ER and PR in Renomedullary Interstitial Cells During Syrian Hamster ...

0013-7227/01/$03.00/0 Printed in U.S.A.

Endocrinology 142(9):4006 – 4014 Copyright © 2001 by The Endocrine Society

ER and PR in Renomedullary Interstitial Cells During Syrian Hamster Estrogen-Induced Tumorigenesis: Evidence for Receptor-Mediated Oncogenesis JONATHAN J. LI, SARAVUT JOHN WEROHA, MARILYN F. DAVIS, OSSAMA TAWFIK, XIAOYING HOU, AND SARA ANTONIA LI Hormonal Carcinogenesis Laboratory, Division of Etiology and Prevention of Hormonal Cancers, Kansas Cancer Institute, and Departments of Pharmacology, Toxicology, and Therapeutics (J.J.L., S.J.W., S.A.L.); Preventive Medicine (J.J.L.); and Pathology and Laboratory Medicine (O.T., M.F.D.), University of Kansas Medical Center, Kansas City, Kansas 66160-7312; and Department of Hematology/Oncology, University of Alabama (X.H.), Birmingham, Alabama 35294 The estrogen-induced and -dependent Syrian hamster renal tumor is the most intensively studied model in estrogen carcinogenesis. Yet, it remains confounding that the kidney of this species behaves as an estrogen target tissue. As both reproductive and urinary systems arise from the same germinal ridge, we propose that some of the germinal cells, normally destined for the uterus, migrate and establish themselves in the renal corticomedullary region in this hamster strain. These ectopically located germinal cells remain dormant unless exposed to estrogen. Supporting this contention, a subset of renal interstitial cells, primarily located in the corticomedullary region, express PR after only 2 wk and ER␣ after 1.5–3.0 months of estrogen treatment. As treatment continues, groups

E

STROGENS HAVE BEEN associated with the causation of a variety of prevalent human cancers, including breast, endometrium, and possibly the ovary (1, 2). Certain studies have also implicated estrogens in the etiology of prostate cancer (3). In the estrogen-induced hamster kidney model, fundamental questions regarding the roles of estrogenic hormones in tumorigenic processes in target tissues can be properly addressed, because no other additional carcinogenic agent is involved. Multiple bilateral renal tumors are specifically induced by either steroid or stilbene estrogens, with an essentially 100% incidence in both intact and castrated male hamsters (4, 5). No spontaneous tumors have been reported in our hamster colony or in other larger colonies at this organ site (6, 7). Prevention of estrogen-induced renal tumorigenesis occurs by the concomitant administration of hormonal estrogen antagonists, such as antiestrogens, androgens, progestins, and, paradoxically, ethinyl estradiol (8 –10). These data provide compelling evidence that the renal tumor induced by estrogens is a hormone-mediated process. There has been considerable controversy regarding the cell of origin, and the developmental processes involved in the estrogen-induced tumorigenesis of the hamster kidney (11– 13). We have provided evidence that the cells of origin are interstitial stem cells found most abundantly in the corticoAbbreviations: DES, Diethylstilbestrol; NIC, nuclear image cytometry; PCNA, proliferating cell nuclear antigen.

of cells of the renal interstitium and small and large renal tumors show ER␣ⴙ and PRⴙ staining. Although ER␣ and PR isoform profiles in estrogen-treated hamster kidneys are distinctly different from corresponding uterine patterns, both receptor isoform profiles in primary renal tumors closely resemble those seen in hamster uteri. Renal ER␣ protein and mRNA expression increased after 2.0 and 4.0 months of estrogen treatment and in all renal tumors examined. Using nuclear image cytometry, both early small and large renal tumors were highly aneuploid, indicating that genomic instability is probably a critical early event in estrogen carcinogenesis. (Endocrinology 142: 4006 – 4014, 2001)

medullary region (14, 15). This view is consistent with previous reports (11, 16) that the renomedullary region coincides with the area of the earliest detectable tumorous foci first reported nearly 4 decades ago (8). Nevertheless, more recently, other investigators have suggested that the estrogen-induced renal tumor may arise from either the juxtaglomerular apparatus (12) or vascular smooth muscle cells (13). The present study characterizes both ER␣ and PR receptors, employing immunohistochemical localization, Western blot, in situ hybridization, and Northern blot analyses, during estrogen-induced renal tumorigenesis in the castrated male Syrian hamster. It provides compelling evidence for the cell of origin of this renal tumor. Moreover, the data presented also support the view that estrogen-induced renal tumorigenesis in the hamster is a solely estrogen-driven process that involves a unique proliferative response leading to genomic destabilization as an early initial critical event in tumor formation. Materials and Methods Animals Adult castrated male Syrian golden hamsters (LAK:LVG), outbred strain, weighing 90 –100 g, were purchased from Charles River Laboratories, Inc., Lakeview Hamster Colony (Wilmington, MA). Animals were housed in facilities certified by the American Association for the Accreditation of Laboratory Animal Care. They were acclimated for at least 1 wk before use, maintained on a 12-h light, 12-h dark cycle, and fed certified rodent chow (5002, Ralston Purina Co., St. Louis, MO) and tap water ad libitum. The animal studies were carried out in adherence

4006

Li et al. • Hormone Receptors and Renal Tumorigenesis

with the guidelines established in the Guide for the Care and Use of Laboratory Animals, U.S. Department of Health and Human Resources (NIH 1985). Hamsters in the treatment groups were implanted sc with 20-mg estrogen pellets as described previously (4, 5, 9, 14, 15). To maintain constant levels, fresh pellets were implanted every 3.0 months, and their mean daily absorption was expressed as follows: diethylstilbestrol (DES), 126 ⫾ 9 ␮g/d; and 96 ⫾ 4 ␮g/d. The pellets were prepared by Hormone Pellet Press (Shawnee Mission, KS). Over a 6.0-month period of E2 treatment, the average E2 concentration in serum was 2.28 ⫾ 0.43 ng/ml, and that in the kidney was 4.57 ⫾ 1.04 pg/mg protein (17). Groups of 6 –10 castrated untreated and either DES- or E2-treated hamsters for 1.0 – 8.0 months were used in this study.

Immunohistochemical analysis The kidneys were excised, trimmed, and fixed in 10% buffered formalin, followed by a rapid paraffin-embedding process. Sections (6 ␮m) were prepared and dewaxed. Antigens were retrieved (Target Retrieval Solution, DAKO Corp., Carpinteria, CA) by heating in a water bath set at 97 C for 40 min for ER␣ and 15 min for PR staining and were treated with 3% H2O2 for 15 min to block endogenous peroxidases. After blocking with 6% of the appropriate serum in 1% BSA, the primary antibodies, mouse monoclonal human ER␣ antibody ID5 (DAKO Corp.) and rabbit polyclonal rat ER␣ MC20 or PR antibody C19 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), were applied to the sections overnight at 4 C. It has been previously shown that the hamster ER␣ C-terminal domain has a 100% homology with that of the human (18). As negative controls, similar sections were incubated replacing the ER␣ antibody (ID5) with normal rabbit IgG or in the presence of the respective blocking peptides for ER␣ MC20 or PR C19. Slides were counterstained with hematoxylin, dehydrated in alcohol, and mounted in Permount medium (Permount-xylene, 1:1) before being examined under the microscope.

Quantification of ER␣ and PR in renal sections The expression of ER␣ and PR was evaluated on a 19-in. Sony TV monitor with a Sony CCD/RGB color video camera, equipped with a 0.5-in. camera format and a chip size of 4.8 ⫻ 6.4 mm (Sony, Tokyo, Japan). This conformation provided a camera TV monitor magnification of ⫻2375. Each field of view was 0.16 mm wide by 0.12 mm deep. Beginning at the corticomedullary junction and counting radially toward the medulla, the ratios of either ER␣- or PR-positive to -negative cells were recorded from 12 successive frames. The total counting area was 1.92 mm when moving the field horizontally or 1.44 mm when proceeding vertically. Values from three to five individual kidney sections per time period (0.5, 1.0, 1.6, and 3.0 months) were counted, averaged, and expressed as the mean ⫾ se.

In situ hybridization Nonradioactive in situ hybridization was performed in sections of formalin-fixed, paraffin-embedded kidneys. The methodology was developed from a protocol published by Roche Molecular Biochemicals (Indianapolis, IN) (19). A 700-bp cDNA probe was synthesized from a linearized human ER␣ cDNA (from Dr. G. Greene, University of Chicago, Chicago, IL), and labeled with digoxigenin-conjugated UTP. Briefly, 6-␮m sections were deparaffinized with xylene, dehydrated in ethanol, treated with proteinase K, fixed again, and incubated for 10 min in 4 ⫻ SSC containing 50% formamide. Hybridization was performed for 24 – 48 h at 42 C in hybridization buffer (10% dextran sulfate, 0.02% Ficoll, 0.02% polyvinylpyrrolidone, 10 mg/ml BSA, 4 ⫻ SSC, 10 mm DTT, 1 mg/ml tRNA, and 1 mg/ml sperm DNA) containing the labeled ER␣ riboprobe, 5 ng/ml hybridization buffer, in a humid chamber. After blocking with 5% normal sheep serum for 30 min, the sections were incubated with sheep-antidigoxigenin IgG conjugated with alkaline phosphatase (1:400) for 3 h at room temperature, followed by PBS buffer rinses. The signal was visualized by color development with 5-bromo4-chloro-3-indol phosphate and nitro blue tetrazolium in the presence of 1 mm levamisole to inhibit endogenous alkaline phosphatases. The specificity of the reaction was determined by ribonuclease treatment and postfixation before hybridization with the probe, hybridization with sense riboprobe, and replacement of antidigoxigenin antibody with BSA. All controls were negative.

Endocrinology, September 2001, 142(9):4006 – 4014 4007

Western blot analysis For Western blot analysis of ER␣, ER␤, and PR, kidney cytosolic fractions from 6 –10 hamsters/group were used. Tissue samples were homogenized by Polytron (Brinkmann Instruments, Inc., Westbury, NY) using the following buffer: 50 mm Tris-HCl (pH 7.4), 0.2 m NaCl, 2 mm EDTA, 0.5% Nonidet P-40, 50 mm NaF, 0.5 mm Na3VO4, 20 mm sodium pyrophosphate, 1 mm phenylmethylsulfonylfluoride, 10 ␮g/ml aprotinin, 10 ␮g/ml leupeptin, and 1 mm dithiothreitol. The tissue lysate was centrifuged at 12,000 rpm for 20 min at 4 C. The supernatant was collected, and its protein content was measured with bicinchoninic acid reagents (Pierce Chemical Co., Rockford, IL). Protein aliquots (20 ␮g) were electrofractionated on SDS-PAGE. Equal loading was determined by staining the gel with Coomassie blue. The proteins were transferred onto nitrocellulose membranes and probed with primary antibody overnight at 4 C, followed by incubation with peroxidase-conjugated second antibody for 2 h. The signals were visualized and amplified by ECL Western blot detection reagents (Amersham Pharmacia Biotech, Arlington Heights, IL). The following primary antibodies were used: rabbit polyclonal ER␣ MC20, raised against the last 20 amino acids of the rat ER␣ C-terminal; PR C-19, raised against amino acids 545–564 of human PR, and their respective blocking peptides; as well as goat polyclonal ER␤ antibodies Y-19, raised against the amino-terminus of the ER␤ of mouse origin, and L-20, raised against the carboxyl-terminus of ER␤ of human origin. All of these antibodies were from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA).

Probe labeling [␣-32P]CTP-labeled riboprobes were generated in vitro using T7 or T3 polymerase with linearized cDNA subclones according to the supplier’s recommendations. The probes used were a 700-bp cDNA synthesized from a linearized human ER␣ cDNA and a 56-bp oligonucleotide probe specific for the rat ER␤ (17). The labeled products were purified by Sephadex G-50 Quick-Spin columns (Boehringer Mannheim Co., Indianapolis, IN) (20).

Preparation of RNA RNA was prepared by the method of Chomczynski and Sacchi (21) with some modifications. Briefly, tissue samples were homogenized for 60 sec in 5 ml 5 m guanidium isothiocyanate using a Polytron (Brinkmann Instruments, Inc.) set at maximum speed, followed by phenolchloroform extraction. The RNA was precipitated with isopropanol and dissolved in ribonuclease-free water. The RNA concentration was determined by absorbance at 260 nm.

Northern blot analysis To reduce the effect of individual variation, Northern blots were prepared using RNA pooled from three hamsters per group. Additional pooled samples were obtained from six to nine groups of hamsters to confirm results obtained. Briefly, 10 ␮g denatured total RNA were loaded onto each lane and fractionated in 2.2 m formaldehyde-1.5% agarose gel. All gels were stained with acridine orange and photographed to ascertain the integrity of RNA samples and to confirm that equal amounts of RNA were loaded. RNA was then transferred to Hybond nylon membranes using a capillary transfer consisting of 10 ⫻ SSC (1 SSC ⫽ 150 mm NaCl and 15 mm sodium citrate, pH 7.0) and UV cross-linked with total energy of 0.3/formamide, 0.5% dextran sulfate, and 50 ␮g/ml yeast tRNA for 3 h at 65 C. Blots were hybridized with labeled (106 cpm/ml) probes for 16 –18 h at 65 C in hybridization mixture. Blots were washed for 1 h in 0.3 ⫻ SSC/0.1% SDS at 65 C and twice for 1 h in 0.1 ⫻ SSC/0.1% SDS at 65 C, then exposed to x-ray film with or without intensifying screens. Control for equal loading and blotting of RNA was performed by stripping blots in boiling 0.05 ⫻ SSC/0.1% SDS and rehybridizing with antisense riboprobe generated from cDNA clones for GAPDH. The hybridization signals were scanned and quantified with a Personal Densitometer SI (Molecular Dynamics, Inc., Sunnyvale, CA).

Nuclear image cytometry (NIC) NIC was used to assess DNA ploidy in hamster renal samples. Untreated and estrogen-treated tissue sections (6 and 8 ␮m) were stained

4008

Endocrinology, September 2001, 142(9):4006 – 4014

with hematoxylin and eosin for histological evaluation. Other serial sections were hydrolyzed with 5 n HCl for 60 min and stained by the Feulgen technique using a CAS Quantitative DNA staining kit (Cell Analysis Systems, Lombard, IL). NIC analysis was performed on a CAS 200, employing Quantitative DNA Analysis software, version 2.5. Small and large renal tumor foci seen at 3.0 –5.0 months of either E2 or DES treatment were confirmed on hematoxylin- and eosin-stained sections, and the corresponding regions were located on Feulgen-stained serial sections. After calibration using hamster kidney cells, the renal samples containing early and established kidney tumors were selected and analyzed at ⫻40 magnification for DNA content (DNA index). Inflammatory, stromal, and nonmalignant cells were eliminated from analysis. A minimum of three to five individual normal, estrogen-treated, and kidney tumor sections were scanned multiple times, totaling 200-1000 cells/ sample. The histograms were classified according to their DNA index as follows: hypoploid, less than 0.85; diploid, 0.85–1.15; aneuploid/hyperdiploid, 1.16 –1.90; and tetraploid or proliferating cells, 1.90 –2.10.

Results ER␣ localization in the renal cortex

In untreated castrated male hamsters, ER␣⫹ staining was consistently observed in proximal tubule cells. Although the ER␣⫹ staining in the nuclei of most proximal tubule cells was distinct, it was uniformly weak throughout the cortex (Fig. 1A). No positive ER␣ staining was observed in any of the samples tested when the ER␣ antibody (ID5) was replaced with normal rabbit IgG (Fig. 1B). When male hamsters were treated continuously with estrogen for 1.0 –2.0 months, the weak ER␣⫹ staining of the proximal tubule cells disappeared, indicating a down-regulation of the ER␣ after estrogen treatment. After 2.0 –3.0 months of either E2 or DES treatment, individual and small groups of renal interstitial cells began to exhibit intense ER␣⫹ staining (Fig. 1C). These ER␣⫹ cells became increasingly evident in the interstitium at the corticomedullary region of the kidney and in nascent tumorous lesions and foci after 3.0 – 4.0 months of treatment. Interestingly, not all of the renal interstitial cells in this region exhibited ER␣⫹ staining (Fig. 1, C and D). With prolonged estrogen treatment, between 4.0 – 6.0 months, virtually all the cells of kidney tumorous foci as well as small and large renal tumor foci displayed marked ER␣⫹ staining (Fig. 1E). This ER␣⫹ staining in renal tumor foci was consistently present throughout these treatment periods. Both DES- and E2treated kidneys exhibited essentially the same pattern of ER␣⫹ staining in all renal tumor foci observed. Similar results were obtained with both ER␣ antibodies tested (ID5 and MC20); however, the quality of the staining was better when the ID5 antibody was used. PR localization in the renal cortex

In contrast, no detectable PR⫹ staining was seen in renal sections from untreated castrated male hamsters. However, after only 2 wk of estrogen treatment, PR⫹ staining was seen in single interstitial cells (Fig. 2, A and C). The pattern of PR⫹ staining was largely confined to interstitial cells located at the renal corticomedullary region, as seen for ER␣⫹-stained cells. Similarly, not all of the kidney interstitial cells in this region exhibited PR⫹ staining (Fig. 2C). After only 1.5 months of estrogen treatment, PR⫹ staining was observed in individual and small clusters of renal interstitial cells (Fig. 2D) as well as in cells of nascent renal tumorous foci consistently seen after 2.0 months of estrogen treatment (Fig. 2E). Correspond-


ingly, in the absence of the PR primary antibody or in the presence of the PR-blocking peptide, no PR⫹ renal interstitial cells were detected (Fig. 2, B and F). After continued estrogen treatment, all small (Fig. 2G) and large renal tumor foci (Fig. 2H) exhibited intense PR⫹ staining. In serial sections, cells from early renal tumorous foci and small and large renal tumors colocalized both ER␣⫹ and PR⫹ staining (data not shown). Quantification of the ER␣ and PR in renal sections

A quantitative evaluation of the changes observed in ER␣⫹ and PR⫹ interstitial cells in the renal corticomedullary region as a function of duration of estrogen treatment is shown in Fig. 3. These data clearly demonstrate an early and gradual rise in the number of PR⫹-stained kidney interstitial cells from 2 wk to 3.0 months of estrogen treatment. The rise in PR⫹ cells in a subset of renal interstitial cells was statistically significant (P ⬍ 0.05) after 1.5 and 3.0 months of estrogen treatment compared with that in animals treated for 0.5 months. Interestingly, an increase in the number of PR⫹ renal interstitial cells preceded the increase in ER␣⫹ interstitial cells, the latter not occurring until after 1.5 months of estrogen treatment (Fig. 3). The rise in ER␣⫹ renal interstitial cells was statistically significant (P ⬍ 0.05) at 3.0 months of estrogen treatment compared with those observed after 0.5 months of treatment or with its age-matched untreated control group. Hamster kidney ER␣ isoforms

Western blot analysis using the ER-MC20 antibody confirmed the presence of ER␣ in the hamster kidney. Untreated, intact or castrated male kidneys exhibited two isoforms of the ER␣, a variant 64-kDa and a 50-kDa truncated isoform. The level of expression of the 64-kDa ER␣ isoform was consistently lower in renal samples from castrated males compared with those from intact animals. After only 1.0 month of estrogen treatment, the level of expression of this ER␣ isoform in castrated hamsters was indistinguishable from that in intact males. Prolonged estrogen treatment (3.0 and 5.0 months) resulted in a further rise in the level of expression of the 64-kDa isoform (Fig. 4). In addition, another ER␣ isoform (58 kDa) present in intact female hamster kidney samples was detected in castrated males after 5.0 months of estrogen treatment (Fig. 4). In addition to the 66-kDa native form of ER␣, both the 58-, and the 50-kDa isoforms were prominent in estrogen-induced renal tumors. These three ER␣ isoforms were also observed in the intact untreated hamster uterus. The 50-kDa ER␣ isoform was present in all hamster kidney samples from untreated males and females and estrogen-treated male, samples from estrogen-induced tumors, as well as intact untreated hamster uteri (Fig. 4). No appreciable changes in the level of expression of this ER␣ isoform were observed during various periods of estrogen treatment. Neutralizing the ER␣ primary antibody with its blocking peptide resulted in the loss of expression of these three ER␣ isoforms. Experiments similar to those described for the detection of ER␣ isoforms were performed employing Y-19 (mouse) and L-20 (human) ER␤ antibodies. We analyzed ER␤ expression in 2.0-, 5.0-, and 8.0-month estrogen-



FIG. 1. Immunohistochemical detection of ER␣ expression in castrated untreated and estrogen-treated male Syrian hamster kidneys. A, Cortical tubule cells (arrows) exhibited weak ER␣⫹ nuclear staining in an untreated, castrated male hamster kidney (magnification, ⫻300). B, Serial section of cortical tubule cells from an untreated castrated male hamster kidney substituting the ER␣ antibody with normal rabbit IgG (magnification, ⫻250). C, The arrows point to individual and small groups of ER␣⫹ interstitial cells from a 3.0-month estrogen-treated hamster kidney (magnification, ⫻200). D and E. ER␣⫹ cells in a small and moderate size renal tumor foci from a 4.0-month (D) and 5.0-month (E; magnification, ⫻250) estrogen-treated hamster kidney (magnification, ⫻200). F, Localization of ER␣ mRNA by nonradioactive in situ hybridization in an estrogen-induced renal tumor foci (magnification, ⫻150).

treated hamster kidneys as well as in primary renal tumors and their respective untreated age-matched controls. Such experiments did not reveal ER␤ protein signals in any of the kidney or tumor samples examined (data not shown). Hamster kidney PR isoforms

Western blot analysis of the PR indicated that renal samples from intact male and female hamsters and castrated male hamsters exhibited negligible to low levels of the PR-B

(120 kDa), PR-A (94 kDa), and PR-C (64 kDa) isoforms (Fig. 4). Estrogen treatment resulted in a modest elevation in the level of expression of the PR-A isoform as well as two PR-A1 and -A2 forms, 103 and 76 kDa (Fig. 4). Additionally, a distinct PR-B isoform was detected in kidney samples derived from 5.0-month estrogen-treated hamsters. Of particular interest in the estrogen-induced renal tumor samples was the marked expression of PR-B, -A, and -C isoforms and the absence of the additional PR forms observed in the un-

4010



FIG. 2. Immunohistochemical detection of estrogen-induced PR expression in A) castrated males. A, 0.5-month, DES-treated hamster kidney, interstitial cells stained for PR (magnification, ⫻150); B, 0.5-month DES-treated serial section to A; interstitial cells at the cortical-medullary



expression of the PR isoforms in the estrogen-induced primary renal tumor resembled the profile seen in the hamster uterus. All signals were blocked when the PR-C19 antibody was neutralized with its blocking peptide. ER␣ and ER␤ mRNA expression

FIG. 3. Time course of ER␣ and PR expression in hamster kidney. The expression of ER␣ (E, control untreated; F, estrogen-treated) and PR (‚, control untreated; Œ, estrogen-treated) positive renal interstitial cells was evaluated on a 19-in. Sony TV monitor with a Sony CCD/RGB color video camera. Beginning at the corticomedullary junction and counting radially toward the medulla, the percentage of either ER␣- or PR-positive to -negative cells was recorded from 12 successive frames. Values from 3–5 individual kidney sections/time period (0.5, 1.0, 1.5, and 3.0 months) were counted, averaged, and expressed as the mean ⫾ SE. PR expression (多), P ⬍ 0.05 compared with 0.5-month estrogen treatment and to their respective untreated age-matched controls. ER expression (夞), P ⬍ 0.05 compared with 0.5-month estrogen treatment and its respective untreated agematched controls.

Northern blot analysis of RNA extracts from age-matched untreated kidneys exhibited very low levels of ER␣ expression. After 2.0 and 4.0 months of estrogen treatment, an increase in ER␣ mRNA expression was evident, and a further rise was observed in all renal tumor samples examined (Fig. 5). This increase in ER␣ mRNA expression in estrogeninduced renal tumors was consistently greater than that observed in RNA extracts from hamster uterus (Fig. 5). Using nonradioactive in situ hybridization, elevated levels of ER␣ mRNA were localized in all small and large primary renal tumor foci induced by either DES or E2 (Fig. 1D). The signal detected was mainly in the nuclei. In contrast, renal ER␤ mRNA expression was undetected by Northern blot analysis in age-matched untreated hamster kidneys, 2.0- and 5.0month estrogen-treated kidneys (data not shown), and all renal tumor samples studied (Fig. 5). Rat ventral and dorsal prostate samples served as positive signal controls. Analysis of genomic instability: aneuploidy

NIC was employed to analyze Feulgen-stained sections of renal samples from untreated, castrated, age-matched hamsters, early renal tumor foci from 3.0- and 4.0-month estrogen-treated animals, and frank renal tumors taken from 6.0to 8.0-month estrogen-treated hamsters. As expected, untreated kidney sections exhibited a normal diploid frequency (n ⫽ 44; Fig. 6A). The same diploid frequency was seen in uninvolved renal tissue adjacent to renal tumor foci (data not shown). In contrast, early kidney tumorous lesions and small renal tumor foci examined after 3.5–5.0 months of estrogen treatment were all highly aneuploid, exhibiting frequencies of 93.5 ⫾ 1.2 (n ⫽ 3; Fig. 6B). The aneuploid frequency of large, well established renal tumor foci was also substantial (91.8 ⫾ 5.4; n ⫽ 5; Fig. 6C). FIG. 4. Western blot analysis of ER␣ and PR during estrogen-induced renal tumorigenesis. Twenty micrograms of total protein extracts from hamster kidneys of intact males (IMK) or castrated males (CMK) treated for 1.0 (E-1), 3.0 (E-3), and 5.0 months (E-5), renal tumor samples from hamsters treated with estrogen for 8.0 –10.0 months (KT), uteri from intact female hamsters (HU), and kidneys from intact females (IFK) were separated by SDS-PAGE, transferred to nitrocellulose membrane, and immunoblotted with ER␣ (MC20) and PR (C19) antibodies. The arrows at the left of the blot are molecular mass markers run along with the samples. Specific bands for ER␣ and PR are indicted by arrows on the right. The results are representative of one of five similar independent experiments.

treated and estrogen-treated kidney samples (Fig. 4). The

Discussion

It has been confounding that the Syrian hamster kidney behaves as an estrogen target tissue, uniquely giving rise to solely estrogen-induced and -dependent neoplasms. A clue to this unusual occurrence may be derived from the observation that the hamster reproductive and urinary systems, as in most, if not all, mammalian species, arise from the same germinal ridge of multipotential cells (8). Based on the results provided herein, it is proposed that some of these germinal or stem cells, which would normally be destined to the uterus, migrate and establish themselves in the renal corticomedullary region (Fig. 7). These ectopically located germ

junction stained in the presence of PR-blocking peptide (magnification, ⫻100), PR⫹ interstitial cells (arrows) at the cortical-medullary junction. C, PR⫹ individual interstitial cells (arrows) at the cortical-medullary junction (magnification, ⫻450). D, PR⫹ cells (arrows) in a small number of interstitial cells after 1.5 months of DES treatment (magnification, ⫻400). E, PR⫹ cells in a nascent tumor foci after 2.0 months of DES treatment (magnification, ⫻200). F, Serial sections of early renal tumor foci (arrow), 2.0 months of DES treatment, stained in the presence of PR blocking peptide (magnification, ⫻200). G, PR⫹ cells in early renal tumor foci, 3.0 months after DES treatment (magnification, ⫻200). H, PR⫹ cells (arrows) in a moderate size renal foci, 5.0 months after DES treatment (magnification, ⫻200).

4012


FIG. 5. Northern blot analysis of ER␣ and ER␤ during estrogeninduced renal tumorigenesis. Total RNA from hamster uteri (HU), age-matched, untreated kidneys from 2.0 (CK-2), 5.0 (CK-5), and 8.0 months (CK-8); estrogen-treated kidneys from 2.0 (EK-2) and 5.0 months (EK-5); renal tumor samples from estrogen-treated hamsters for 8.0 –10.0 months (KT1 and KT2); and rat ventral (RVP) and dorsal prostates (RDP) was isolated for Northern blot analysis, as described in Material and Methods. GADPH served as the internal control. Each slot contained three pooled samples from individual hamster kidneys, tumors, uteri, or rat prostates. The results represent one of four similar independent experiments.

cells would remain dormant unless exposed to a sustained level of estrogen. This idea is supported by the resemblance of early renal interstitial lesions to blastema and their strong staining using mesenchymal markers (i.e. vimentin and desmin) with only a trace of cytokeratin (14, 15). It is further supported by our finding that only a subset of multipotential renal interstitial cells responds to estrogen treatment by expressing PR and ER␣, and by the close resemblance of the ER␣ and PR isoform profiles of the hamster primary estrogen-induced and -dependent renal tumor to those found in the uterus. Moreover, the lack of ER␤ expression in estrogeninduced renal neoplasms is consistent with the established selective expression and proliferative role of ER␣ in uterine tissue (22–24). In addition, the ability of progesterone to inhibit these renal tumors is consistent with the antiproliferative role of progesterone in the uterus in contrast to the mammary gland, where estrogen and progesterone are both mitogenic. The overexpression of early estrogen response genes (c-myc, c-fos, and c-jun) (25), the high proliferative activity (PCNA labeling) observed only in these interstitial renal cells in response to estrogen treatment (26), and the 2.0to 3.0-fold increase in hamster renal cell number observed after 1 nm E2 treatment in serum-free, chemically defined culture conditions, which was inhibited by tamoxifen (27), indicate that this subset of renal interstitial cells are bona fide estrogen target cells, that is, estrogen responsive and dependent. This renal interstitial cell estrogen responsiveness, however, may be unique to the Syrian hamster. The untreated or estrogenized Turkish hamster does not express


either PR or ER␣ in its renal interstitial cells when assessed by immunohistochemistry and Western blot analysis (Li, J. J., J. Coe, and S. A. Li, unpublished data). Interestingly, it has been reported that Turkish hamsters chronically treated with estrogens do not develop renal tumors (28). The expression of ER␣ mRNA and protein during early murine embryogenesis has been shown (29 –31). Interestingly, in newborn mouse uterine cells, ER␣ was demonstrated in uterine stromal cells at a much higher level than in epithelial cells (32, 33). Although numerous reports have characterized ER␣ isoforms in target tissues of a variety of species, none has as yet been shown for hamster tissues. Earlier reports by our laboratory, however, have shown the estrogen specificity and sucrose gradient sedimentation characteristics of an ER in hamster kidneys derived from untreated and estrogen-treated animals and in primary renal tumors (34, 35). In murine uteri it is now evident that the native form of the ER␣ has a molecular mass of 65– 67 kDa (36, 37). The ER␣ protein profile (66, 58, and 50 kDa) of the E2-induced and -dependent primary renal tumor is essentially identical to that of the hamster uterus. In contrast, the ER␣ protein profiles of estrogen- and estrogen- plus androgen-induced primary mammary neoplasms of ACI and Noble rats, respectively, differ markedly from their respective ER␣ uterine profiles (38, 39). The rise in the ER␣ 64-kDa isoform in the hamster kidney with increasing duration of estrogen treatment indicates that this isoform may contribute to the growth advantage of nascent and early interstitial renal foci, as it has been suggested that specific ER␣ isoforms may differentially affect signaling mechanisms and, thus, specific gene activities within target tissues (40, 41). Consistent with the rise in ER␣ immunostaining, and particularly the 64-kDa ER␣ isoform concentration, a concomitant rise in renal ER␣ mRNA expression was observed after 2.0 and 4.0 months of estrogen treatment and in estrogen-induced kidney tumors. These findings extend previous ER␣ mRNA data in hamster kidneys treated for 7.0 months with estrogen (42). The lack of ER␣ immunostaining in untreated normal kidneys and the relatively poor ER␣ immunostaining in the primary renal tumor in the same report (42) were probably due to the much weaker ER␣ antibody used by these investigators. Similar to ER␣, PR has also been detected in fetal uterine cells (43). At least three major PR isoforms have been identified in various target tissues (PR-B, -A, and -C) (44 – 47). The specificity and binding characteristics of the hamster kidney PR during estrogen-induced tumorigenesis and renal tumor development have been previously reported by us (48, 49). The levels of PR-B and PR-A in the primary renal tumor were consistently equal in all of the renal tumor samples examined. Although the level of PR-A was lower than that of PR-B in hamster uterine samples from untreated intact female hamsters, it is known that PR-B concentrations vary considerably throughout the estrous cycle (50). The presence of PR-C in the primary renal tumor, evidently localized in the cytosol, may also play an important role in modulating progesterone action during estrogen-induced renal tumorigenesis. Furthermore, when progesterone is administered concomitantly with estrogen, the data suggest that the estrogen induction of PR-A alone, or possibly together with PR-A1 and -A2, may be sufficient to block renal tumorigenesis in the absence of



FIG. 6. Representative NIC histogram from A) 3.5-month, untreated castrated hamster kidney (DNA index, 0.99; diploid, 100%), B) early tumorous lesions, 3.5-month, DES-treated (DNA index, 1.18; aneuploid, 94%), and C) well established kidney tumor foci, 6.0-month, DES-treated (DNA index, 1.31; aneuploid, 100%). The data represent one of three to five similar independent experiments.

mediated carcinogenesis in the kidney. The aneuploidy seen in nascent renal tumorous lesions and early small renal tumor foci is accompanied by both ER␣ and PR positivity and increased proliferative activity (PCNA labeling) (26). Moreover, these findings are consistent with both estrogen- and estrogen- plus androgen-induced mammary tumors in ACI and Noble rats, respectively; that is, genomic destabilization is found in both early and well established tumors (37). Taken together, these data indicate that genomic instability is probably a critical early event in estrogen carcinogenesis. Acknowledgments

FIG. 7. Schematic representation of the germinal ridge in the Syrian hamster. The hamster reproductive tract (round cells) and urinary tract (elongated cells) systems arise from the same germinal ridge of multipotential cells. The reproductive germinal cells (round), which would normally be destined to reside in uterus, migrate and establish themselves in the corticomedullary region of the kidney. These ectopically located germ cells remain dormant unless exposed to a sustained level of estrogen.

appreciably levels of PR-B and PR-C (8, 9). In this regard, recent evidence indicates that PR-A and PR-B may differentially regulate a subset of progesterone-responsive target genes (51), ultimately affecting progesterone action. The immunoblots clearly show that the PR expression profile of the renal tumor closely resembles that of the uterus. The results presented herein, employing the expression of PR and ER␣ as biomarkers during estrogen-induced renal tumorigenesis, are consistent with the view that renal tumors arise from a subset of multipotential interstitial cells driven to proliferate by estrogens. Moreover, there is no evidence of distinct intervening dysplastic changes during estrogen treatment, but, rather, there is a progressive continuum leading to renal tumor development. The data from NIC analyses, performed in tissue sections of renal tumorous foci and frank kidney tumors, support our recent karotypic studies derived from metaphases of cultured cells from either estrogen-treated kidneys or well established renal tumors (26); that is, genomic instability is an early event in estrogen-

Received February 12, 2001. Accepted May 3, 2001. Address all correspondence and requests for reprints to: Dr. Jonathan J. Li, Division of Etiology and Prevention of Hormonal Cancers, Kansas Cancer Institute, University of Kansas Medical Center, 1043 Lied Biomedical Research Facility, 3901 Rainbow Boulevard, Kansas City, Kansas 66160-7312. E-mail: [email protected]. This work was supported by NIH Grant 2-R01-CA-58030-08.

References 1. Hoover RN 1995 Sex hormones and human carcinogenesis. In: Becker KL, ed. Principles and practice of endocrinology and metabolism, chapt 216, 2nd Ed. New York: Lippinicott; 1861–1868 2. Riman T, Persson I, Nilsson S 1998 Hormonal aspects of epithelial ovarian cancer: review of epidemiological evidence. Clin Endocrinol (Oxf) 49:695–707 3. Chang WY, Prins GS 1999 Estrogen receptor-␣: implications for the prostate gland. Prostate 40:115–124 4. Li JJ, Li SA, Klicka JK, Parsons JA, Lam LKT 1983 Relative carcinogenic activity of various synthetic and natural estrogens in the hamster kidney. Cancer Res 43:5200 –5204 5. Li JJ, Li SA, Oberley TD, Parsons JA 1995 Carcinogenic activities of various steroidal and nonsteroidal estrogens in the hamster kidney: relation to hormonal activity and cell proliferation. Cancer Res 55:4347– 4351 6. Li JJ, Li SA 1990 Estrogen carcinogenesis in hamster tissues: a critical review. Endocr Rev 11:524 –531 7. Pour P, Althuff J, Salmasi SZ, Stephen K 1979 Spontaneous tumors and common diseases in two colonies of Syrian hamsters. III. Urogenital system and endocrine glands. J Natl Cancer Inst 56:949 –961 8. Kirkman H 1959 Estrogen-induced tumor of the kidney in the Syrian hamster. J Natl Cancer Inst Monogr 1:1–139 9. Li JJ, Cuthbertson, Li SA 1980 Inhibition of estrogen carcinogenesis in the Syrian golden hamster kidney by antiestrogens. J Natl Cancer Inst 64:795– 800 10. Li JJ, Hou X, Bentel JM, Yazlovitskaya EM, Li SA 1998 Prevention of estrogen carcinogenesis in the hamster kidney by ethinylestradiol: some unique properties of a synthetic estrogen. Carcinogenesis 19:471– 477 11. Llombart-Bosch A, Peydro A 1975 Morphological, histochemical, and ultrastructural observations of DES-induced kidney tumors in the Syrian golden hamster. Br J Cancer 11:403– 412 12. Dodge AH, Brownfield M, Reid IA, Inagami R 1988 Immunohistochemical

4014 Endocrinology, September 2001, 142(9):4006 – 4014

13. 14. 15. 16.

17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29.

30. 31. 32.

renin study of DES-induced renal tumor in the Syrian hamster. Am J Anat 182:347–352 Hacker HJ, Bannasch P, Liehr JG 1988 Histochemical analysis of the estradiolinduced kidney tumors in male Syrian hamsters. Cancer Res 48:971–976 Gonzalez A, Oberley TD, Li JJ 1989 Morphological and immunohistochemical studies of the estrogen-induced Syrian hamster renal tumor: probable cell of origin. Cancer Res 49:1020 –1028 Oberley TD, Gonzalez A, Lauchner LJ, Oberley LW, Li JJ 1991 Characterization of early lesions in estrogen-induced renal tumors in the Syrian hamster. Cancer Res 51:1922–1929 Cortes-Vizcaino V, Peydro-Olaya A, Llombart-Bosch A 1994 Morphological and immunohistochemical support for the interstitial cell origin of oestrogeninduced kidney tumors in the Syrian golden hamster. Carcinogenesis 15:2155– 2162 Li SA, Xue QY, Xie Q, Li CI, Li JJ 1994 Serum and tissue levels of estradiol during estrogen-induce renal tumorigenesis in the Syrian hamster. J Steroid Biochem Mol Biol 48:283–286 Bhat HK, Vadgama JV 2000 Hamster estrogen receptor cDNA: Cloning and mRNA expression. J Steroid Biochem Mol Biol 72:47–53 Roche Molecular Biochemicals 2000 Nonradioactive in-situ hybridization application manual, 2nd Ed. Indianapolis: Roche Molecular Biochemicals Sanbrook J, Fritch EF, Maniatis T 1989 Molecular cloning: a laboratory manual. Cold Spring Harbor: Cold Spring Harbor Laboratory Chomczynski P, Sacchi N 1987 Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 162: 156 –159 Gustafsson JA 2000 Novel aspects of estrogen action. J Soc Gynecol Invest 7(Suppl 1):58 –59 Taylor AH, Al-Azzawi F 2000 Immunolocalization of estrogen receptor ␤ in human tissues. J Mol Endocrinol 1:145–155 Pelletier G, Labrie C, Labrie F 2000 Localization of oestrogen receptor ␣, oestrogen receptor ␤, and androgen receptors in the rat reproductive organs. J Endocrinol 165:359 –370 Hou X, Li JJ, Chen W, Li SA 1996 Estrogen-induced proto-oncogene and suppressor gene expression in the hamster kidney: significance for estrogen carcinogenesis. Cancer Res 56:2616 –2620 Li JJ, Hou X, Banerjee SK 1999 Overexpression and amplification of c-myc in the Syrian hamster kidney during estrogen carcinogenesis: a probable critical role in neoplastic transformation. Cancer Res 59:2340 –2346 Oberley TD, Lauchner LJ, Pugh TD, et al. 1989 Specific estrogen-induced cell proliferation of cultured Syrian hamster renal proximal tubular cells in serumfree chemically defined media. Proc Natl Acad Sci USA 86:2107–2111 Coe JE, Cieplak W, Hadlow WJ, Ross MJ 1997 Female proteins, amyloidosis, and hormonal carcinogenesis in Turkish hamster: differences from Syrian hamster. Am J Physiol 273:R934 –R941 Greco TL, Duello TM, Gorski J 1993 Estrogen receptors, estradiol, and diethylstilbestrol in early development: the mouse as a model for the study of estrogen receptors and estrogen sensitivity in embryonic development of male and female reproductive tracts. Endocr Rev 14:59 –71 Lemmen JG, Brockhof JLM, Kuiper GG, Gustafsson J-A, vanderSaag PT, vanderBurg B 1999 Expression of estrogen receptor ␣ and beta during mouse embryogenesis. Mech Dev 81:163–167 Hiroi H, Momoeda M, Inoue S, et al. 1999 Stage-specific expression of estrogen receptor subtypes and estrogen responsive finger protein in preimplantational mouse embryos. Endocr J 46:153–158 Yamashita S, Newbold RR, McLachlan JA, Korach KS 1989 Developmental pattern of estrogen receptor expression in female mouse genital tracts. Endocrinology 125:2888 –2896


33. Bigsby RM, Li AX, Luo K, Cunha GR 1990 Strain differences in the ontogeny of estrogen receptors in murine uterine epithelium. Endocrinology 126:2592– 2596 34. Li JJ, Talley DJ, Li SA, Villee CA 1974 An estrogen binding protein in renal cytosol of intact, castrated, and estrogenized Golden hamsters. Endocrinology 95:1134 –1141 35. Li JJ, Talley DJ, Li SA, Villee CA 1976 Receptor characteristics of specific estrogen binding in the renal adenocarcinoma of the Golden hamster. Cancer Res 36:1127–1132 36. Horigome T, Ogata F, Golding TS, Korach KS 1988 Estradiol-stimulated proteolytic cleavage of the estrogen receptor in mouse uterus. Endocrinology 123:2540 –2548 37. Furlow JD, Ahrens H, Mueller GC, Gorski J 1990 Antisera to a synthetic peptide recognize native and denatured rat estrogen receptors. Endocrinology 127:1028 –1032 38. Li SA, Liao JD, Li JJ 2001 Estrogen-induced breast cancer in female ACI rats. In: Li JJ, Li SA, Daling JR, eds. Hormonal carcinogenesis, chapt 13. New York: Springer-Verlag; vol 3:178 –188 39. Liao JD, Pantazis CG, Hou X, Li SA 1998 Promotion of estrogen-induced mammary gland carcinogenesis by androgen in the male Noble rat: probable mediation by steroid receptors. Carcinogenesis 19:2173–2180 40. Horigome T, Ogata F, Golding TAS, Korach KS 1988 Estradiol-stimulated proteolytic cleavage of the estrogen receptor in the mouse uterus. Endocrinology 123:2540 –2548 41. Hu C, Hyder SM, Needleman DS, Baker VV 1996 Expression of estrogen receptor variants in normal and neoplastic human uterus. Mol Cell Endocrinol 118:173–179 42. Bhat HK, Hacker H-J, Bannasch P, Thompson EA, Liehr JG 1993 Localization of estrogen receptors in interstitial cells of hamster kidney and in estradiolinduced renal tumors as evidence of the mesenchymal origin of this neoplasm. Cancer Res 53:5447–5451 43. Sumida C, Lecerf F, Pasqualini JR 1988 Control of progesterone receptors in fetal uterine cells in culture: effects of estradiol, progestins, antiestrogens, and growth factors. Endocrinology 122:3–11 44. Ogle TF, Dai D, George P, Mahesh VB 1997 Stromal cell progesterone and estrogen receptors during proliferation and regression of the decidua basalis in the pregnant rat. Biol Reprod 57:495–506 45. Park-Sarge O, Parmer TG, Gu Y, Gibori G 1995 Does the rat corpus luteum express the progesterone receptor gene? Endocrinology 136:1537–1543 46. Shyamala G, Louce SG, Camarillo IG, Talamantes F 1999 The progesterone receptor and its isoforms in mammary development. Mol Gen Metab 68:182– 190 47. Wei LL, Miner R 1994 Evidence for the existence of a third progesterone receptor protein in human breast cancer cell line T47D. Cancer Res 54:340 –343 48. Li SA, Li JJ, Villee CA 1977 Significance of the progesterone receptor in the estrogen-induced and dependent renal tumor of the Syrian golden hamster. Ann NY Acad Sci 286:369 –382 49. Li SA, Li JJ 1978 Estrogen-induced progesterone receptor in the Syrian hamster kidney. I. Modulation by antiestrogens and androgens. Endocrinology 103: 2119 –2128 50. Mangal RK, Wiehle RD, Pomdexter III AN, Weigel NL 1997 Differential expression of uterine progesterone receptor forms A and B during the menstrual cycle. J Steroid Biochem Mol Biol 63:195–202 51. Mulac-Jericevic B, Mullinax RA, DeMayo FJ, Lydon JP, Conneely OM 2000 Subgroup of reproductive functions of progesterone mediated by progesterone receptor-B isoform Science 289:1751–1754