating parameters. A minimum of 20,000 events were recorded for each condition. The data were analyzed using BD FACSChorus™ 3.0 software (BD Biosciences). Cell proliferation percentages were established by comparing to control cells, and histograms were plotted using GraphPad Prism 5 (Dotmatics).
Statistical analyses
GraphPad Prism 5 was used for statistical analyses. All experiments were repeated at least three times and the results are presented as the mean ± SEM. For each data point, a two-tailed, unpaired, Student's t-test was performed to determine the significance of the differences between two groups, and one-way analysis of variance (ANOVA) followed by Bonferroni's post hoc test was performed to compare three or more groups. When analyzing multiple variables a two-way ANOVA analysis followed by Bonferroni's post hoc test was performed to determine significance. P<0.05 was considered to indicate a statistically significant difference.
Results
ABA treatment inhibits PCa cell viability
To determine the effects of ABA on PCa DTC viability, human and mouse PCa cells were exposed to increasing doses of ABA (0, 25, 50, 100 and 200 µM) for 72 h in culture medium. ABA significantly reduced the viability of the PCa cell lines in a dose-dependent manner (Fig. 1A-D). Specifically, ABA at concentrations of 100 and 200 µM decreased the viability of the PC3, C4-2B, DU145 and Myc-CaP cells by ~23, 33, 26 and 25%, and by ~24, 52, 36 and 48% compared with the control groups, respectively (Fig. 1A-D).
To determine if the inhibition of viability by ABA was reversible, a pulse-chase type of investigation was performed. Here, the metastatic PCa cells were treated with vehicle or ABA (100 µM) for 72 h. After the initial culture, the cells were washed, trypsinized and secondary cultures were established without additional ABA treatment. To account for the differences in viability that ABA induces, the secondary cultures were established with equal numbers of viable cells. As expected, the ABA-treated PC3 and C4-2B primary cultures had fewer cells relative to the vehicle-treated cultures (Fig. 1E and F). In the secondary cultures, the cells grew equally as well, regardless of their prior exposure in primary culture to ABA, thus demonstrating that the ABA-induced inhibition of viability was reversable (Fig. 1E and F). Notably, it was determined that ABA did not induce cell death of any of the PCa cells using a non-radioactive cytotoxicity assay to measure LDH release (Fig. 1G). These data collectively suggested that ABA induces reversible arrest of metastatic PCa cells.
ABA inhibits PCa cell viability by signaling through LANCL2 and PPARγ
To assess the molecular mechanisms by which ABA signaling influences PCa viability, stable KD of the ABA receptors LANCL2 and PPARγ were performed in metastatic PCa cell lines using shRNAs. To evaluate the efficiency of the KD, LANCL2 or PPARγ mRNA expression levels were confirmed by RT-qPCR (Fig. 2A-D) and protein expression levels were detected using western blotting (Fig. 2E-H). The targeting sequence #1 for LANCL2 and PPARγ were selected for all subsequent studies. After validation of the KD, the sensitivity of PC3 and C4-2B cells to ABA in vitro was assessed using cell viability assays. shRNA KD of LANCL2 or PPARγ conferred partial resistance to ABA in both PC3 and C4-2B cells (Fig. 2I and J).
As a surrogate assay for estimating in vivo tumorigenesis, anchorage-independent growth in soft-agar assays was performed to evaluate the effects of ABA and its signaling through its receptors (41,42). ABA significantly decreased the ability of PCa PC3 and C4-2B cells to form colonies in the NS groups; however, when LANCL2 or PPARγ expression was downregulated by shRNA, ABA treatment did not affect colony formation in soft agar (Fig. 3A-D). These observations suggested that LANCL2 or PPARγ are required for ABA signaling to induce anchorage-independent growth. Given the functional link between colony formation and cancer-stem cell (CSC) activities (43), these data also indicated that ABA, and its signaling though its receptors, may downregulate CSC proliferation; however, the impact on CSCs was not formally tested.
ABA inhibits PCa cell migration and invasion through LANCL2 and PPARγ receptors
Given the effects of ABA on viability and colony formation, the impact of ABA on PCa invasion was next evaluated. For these studies, the migration of cells treated with or without ABA was detected in response to FBS as a chemoattractant in Transwell plates. ABA decreased the invasion of PCa cells, whereas targeted KD of LANCL2 or PPARγ expression reduced the effects of ABA treatment on migration (Fig. 4A-D). To evaluate the effects of ABA and the respective receptor KD on invasion, Transwell plates covered in Matrigel were utilized. As with the migration assays, ABA decreased cell invasion in response to FBS, whereas KD of LANCL2 or PPARγ expression abrogated this effect (Fig. 4E-H).
To evaluate the mechanisms by which ABA affects invasion, the present study evaluated the mRNA expression levels of E-cadherin, Vimentin and ZEB2 by RT-qPCR. The results showed that ABA increased E-cadherin, and decreased Vimentin and ZEB2 expression levels in PCa cells in the NS group (Fig. S1). However, shRNA KD of LANCL2 or PPARγ receptors mitigated the effects of ABA on the majority of the cells. The exception being that the mRNA expression levels of Vimentin in ABA-treated shRNA PPARγ KDC4-2B cells were reduced compared with in those cells not treated with ABA, which could be due to a compensatory effect of the LANCL2 receptor (Fig. S1). Collectively, these results indicated that LANCL2 and PPARγ are necessary for the effects of ABA on PCa cell migration and invasion, and suggested that these receptors play a critical role in the metastatic attributes of PCa cells.
Inhibition of PCa cell viability in co-culture is enhanced by ABA
To explore the role that ABA and its receptors play in promoting dormancy in bone marrow, in vitro studies were performed. For these studies, PC3 cells were cultured in the presence of confluent monolayers of bone marrow stromal cells isolated from C57BL/6J mice. The co-cultures were grown for 3 days in the presence or absence of ABA. To distinguish the PC3 cells from the stromal layers, PC3-GFP cells were used. The viability of PC3-GFP cells was evaluated upon recovery of the cells from culture with trypsin by FACS probing for GFP. In co-culture, PC3-GFP viability was significantly decreased when ABA was included in the co-cultures (Fig. 5A, B and G). Notably, when the PCa cells with KD of LANCL2 or PPARγ were cultured with the stromal cells, they exhibited increased viability relative to the NS control group (Fig. 5C-G). As expected, the addition of exogenous ABA in the absence of LANCL2 or PPARγ expression led to little to no impact on cellular viability in the co-cultures (Fig. 5C-F). These results indicated that ABA inhibited PCa cell viability in co-culture, whereas when LANCL2 or PPARγ expression was reduced, either the PCa cells were no longer sensitive to ABA expressed by the bone marrow cells (44), or they become insensitive to another negative cell growth regulator produced by stomal cells which signals through LANCL2 or PPARγ.
ABA regulates transcription of p21, p27 and p16 through LANCL2 and PPARγ in PCa cells
The role of p21, p27 and p16 in cell cycle progression is well established. Activation of p21, p27 and p16 inhibits phosphorylation of CDK1 and CDK2 resulting in cell cycle arrest. To explore whether ABA signaling through LANCL2 and PPARγ impacts the expression of p21, p27 or p16, the PCa control or LANCL2 and PPARγ KD cells were treated with ABA and were examined by RT-qPCR analysis. Unexpectedly, when PPARγ expression was knocked down by shRNA ABA increased the expression levels of LANCL2 (Fig. 6A and F), and had variable impact on PPARγ expression when LANCL2 expression was knocked down by shRNA (Fig. 6B and G), thus suggesting that the expression of these ABA receptors establish a feedback loop in PC3 (Fig. 6A and B) and C4-2B cells (Fig. 6F and G). Notably, ABA induced the mRNA expression levels of p21, p27 and p16 in both PC3 (Fig. 6C-E) and C4-2B (Fig. 6H-J) cells. In the predominance of the studies, KD of LANCL2 and PPARγ by shRNA markedly abrogated the effect of ABA on the mRNA expression levels of p21, p27 and p16 (Fig. 6C-E and H-J). Collectively, these results suggested that ABA induces transcriptional regulation of several cell cycle inhibitor proteins in PCa cells through LANCL2 and PPARγ signaling.
ABA induces PCa dormancy through LANCL2 and PPARγ and p38MAPK pathway activation
Phosphorylation of p38MAPK is one of the major signaling pathways involved in the induction of dormancy by molecules that inhibit cell proliferation, including TGFβ, BMPs and GAS6 (15,17). Notably, activation of p38MAPK signaling by phosphorylation, and enhanced expression of the cell cycle inhibitors p21 and p27, are considered as dormancy markers in cancer cells (15,19,45). Therefore, the present study evaluated the effects of ABA signaling on p38MAPK activation. Treatment of the PCa cells with ABA resulted in phosphorylation 38MAPK (Fig. 7A, B, D and E), as well as stimulation of NR2F1 expression, a dormancy marker (Fig. 7A, C, D and F). ABA also enhanced the expression levels of the downstream targets of p38MAPK, including p21 and p27 (Fig. 7G and H). As expected, KD of the ABA receptors LANCL2 and PPARγ negated the ABA-induced effects on p21, p27 and NR2F1 expression (Fig. 7A-H). Collectively, these findings indicated that ABA affects the viability arrest of PCa cells through LANCL2 and PPARγ receptors, in part by signaling through p38MAPK.
Discussion
PCa is a heterogenous disease, which has variable clinical outcomes ranging from early-stage, curable, advanced and terminal disease. Although the majority of men are cured soon after diagnosis at an early stage, a subset of men develop recurrent disease, or present with de novo metastatic disease (46). The bone is a common site for PCa metastasis (47), where metastatic cells that have escaped the prostate early in disease are likely to have entered a dormant state for years or even decades. Notably, ~90% of patients who succumb to metastatic PCa have some level of bone involvement (48). Several new therapeutic strategies, including androgen receptor-targeted therapies, chemotherapy, poly ADP-ribose polymerase inhibitors, sipuleucel-T and radium-223, have been approved for the treatment of patients with advanced PCa and bone metastasis. However, this disease still represents the second most frequent cause of cancer-related death in men in the United States (49). The challenges in treating bone metastases at the advanced stage include a high level of genomic heterogeneity (50), a pro-immunosuppressive environment (51) and several emerging mechanisms of androgen independence (52). Therefore, new therapeutics are urgently needed.
ABA is known to regulate cell responses and biological processes in plants, mammals and other organisms (53,54). ABA is a key hormone that promotes seed dormancy during development in plants and after seed dispersal (55). ABA controls the induction of dormancy, and ultimately the release and germination of seeds in response to environmental signals (55). ABA has also been recognized as an endogenous hormone in humans (29). It has a role in the inflammatory response, as well as in immunoregulatory activities, antioxidant properties and the maintenance of glycemic control in pre-clinical models of diabetes and inflammatory diseases (56,57). Despite the advances in the understanding of the role that ABA plays in several physiological processes, the role of ABA in tumor dormancy remains to be elucidated.
The present study demonstrated that ABA suppressed the viability of four distinct PCa cell lines (PC3, C4-2B, DU-145 and Myc-CaP) in a dose-dependent manner; however, exposure to ABA was not in and of itself cytotoxic to the cancer cells. Notably, ABA-induced inhibition of PCa cell viability was reversible, as cells which were removed from ABA treatment were able to resume their proliferation. There are at least two receptors that are responsible for transducing ABA signaling; LANCL2 and PPARγ. To explore how ABA induced proliferative quiescence, targeted KD of LANCL2 and PPARγ was established using shRNAs in PC3 and C4-2B cell lines. In almost every instance where ABA signaling was examined, including viability, migration, invasion and downstream signaling, reduced expression of LANCL2 and PPARγ abrogated the impact of ABA on the cancer cells. Furthermore, it was observed that ABA inhibited the formation of PCa colonies in soft agar, whereas LANCL2 and PPARγ KD alone did not. Mechanistically, ABA treatment induced p21, p27 and p16 mRNA and protein expression, whereas LANCL2 and PPARγ shRNA KD did not significantly inhibit either p21, p27 or p16 mRNA or protein expression relative to the controls. These data suggested that ABA may inhibit viability through inducing the expression levels of p21, p27 and p16, but when the expression of ABA receptors is reduced, there is little impact on p21, p27 and p16 expression.
LANCL2 and PPARγ are known to mediate a range of physiological responses of ABA in a number of systems, including granulocytes and rat insulinoma cells (31,58). ABA signaling in mammalian immune cells, keratinocytes and pancreatic cells requires LANCL2 expression, as small interfering RNA targeting of LANCL2 has been shown to abrogate cellular responses (23,26,58). Several studies have reported that activation of PPARγ by agonists markedly reduces cell proliferation, including in hepatocellular carcinoma (59), PCa (60) and gastric cancer (61). Other studies have demonstrated that PPARγ agonists promote terminal differentiation, inhibit cell proliferation and increase the apoptosis of human cancer cell lines, as well as tumor growth in animal models (33,62). Notably, in some cases, PPARγ agonists have demonstrated modest efficacy as chemopreventives in clinical trials (33,62). It is not surprising that clinical studies have reported that the overall survival of patients with colorectal cancer is better when PPARγ expression is detectable in the primary tumors (63). In part, this may be due to the findings that PPARγ modulates the expression of different cell cycle regulators, including decreasing the expression of cyclin D1 (64), and increasing the expression of the CDK inhibitors p21 (65) and p27 (66).
The present study reported on notable findings in co-culture studies of PCa with bone marrow stromal cells. In the presence of marrow stroma, ABA is known to inhibit PCa cell viability (44). However, when PCa cells with reduced expression of LANCL2 and PPARγ were grown on the bone marrow stromal cells, the cells proliferated at the same rate as the control cells. These results suggested that PCa cells with decreased expression of LANCL2 and PPARγ may no longer be sensitive to ABA produced by the mesenchymal stromal cells, as shown in a previous study (67), or are no longer sensitive to growth inhibitors secreted by the stomal cells, which also signal through LANCL2 and PPARγ. Very little is known as to what activates LANCL2. In glioma it has been demonstrated that LANCL2 expression may be co-regulated by epidermal growth factor receptor (68). Activation of PPARγ can inhibit cell proliferation, induce differentiation and promote metastasis in PCa cells (60). In other contexts, PPARγ is activated by fatty acids and derivatives including polyunsaturated fatty acids, derivatives of arachidonic acid and prostaglandins (69). However, what other negative regulators of PCa cell viability are produced by bone marrow stromal cells, which signal through LANCL2 and PPARγ remain unclear.
In the PCa cell lines tested there appears to be some diversity in signaling in response to ABA through PPARγ and LANCL2. For example, p27 activity was enhanced by ABA treatment in both cell lines, but when LANCL2 was knocked down by shRNA, the p27 response was altered in the C4-2B and PC3 cells, although the response was in the opposite direction. Currently, there is no specific explanation as to the basis of the differences in these observations. A reasonable possibility is that the cells have other receptors for ABA in addition to LANCL2 and PPARγ. In other studies, LANCL1 and CD38 have been suggested to serve as receptors or co-stimulatory molecules for LANCL2 (58,70). Unfortunately, we were not able to achieve functional reductions of CD38 in the cell lines used in the present study (C4-2B and PC3); therefore, it is possible that other players participate in the signaling. Furthermore, there are known ABA inhibitors in plants, including gibberellins, which can in some cases serve as allergens and of which derivatives may have anti-neoplastic activities (71–77). Whether equivalent proteins function in human systems remains unclear. However, in preliminary studies we did not observe any significant impact of gibberellin on the cell cycle of metastatic cancer cell lines (Jung et al, unpublished data). Further studies are needed to determine the basis of the variations in cell cycle inhibitors.
To further understand the mechanisms by which ABA signals through LANCL2 and PPARγ to induce dormancy, western blot analysis was performed to detect p38 phosphorylation in PCa cells. It was revealed that KD of LANCL2 and PPARγ inhibited the activation of p38 by ABA. In addition, NR2F1 has been reported to act as a master regulator of tumor cell dormancy (78). By binding to DNA and recruiting coactivator or corepressor complexes, NR2F1 is able to serve as a cofactor to other nuclear receptors. Notably, NR2F1 expression, one of the best known dormancy markers, has been reported to be directly regulated by ABA (78), and its expression in this previous study was associated with p38 signaling (78). Activation of p38 signaling induces expression of the cell cycle inhibitors p21 and p27, also considered as dormancy markers in cancer cells (15,19,45). These findings make sense given the proposed hypothesis that the ratio between the p38 MAPK stress response signaling pathway and ERK guides cancer cells into a proliferative or dormant state (79).
The regulation of cell proliferation in the bone marrow is a critical function of the marrow microenvironment to protect and promote proliferation of hematopoietic and mesenchymal stem cells, and its integration across an organism is critical to the survival of mammals. The present study assessed the role of ABA and its signaling pathways in regulating cell proliferation/quiescence of cells that do not belong in the marrow, specifically cells of epithelial origin. Given that there are a number of pathways that regulate the proliferation and dormancy of stem and progenitor cells in the marrow, it is not surprising that there are multiple pathways that can regulate stem cell proliferation (16,80–82). That tumor cells exploit the niches used by the bone marrow to regulate stem and progenitor cells is not surprising and makes sense from an ecological perspective (80). For example, osteoblasts that participate in forming hematopoietic stem cell (HSC) niches produce GAS6, which not only regulates HSC function, but limits the proliferation of PCa cells in vitro (83) and in vivo (84–86). Other studies have shown that GAS6 and its receptor AXL are required for TGFβ2-mediated cell proliferation suppression in PCa, where AXL positively regulates the expression of TGFβ and TGFβ receptor 2 (18). In the current study, it was demonstrated that ABA in co-culture of PCa cells with murine bone marrow significantly inhibited PCa cell viability. However, LANCL2 and PPARγ KD abrogated the response to ABA. Together, these data suggest the following model: ABA signaling through LANCL2 or PPARγ receptors activates signaling via p38MAPK, NR2F1, p27, p16 and p21, which ultimately leads to dormancy (Fig. 8). These results suggest that interference with ABA signaling under the appropriate conditions, or perhaps even from dietary sources such as fruits and vegetables, may prove beneficial in treating men with prostate cancer, by either keeping DTCs as dormant cells or by interfering with ABA signaling together with chemotherapy to selectively kill DTCs.
In conclusion, the present findings suggested that ABA may induce cell cycle arrest in PCa. ABA does so by signaling through LANCL2 and PPARγ, which activate p38MAPK and the CDK inhibitors, p21, p16 and p27, and NR2F1 resulting in PCa cell dormancy. Further investigation into how the ABA signaling pathway results in dormancy may reveal novel opportunities for eradicating dormant cancer cells and/or keeping them in a perpetual dormant state, thus preventing PCa metastasis.
Supplementary Material
Supporting Data
Acknowledgements
The authors would like to express their gratitude to Professor Amjad Javed (University of Alabama at Birmingham) for generously providing the mice used for isolating primary bone marrow and conducting the co-culture experiments with prostate cancer cells.
Funding
This work was supported by the NIH/NCI (grant no. 3P01CA093900-018), the Prostate Cancer Foundation (2014 Challenge Award) and the Department of Defense (grant no. PC140665).
Availability of data and materials
The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.
Authors' contributions
KRP, YJ and RST conceived or designed the study. KRP collected the data. KRP and RST analyzed and interpreted the data. KRP drafted the article. YJ and RST critically revised the article. KRP and RST confirm the authenticity of all the raw data. All authors read and approved the final manuscript.
Ethics approval and consent to participate
The University of Alabama at Birmingham Institutional Animal Care and Use Committee (IACUC) approved the present study (approval no. IACUC-21928).
Patient consent for publication
Not applicable.
Competing interests
The authors declare that they have no competing interests.
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