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Epidemiology and Prevention |
and 
Agonists Differentially Alter Tumor Differentiation and Progression during Mammary Carcinogenesis
1 Department of Oncology, Georgetown University, Washington, District of Columbia; 2 Department of Medicinal Chemistry and Pharmacognosy, University of Illinois at Chicago, Chicago, Illinois; and 3 Division of Chemoprevention, National Cancer Institute, Bethesda, Maryland
Requests for reprints: Robert I. Glazer, Department of Oncology, Georgetown University School of Medicine, Room W318, Research Building, 3970 Reservoir Road, Northwest, Washington, DC 20057. Phone: 202-687-8324; Fax: 202-687-7505; E-mail: glazerr{at}georgetown.edu.
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Abstract |
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and PPAR
agonists in a progestin- and carcinogen-induced mouse mammary tumorigenesis model. Animals treated with the PPAR agonist GW7845 exhibited a moderate delay in tumor formation. In contrast, animals treated with the PPAR agonist GW501516 showed accelerated tumor formation. Significantly, tumors from GW7845-treated mice were predominantly ductal adenocarcinomas, whereas tumors from GW501516-treated animals were adenosquamous and squamous cell carcinomas. Gene expression analysis of tumors arising from GW7845- and GW501516-treated mice identified expression profiles that were distinct from each other and from untreated control tumors of the same histopathology. Only tumors from mice treated with the PPAR agonist expressed estrogen receptor-
in luminal transit cells, suggesting increased ductal progenitor cell expansion. Tumors from mice treated with the PPAR agonist exhibited increased PPAR levels and activated 3-phosphoinositide–dependent protein kinase-1 (PDK1), which co-associated, suggesting a link between the known oncogenic activity of PDK1 in mammary epithelium and PPAR activation. These results indicate that PPAR and PPAR agonists produce diverse, yet profound effects on mammary tumorigenesis that give rise to distinctive histopathologic patterns of tumor differentiation and tumor development. |
Introduction |
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The role of the PPAR family has been controversial in the field of cancer cause and prevention. In colon cancer models, such as the APCMin mouse, PPAR expression resides downstream of ß-catenin activation (5) and influences adenoma proliferation (6) and tumor development (7). These activities may be, in part, related to activation of the downstream PPAR gene target, 3-phosphoinositide-dependent protein kinase-1 (PDK1; ref. 8), which itself is oncogenic when expressed in mammary epithelial cells (9, 10), as well as to activation of proliferation and cell survival pathways (8, 11). PPAR function may also be associated with ligand-independent actions as shown by the paradoxical finding that PPAR agonist GW501516 produced a similar anti-inflammatory phenotype in macrophages as in cells nullizygous for PPAR through its interaction with the BCL-6 corepressor (12).
In some instances, PPAR agonists have been associated with tumor promotion as shown by the ability of the thiazolidenedione class of agonists, troglitazone, and rosiglitazone to promote colon polyp formation (13, 14). Other studies suggest a tissue-specific role for PPAR activation in tumorigenesis. Mice expressing a constitutively active PPAR transgene did not exhibit a tumorigenic phenotype, but markedly accelerated mammary tumorigenesis when crossed with mouse mammary tumor virus–polyoma middle T-antigen transgenic mice (15). However, mice nullizygous for PPAR did not differ in their susceptibility to prostate carcinogenesis initiated by the probasin-SV40 T-antigen transgene (16). With respect to human disease, inactivating polymorphisms in PPAR have been reported in sporadic colon cancers (17, 18) as well as findings to the contrary (19). As yet untested as a susceptibility factor is the provocative finding that a single point mutation may alter PPAR selectivity as shown by the ability of PPARM417V to function as PPAR (20).
In contradistinction to the aforementioned studies, PPAR agonists have been reported to possess chemopreventive activity. PPAR agonist GW7845 and PPAR agonist WY14643 reduced 7,12-dimethylbenz(a)anthracene (DMBA)–mediated mammary carcinogenesis in rats (21, 22) and troglitazone inhibited DMBA-induced preneoplastic lesions in mammary gland organ culture (23). Although it is presumed that the effects of PPAR agonists are receptor mediated, there is evidence to suggest that troglitazone can act as a partial agonist (24), exhibit concentration-dependent biphasic activity (25), and elicit nonspecific antioxidant effects (26). This is consistent with reports of thiazolidenedione PPAR agonists acting through receptor-independent mechanisms (27, 28).
In the present study, we wished to determine whether the highly potent and selective PPAR agonist GW7845 and PPAR agonist GW501516 possessed chemopreventive activity in a mouse mammary carcinogenesis model. Our results show the unexpected findings that at bioequivalent doses, GW501516 acted as a tumor promoter, whereas GW7845 produced a delay in tumor formation. In addition, GW7845-treated mice presented with ductal adenocarcinomas, in contrast to GW501516 treatment where adenosquamous and squamous cell carcinomas predominated. These findings suggest that PPAR and PPAR agonists not only affect tumor formation in divergent ways, but that they can act as lineage fate determinants influencing tumor differentiation and development.
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Materials and Methods |
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Mammary carcinogenesis. Six-week-old female FVB mice received an injection of 15 mg medroxyprogesterone acetate (Depo-Provera, Pharmacia & Upjohn, Kalamazoo, MI) s.c. 1 week before administration of 1 mg DMBA in 0.1 mL cottonseed oil each week for 4 consecutive weeks. Each experimental group consisted of 10 to 11 mice, and animals were placed on a control diet of Purina rodent chow 5001 or diets containing either 0.005% GW7845 or 0.005% GW501516 1 day after the last DMBA dose, and maintained on the diets ad libitum for the duration of the study.
Histopathology. Tumor samples were dissected free of necrotic and connective tissue and preserved in formalin. Paraffin blocks were prepared for H&E staining and immunohistochemistry by the Histopathology and Tissue Shared Resource, Lombardi Comprehensive Cancer Center, Georgetown University. All samples were evaluated in a blinded fashion by the pathologist (Dr. Robert G. Russell). Tumors were classified using the histologic nomenclature recommended by Cardiff et al. (30), and arranged into three groups: group I, adenocarcinoma including acinar and solid lobular arrangements; group II, squamous cell carcinoma and adenosquamous carcinoma; and group III, myoepithelial carcinoma and undifferentiated carcinoma.
Immunohistochemistry. Sections of tumors were cut at 5 µm and deparaffinized. Antigen retrieval was conducted for 25 minutes using Citra Plus (Biogenex HK080-9k) in a steamer.
Sections were then blocked with 10% normal goat serum for 30 minutes at room temperature and subsequently incubated with an anti-estrogen receptor- (ER-) antibody (sc-562, Santa Cruz Biotechnology, Santa Cruz, CA) diluted 1:200 (1 µg/mL) overnight at 4°C in a humidified chamber. Sections were washed in PBS and incubated with a polymer-labeled secondary antibody for 30 minutes at room temperature, washed in PBS, and reactions were visualized with 3,3'-diaminobenzidine as substrate (DAKO Envision Plus Staining System, DAKO, Carpenteria, CA). Sections were counterstained with Harris hematoxylin. Appropriate controls for immunostaining included a breast carcinoma as a positive control and replacing the primary antibody with normal IgG as a negative control.
Gene microarray. Solid lobular adenocarcinomas and squamous cell carcinomas with no evidence of other cellular elements were used for gene array analysis. Mammary gland tumors were removed and kept in RNAlater (Ambion, Austin, TX) at –20°C until RNA was prepared using TRIzol (Invitrogen, Carlsbad, CA). RNA quality was monitored by electrophoresis in 1% denaturing agarose gels, and purity was assessed by a A260/A280 ratio of 
1.9. cRNA synthesis was done using the Affymetrix protocol with minor modification. First- and second-strand cDNA synthesis was initiated with 12 µg total RNA and purified with DNAclear (Ambion). In vitro transcription was done with MEGAscript T7 (Ambion) using biotin-11-CTP (Enzo Diagnostics, Farmingdale, NY), and biotin-labeled cRNA was purified with MEGAclear (Ambion). Twenty micrograms of biotin-labeled cRNA were fragmented at 94°C for 35 minutes and used for hybridization overnight on an Affymetrix MG-U74Av2 GeneChip representing 12,473 open reading frames by the Macromolecular Analysis Shared Resource, Lombardi Comprehensive Cancer Center. The processed chips were scanned using an Agilent Gene Array scanner and grid alignment and raw data generation were done using Affymetrix GeneChip 5.0 software. For quality control, oligo-B2 was hybridized to analyze the checkerboard pattern in each corner of the chip and bioB, bioC, and bioD probes were added to each sample with varying concentrations to standardize the hybridization, staining, and washing procedures. Raw data representing the average difference in hybridization intensity between oligonucleotides that perfectly match the transcript sequence and oligonucleotides containing single bp mismatches were measured. A noise value (Q) based on the variance of low-intensity probe cells was used to calculate a minimum threshold for each GeneChip. Data generated after scanning was subjected to comparison analysis to select change calls at 100% increase or decrease compared with control for each gene. Gene array analysis was further refined by evaluating differences between paired tumor samples and ranking changes by their log2 ratio. Only differences in signal ratio >log2 2.0 and <log2 –2.0 were ranked, and genes with low constitutive expression (signal intensity <500) in both sample pairs were excluded. This allowed culling the data set to genes with high constitutive expression and minimizing variability due to small changes in expression. Each cRNA was prepared from equal amounts of RNA pooled from three individual tumors. Each hybridization was repeated twice.
Quantitative reverse transcription-PCR. Total RNA was extracted from mammary gland tumors as described above for gene microarray analysis. RNA (2 µg) was predigested with DNase I (Invitrogen) for 15 minutes and initiated for cDNA synthesis with Superscript II RNase H– reverse transcriptase (Invitrogen) and random primers following the manufacturer's protocol. Quantitative reverse transcription-PCR (qRT-PCR) was done in triplicate in an ABI-Prism 7700 sequencing instrument (Applied Biosystems, Foster City, CA) using SYBR green I detection as described (31). Briefly, the 20 µL PCR reaction contained 1x Platinum SYBR Green qPCR SuperMix UDG (Invitrogen), 300 nmol/L of each primer, Rox-Dye, and the cDNA sample. qRT-PCR conditions were 2 minutes at 50°C and 10 minutes at 95°C, followed by 45 cycles of 15 seconds denaturation at 95°C, and 30 seconds annealing/extension at 60°C. The fluorescent signal intensities were recorded during the PCR reaction and analyzed using SDS v1.91 software. Dissociation curves of the amplicons were generated after each run to ensure that increased fluorescent intensity was not due to primer dimerization. The increase in fluorescent signal was associated with exponential formation of PCR product during the linear log phase. The threshold cycle (CT) value is the cycle at which a significant increase in the reaction product is first detected. The higher the initial amount of cDNA, the sooner accumulated product is detected in the PCR process, and the lower the CT value. The expression of each target gene was normalized to the expression of ß-actin and is presented as the ratio of the target gene to ß-actin gene calculated by 2-
Ct, where Ct=CtTarget–Ctß-actin. The following pairs of primers were used for qRT-PCR: adipsin: forward 5'-CAA TCA TGA ACC GGA CAA CCT G-3', reverse 5'-CGC GAG AGC CCC ACG TAA CCA CA-3'; proliferin: forward 5'-ATG CTC CCT TCT TCG ATT CAA C-3', reverse 5'-CTC TGA GCC CAG ACA CGT TAG A-3'; receptor activator of nuclear factor-
B ligand (RANKL): forward 5'-TTT GCA CAC CTC ACC ATC AA-3', reverse 5'-GAA AGC AAA TGT TGG CGT ACA G-3'; IGFBP-5: forward 5'-TAC GGC GAG CAA ACC AAG ATA GAG-3', reverse 5'-GTC GAC GGA AAT GCG AGT GTG CT-3'; MMP-9: forward 5'-GGC GTG TCT GGA GAT TCG ATT TGA-3', reverse 5'-GGA AAC TCA CAC GCC AGA AGA A-3'.
Statistical analysis. Tumor-free intervals were evaluated by Kaplan-Meier analysis, and tumor formation by the Wilcoxon rank log test. Differences were considered statistically significant at P < 0.05. Statistical analyses were contributed by Drs. Ying Zhang and Ed Gehan (Biostatistics Shared Resource, Lombardi Comprehensive Cancer Center).
Western blot. Tumor samples were frozen immediately in dry ice and stored at –80°C. Samples were homogenized with radioimmunoprecipitation assay buffer (RIPA: 1x PBS, 1% NP40, 0.5% sodium deoxycholate, 1% SDS, 1 mmol/L sodium orthovanadate, 0.1 mg/mL phenylmethylsulfonyl fluoride, protease inhibitor cocktail, Boehringer-Mannheim, Mannheim, Germany), allowed to remain on ice for 60 minutes, and then centrifuged at 12,000 x g for 15 minutes. Equal amounts of protein were separated in 10% polyacrylamide gels by SDS-PAGE. Gels were blotted onto nitrocellulose (Optitran, Schleicher & Schuell, Keene, NH) and used for immunoblotting as described (32) using polyclonal antibodies against PPAR antibody (sc-1986, Santa Cruz Biotechnology), PDK1 (06-637, Upstate Biotechnology, Lake Placid, NY), and PDK1pSer241 (3061, Cell Signaling Technology, Beverly, MA).
Immunoprecipitation. Lysates were prepared by homogenization of a squamous cell carcinoma using modified RIPA buffer [50 mmol/L Tris (pH 7.4), 0.5% NP40, 0.25% Na-deoxycholate, 125 mmol/L NaCl, 1 mmol/L EDTA, 1 mmol/L EGTA, 1 mmol/L Na3VO4, and protease inhibitor cocktail (Boehringer-Mannheim)]. For 293T cells, lysates were prepared without homogenization. Lysates were kept on ice for 1 hour, sonicated for 10 seconds, and centrifuged at 12,000 x g for 15 minutes at 4°C, and the supernatant removed for immunoprecipitation. Tumor lysates containing 1.5 mg protein or 293T cell lysates containing 0.5 mg protein were precleared for 30 minutes with 30 µL Protein G Plus/Protein A agarose (Oncogene Research Products, Cambridge, MA) and incubated at 4°C with the appropriate antibodies or normal IgG preadsorbed to Protein G Plus/Protein A agarose. Immune complexes were collected after incubation overnight at 4°C, washed once with lysis buffer, and washed thrice with PBS. Protein was eluted by boiling in 50 µL 2x SDS sample buffer [125 mmol/L Tris (pH 6.8), 10% 2-mercaptoethanol, 4% SDS, 20% glycerol], and eluted proteins were analyzed by SDS-PAGE and Western blotting using the appropriate antibodies.
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Results |
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agonist GW501516 exhibited tumor formation within 2 months (Fig. 1) but no significant change in tumor multiplicity (1.3 tumors per mouse). In contrast, animals treated with PPAR agonist GW7845 showed reduced tumor formation beginning 4 months after initiating the PPAR regimen (Fig. 1) and no change in tumor multiplicity.
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in what appeared to be early or late ductal transit cells (37), whereas adenocarcinomas from control mice and GW501516-treated mice were largely ER negative.
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/ß, proliferin 2/3, and PAI-II expression. Gene expression changes associated with GW7845 treatment were reduced TCA4, FGFR1, TIMP-3, Twist, IGFBP-3 and placental growth factor and increased MMP-9 and RANKL. Several of these changes were corroborated by qRT-PCR (Fig. 4). Among the changes associated with GW501516 treatment were increased proliferin-2 and proliferin-3 expression and pronounced reduction in mitochondrial uncoupling protein UCP as well as reduced IGFBP-5 and adipsin (Fig. 4). GW7845-treated tumors exhibited higher MMP-9 and RANKL and reduced TCA4 expression.
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and the PPAR target gene, PDK1 (Fig. 5; ref. 8). Treatment with GW501516 increased PPAR expression in tumors that was accompanied by increased PDK1 activity as assessed by PDK1pSer241 reactivity (Fig. 5A). Tumor lysates were then prepared from GW501516-treated animals and immunoprecipation done with a PPAR antibody (Fig. 5B). Western blotting revealed that PDK1 co-associated with PPAR. Reprobing the blot with a PPAR antibody was not possible due to interference by the IgG heavy chain of similar molecular weight. Co-association of PDK1 and PPAR was confirmed by transient expression of both genes in 293 T cells using Flag-tagged PPAR (Fig. 5C). PDK1 and PPAR associated with a high degree of efficacy as noted by the percentage of input Flag-PPAR in the immunoprecipitate.
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Discussion |
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and PPAR agonists on tumor latency. PPAR agonist GW501516 increased tumor development subsequent to carcinogen administration and, hence, fit the operational definition of a tumor promoter. In contrast, a bioequivalent regimen of PPAR agonist GW7845 exhibited a late chemoprotective effect as evidenced by the 2-month delay to tumor formation in all animals, a finding consistent with previous studies in a rat DMBA carcinogenesis model (22). Although PPAR activation has previously been associated with a proliferative or tumorigenic response in the colon (6, 7), our study is apparently the first to report that a PPAR agonist has tumor promoting activity. There has been much contention about whether PPAR agonists are chemoprotective or tumor promoting. PPAR and PPAR agonists have been reported to inhibit tumor cell growth and to increase cell survival (38–41). In interpreting the latter results, it is important to note that PPAR agonists of the thiazolidenedione class can act as partial agonists (24), exhibit dose-dependent biphasic activity (25), and act by receptor-independent mechanisms (27, 28, 42, 43). This is particularly true for analogues, such as troglitazone, which contains a vitamin E moiety that can elicit nonspecific antioxidant effects (26). Nonspecific actions have not been reported for the more selective and chemically unrelated PPAR agonists GW7845 and GW501516 used in the present study.
The second salient finding in our study was the pronounced and selective increase in adenocarcinoma formation by GW7845. This was associated with increased ER expression in what appeared to be transit or progenitor cells (37), suggesting that PPAR activation may promote differentiation along a ductal epithelial lineage. Notwithstanding nonspecific actions, there is precedent for PPAR agonists to influence tumor differentiation. Troglitazone promoted a more differentiated phenotype in breast carcinoma (44) and colon carcinoma (45) cells and in squamous metaplasia (46), and pioglitazone induced terminal differentiation of liposarcoma cells (47). PPAR overexpression in hepatocytes induced a phenotypic switch from activated to quiescent stellate cells and blocked myoepithelial differentiation (48). However, the gene expression profile in tumors from GW7845-treated animals revealed several biomarker changes that reflected both a more differentiated and a more invasive phenotype. Increased MMP-9 and reduced TIMP-3, TCA4, Twist, IGFBP3, and INK4a (49–52) expression would be expected to be indicative of proliferation and invasion, but increased expression of ER and lactogenic genes, such as casein and whey acidic protein, suggest a more differentiated, less tumorigenic phenotype as evidenced by the increase in tumor latency. It is uncertain if the changes in tumor histopathology elicited by GW7845 are direct or indirect because tumor development is influenced by the secretion of adipokines, such as IGFBP-2, MMP-1, and IGF-2 (53). The abundance of adipose tissue in the mouse mammary gland, which contains plentiful levels of PPAR (54), also suggests that the adipogenic effects of PPAR and PPAR activation (55, 56) may have influenced tumor development. Although PPAR agonists are also known to elicit proangiogenic (57–60) and antiangiogenic (61) effects, we did not observe a gene expression profile for GW7845 treatment indicative of such an action as reported for rosiglitazone (62, 63).
In contrast to the PPAR agonist, GW501516 induced tumor promotion that was associated exclusively with adenosquamous and squamous cell differentiation. This could have resulted from direct activation of PPAR or to redirection of lineage fate specificity resulting in transdifferentiation. Nevertheless, the pattern of gene expression for GW501516 treatment indicated several changes indicative of tumor-promoting activity. Proliferin-2 and proliferin-3, members of the prolactin gene family and ligands for the mannose-6-phosphate/IGF-II receptor (64), have been associated with proliferation (65), angiogenesis (66), and transformation (67). Reduced IGFBP5 and TCA4 and increased Notch1, PAI-2, and IL-1/ß gene expression (51, 68–71) have been linked to increased tumorigenesis and invasion. This profile correlates closely with the observed tumor promoting action of GW501516. Perhaps of more interest is the increase in activated PDK1pSer241 and PPAR observed in GW501516-treated tumors, and their co-association in tumor lysates. PDK1 is transforming in mammary epithelial cells due to activation of ß-catenin signaling (9, 10), and is a gene target of PPAR transactivation (8). This is the first demonstration of this protein-protein interaction, and suggests the possibility that PDK1 and PPAR may form an autoactivating oncogenic cycle that may contribute to the enhanced tumor promoter activity of GW501516.
In summary, our results show that a PPAR agonist accelerates mammary carcinogenesis, which contrasts the tumor-delaying action of a PPAR agonist. These biological activities could be distinguished on the basis of tumor histopathology and gene expression for the respective drugs. These studies suggest that these new classes of PPAR agonists warrant further evaluation, as well as caution in the context of hormone-dependent breast cancer.
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Acknowledgments |
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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.
We thank Dr. Clint Grubbs (University of Alabama, Tuscaloosa, AL) for preparation of the diets, Drs. Ying Zhang and Ed Gehan (Georgetown University, Washington, DC) for statistical analyses, and Carlos Benitez and Aaron Foxworth (Georgetown University Animal Research Resource, Washington, DC) for tumor measurements and diligent animal care. We thank Dr. Paul Grimaldi (University of Nice, France) for providing the PPAR cDNA.
Received 11/ 8/04. Revised 1/11/05. Accepted 2/14/05.
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References |
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), 0.005% GW7845 (
), or 0.005% GW501516 (
). Differences in tumor formation by GW501516 and GW7845 were statistically significant (P < 0.05) by the Wilcoxon rank test.
