SEARCH

SEARCH BY CITATION

Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. References

The effects of 17β-estradiol and the important estrogen metabolites, 2-hydroxyestrone (2-OHE1) and 16α-hydroxyestrone (16α-OHE1) on bone, mammary gland, and uterine histology, and on blood cholesterol were investigated in ovariectomized growing rats. Rats were treated with 200 μg/kg of body weight/day of each of the test compounds for 3 weeks. Ovariectomy resulted in uterine and mammary gland atrophy, increased body weight, bone turnover and tibia growth, and hypercholesterolemia. 17β-estradiol treatment prevented these changes, with the exception that this high dose of estrogen did not prevent hypercholesterolemia. 2-OHE1 had no effect on any of the measurements. 16α-OHE1 resulted in bone measurements that did not differ from the 17β-estradiol–treated rats and prevented the increase in serum cholesterol. In contrast, 16α-OHE1 resulted in increases in uterine weight, uterine epithelial cell height, and mammary gland cell proliferation that were significantly less than the 17β-estradiol treatment. These findings demonstrate that 16α-hydroxylation of estrone results in tissue-selective estrogen agonistic activity, whereas 2-hydroxylation resulted in no measured activity. Furthermore, they suggest that factors that modulate the synthesis of these metabolites could selectively influence estrogen target tissues.


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. References

17β-ESTRADIOL IS METABOLIZED through a series of reversible and nonreversible enzyme-mediated steps. The initial step involves 17β-dehydrogenase–mediated reversible formation of estrone.1 This is followed by hydroxylation at either the C-2 or C-16α site.2 Although other hydroxylation sites exist, the majority of circulating 17β-estradiol is metabolized through these two mutually exclusive pathways. Originally, it was thought that the two metabolites (2-hydroxyestrone [2-OHE1] and 16α-hydroxyestrone [16α-OHE1]) generated through these pathways were without biological activity and simply represented breakdown products. This belief, however, has not been supported by subsequent in vivo and cell culture studies.

16α-OHE1 is an estrogen agonist in some model systems. It stimulates cell proliferation and differentiation in cultured murine and human breast epithelial cells,3,4 increases uterine weight in ovariectomized (OVX) rats,3 and alters gonadotropin secretion in OVX rats,3 similar to 17β-estradiol. Its affinity for sex hormone binding globulin is low, rendering it available in circulation to produce estrogenic effects, and it has the capacity to bind covalently to nuclear estrogen receptors.5 In contrast, 2-OHE1 is not an estrogen agonist6 and it has been reported to have antiestrogenic activity.4,7 By tightly binding to the estrogen receptor, 2-OHE1 may prevent other circulating estrogens from binding and interacting with the receptor to produce their biological effect.4

Originally it was suggested that the metabolites of 17β-estradiol and estrone may predict the risk of breast8 and other hormone-related cancers.9 Specifically, it has been reported that higher 16α-OHE1 levels, or alternatively a greater proportion of estrogens metabolized via 16α-hydroxylation, may confer an increased risk for developing breast2,10,11 and cervical9 cancer. In animal models, mice that spontaneously develop mammary tumors have increased 16α-hydroxylation of estrogen,12 and in the few human studies performed to date, most,2,10,11,13 but not all,14,15 indicate that breast cancer patients metabolize a greater percentage of their estrogen through the C-16α-pathway. 2-hydroxylation has been suggested to confer a protective effect against breast cancer.16

Recently, it has been suggested that women who proportionally metabolize more estrogen through the 2-hydroxylation pathway and less through the 16α-hydroxylation pathway are at greater risk for developing osteoporosis.17–19 In a small study of Korean women, Lim and colleagues reported that 16α-OHE1 was lower in osteopenic subjects than in nonosteopenic subjects and 2-OHE1 was significantly higher, although there were no differences in serum estradiol or estrone levels. Further, the ratio of the two metabolites (16α/2-OHE1) was positively correlated with spinal bone mineral density (BMD). Other investigators have reported that low levels 16α-OHE1 are associated with an increased rate of bone loss.19 This association is supported by epidemiological data, demonstrating an inverse relationship between breast cancer and osteoporosis risk.20,21 Women in the highest quartiles of BMD had a significantly increased relative risk of developing breast cancer. The data suggest a biological link, perhaps estrogen metabolism, between these two diseases.

The skeleton is well recognized as an estrogen-responsive tissue. The presence of estrogen receptors and rapid responses to the hormone suggests that estrogen has direct actions on the skeleton.22–24 As discussed in recent reviews,25,26 it is well established that the decline in ovarian hormone production with menopause results in bone loss and increased risk for osteoporotic fractures. However, treatment with estrogen inhibits bone loss and reduces fracture risk.25,26 The relationship between estrogen metabolism and the risk of developing osteoporotic fractures is largely unknown. The purpose of the present investigation was to utilize the OVX rat model to assess the estrogen agonism of the two metabolites, 2-OHE1 and 16α-OHE1, on the skeleton and other estrogen-responsive tissues. These effects were compared with OVX rats receiving 17β-estradiol, no treatment, as well as ovary-intact animals.

MATERIALS AND METHODS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. References

Animals

All procedures were approved by the Institutional Animal Care and Use Committee at the AMC Cancer Research Center before beginning the study.

Fifty-four Sprague-Dawley rats were obtained from Taconic Farms (German Town, NY, U.S.A.) at ∼8 weeks of age (mean [± SE] body weight [bw] = 160 ± 3 g). Bilateral ovariectomy (n = 48, OVX) or sham surgery (n = 6, intact) was performed at 9 weeks of age. Two days postsurgery, OVX rats were stratified by weight and randomized to a baseline (n = 8) or one of four treatment groups: 2-OHE1 (n = 10), 16α-OHE1 (n = 10), 17β-estradiol (n = 10), or vehicle (n = 10, OVX-vehicle).

Treatment was not started until 1 week post-OVX to allow for estrogen levels to become depleted. Treatment consisted of daily subcutaneous injections of 2-OHE1, 16α-OHE1, 17β-estradiol, or vehicle (50% ethanol). The dose was set at 200 μg/kg based on an initial group mean bw (180 g) and was not altered over the treatment period as the animals gained weight. This supraphysiological dose was chosen to ensure saturation of estrogen receptor binding sites with each metabolite.3,4,27 2-OHE1, 16α-OHE1, and 17β-estradiol were obtained from Sigma Chemical Co. (St. Louis, MO, U.S.A.) and mixed to appropriate concentrations in 50% ethanol. Ascorbic acid (1 mg/ml) was added in the preparation of 2-OHE1 to prevent oxidation of this labile metabolite. Animals were injected with 200 μl of the individual treatment at 8:00 a.m. daily. Treatment duration was 3 weeks.

Fluorochromes to label mineralizing bone matrix were administered by juxta-tail vein injection 1 day before starting treatment (tetracycline-HCl, 20 mg/kg of bw; Sigma), 7 days before sacrifice (calcein, Sigma; 20 mg/kg of bw) and 1 day before sacrifice (tetracycline, as described). The two fluorochrome labels are readily differentiated under UV illumination, because tetracycline fluoresces pale yellow, while calcein fluoresces bright green.

Animals were sacrificed by CO2 inhalation 24 h after their last treatment injection. Baseline animals were sacrificed on the day that treatment was started. Blood was collected via the periorbital route for measurement of serum cholesterol and estrogen metabolite levels. The uterus was excised for wet weight determination and for measurement of epithelial cell height. Mammary glands were excised for histologic evaluation and proliferating cell nuclear antigen (PCNA) immunohistochemistry quantitation. The right tibia was removed and fixed in 70% ethanol for static and dynamic histomorphometry.

Bone histomorphometry

Histomorphometric measurements were performed with the SMI-Microcomp-P.M. semiautomatic image analysis system (Southern Micro Instruments, Inc., Atlanta, GA, U.S.A.), which consists of a computer (Compak 285, Compaq Computer Corp., Houston, TX, U.S.A.) coupled to a photomicroscope and image analysis system.

Longitudinal growth rate

Longitudinal growth rate was measured in the proximal tibial metaphysis and is the mean distance between the calcein-labeling front located in the primary and secondary spongiosa and the final tetracycline label in the mineralizing growth plate cartilage divided by the labeling interval of 6 days.

Cortical bone measurements

Ground transverse sections were used for histomorphometric analysis of cortical bone. Cross-sections 150-μm-thick were cut at a site just proximal to the tibia–fibula synostosis with a low-speed saw (Isomet, Buehler, Lake Bluff, IL, U.S.A.) equipped with a diamond wafer blade. The sections were ground to a thickness of 15–20 μm on a roughened glass plate and mounted in glycerin before microscopic examination under UV illumination to visualize fluorochrome labeling. The following measurements were performed as described27: cross-sectional area, defined as the area of bone and marrow cavity bounded by the periosteal surface of the specimen; medullary area, defined as the area delineated by the endocortical surface of the specimen; cortical bone area, calculated as the difference between the cross-sectional and medullary area; periosteal perimeter, defined as the total perimeter enclosing the cross-section (periosteal perimeter includes fluorochrome-labeled and nonlabeled perimeters); periosteal bone formation rate, calculated as the area bounded by the tetracycline labels and divided by the labeling period of 21 days; and periosteal mineral apposition rate (MAR), defined as the periosteal bone formation rate divided by the label perimeter. These measurements have been described previously27 and conform to the standard nomenclature proposed by Parfitt et al.28

Cancellous bone measurements

The tibia was dehydrated in a series of increasing concentrations of ethanol, embedded without demineralization in a mixture of methylmethacrylate-2-hydroxyethyl-methacrylate (12.5:1) to retain the fluorochrome labels, and sectioned at a thickness of 5 μm.

The sampling site included the entire tibial epiphysis and represented an area ∼2.8 mm2. Measurements were performed as described.27 Cancellous bone area was determined as the area of total cancellous bone per square millimeter of epiphyseal area and expressed as a percentage. The cancellous bone perimeter was calculated as the perimeter of cancellous bone perimeter per square millimeter of epiphyseal area. The bone formation rate was calculated as the product of the double label surface and MAR. The MAR (μm/day) was the mean distance between the calcein and second tetracycline label divided by the labeling interval of 6 days. Double-labeled surface was determined as the bone surface with the calcein and second tetracycline labels.

Uterus

The uterus was weighed for an initial wet weight and then frozen in liquid nitrogen. Uteri were thawed in 10% neutral buffered formalin for 4–6 h, then transferred to 70% ethanol until processing for conventional paraffin embedding. Five-micrometer sections were cut and stained with hematoxylin and eosin (H&E) to measure epithelial height. Uterine epithelial height, expressed in micrometers, was measured at ×20 magnification with an Olympus BH-2 microscope (Olympus, New Hyde Park, NY, U.S.A.). A minimum of 20 sites were measured from each section/animal.

Mammary glands

The right and left mammary gland chains were removed. Lymph node regions were removed and fixed in 10% neutral buffered formalin. These regions were used for histologic evaluation with standard H&E staining and PCNA immunohistochemistry.

PCNA marks cells in S-phase and is an index of proliferation. Five-micrometer tissue sections were deparaffinized in xylene and rehydrated through a descending series of alcohol to water. Antigen retrieval was used via a citrate buffer (pH 8.0). After heating and addition of distilled water, endogenous peroxidase activity was blocked using 3% hydrogen peroxide. The primary antibody, PCNA (Dako, Carpenteria, CA, U.S.A.) was applied at a dilution of 1:50 for 60 minutes. The secondary antibody, a biotinylated rabbit anti-mouse antibody, was diluted 1:200 in 10% normal rabbit serum and incubated for 30 minutes. Finally, Streptavidin-HRP (Dako) at a dilution of 1:1000 was applied to the sections, which were then incubated for 30 minutes. The PCNA signal was visualized using 3,3′-diaminobenzidine. All washes were performed in modified phosphate-buffered saline (0.04 M K2HPO4, 0.01 M NaH2PO4, 0.13 M NaCl). Sections were lightly counterstained in Harris hematoxylin followed by an ascending series of alcohol and mounted with Permount (Fischer Scientific, Pittsburgh, PA, U.S.A.). Quantitation was performed by photographing five representative sections/animal (×400 magnification), counting stained and unstained cells (∼150–200 total cells/photograph), and calculating the percentage of those stained with the antigen.

Serum estrogen metabolites

Concentrations of circulating 2-OHE1 and 16α-OHE1 were determined by a direct monoclonal antibody-based competitive enzyme immunoassay modified by the manufacturer from that published for the measurement of urinary estrogen metabolites.29 All samples were assayed in duplicate with appropriate controls and standards run with each assay. Assay sensitivities for serum 2-OHE1 and 16α-OHE1 are <10 pg/ml, and the intra- and interassay coefficients of variation were 5% and 12%, respectively.

Serum cholesterol

Blood samples were allowed to clot at room temperature for 2 h, and serum was obtained after centrifugation at 2000 rpm for 15 minutes. Serum samples were stored at −80°C until analysis. Serum cholesterol was determined using a Cobas-Mira high-performance cholesterol assay with Roche Reagents. Briefly, serum cholesterol is released from its esters by enzymatic action of an ester hydrolase and then oxidized by cholesterol oxidase to produce hydrogen peroxide. The hydrogen peroxide, when combined with 4-aminoantipyrine and phenol, forms a chromophore which is visible at 500 nm and is directly proportional to the cholesterol concentration.

Statistical analysis

Analyses of variance were performed on all variables. Student–Neuman–Keuls post hoc multiple comparison tests were performed to assess between group differences when appropriate. A p value of ≤0.05 was designated as statistically significant.

RESULTS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. References

The effect of OVX, 17β-estradiol, 2-OHE1, and 16α-OHE1 on bw, serum cholesterol, and longitudinal growth rate, are presented in Table 1. OVX resulted in a significant increase in bw, serum cholesterol, and tibial longitudinal growth rate. Treatment with 17β-estradiol prevented the OVX-induced weight gain and increased the tibial longitudinal growth rate; values were not different from those of the intact group. No change in serum cholesterol levels was observed in 17β-estradiol–treated rats relative to the OVX group. 2-OHE1 had no effect on bw, serum choesterol or longitudinal growth rate. Values were not significantly different from those of the OVX animals. 16α-OHE1 treatment reduced the OVX-induced increase in bw but not as much as 17β-estradiol treatment; weight gain in the 16α-OHE1 group was significantly greater than the intact animals. The longitudinal growth rate of the 16α-OHE1 group was not different from the intact or 17β-estradiol–treated group and was significantly less than the OVX group. Similarly, cholesterol levels in 16α-OHE1–treated rats were significantly lower than the OVX or 17β-estradiol–treated animals and did not differ from the intact animals.

Table Table 1. Effects of OVX, 17β-Estradiol, 2-OHE1, and 16α-OHE1 on Body Weight, Serum Cholesterol, and Longitudinal Growth Rate at the Proximal Tibial Metaphysis
Thumbnail image of

Uterine weight and uterine epithelial height data are presented in Table 2. OVX resulted in a significant decrease in wet weight and epithelial height relative to the intact animals. 17β-estradiol treatment resulted in uterine weight comparable to the intact animals and epithelial height 70% larger than the intact group. 16α-OHE1 treatment resulted in an intermediate response; uterine weight was increased compared with the OVX-vehicle–treated group but decreased compared with the intact and 17β-estradiol–treated groups. Epithelial height in 16α-OHE1–treated rats was significantly greater than OVX, not different from the intact and significantly less than the 17β-estradiol–treated group. 2-OHE1 treatment had no effect on uterine weight or epithelial cell height. Photomicrographs, at ×400 magnification, illustrate the histologic differences in the uterus between the treatment groups (Fig. 1).

Table Table 2. Effects of OVX, 17β-Estradiol, 2-OHE1, and 16α-OHE1 on Uterine Weight, Uterine Epithelial Height, and Mammary Gland Proliferating Cell Nuclear Antigen Labeling
Thumbnail image of
thumbnail image

Figure FIG. 1. Photomicrographs (×400) of representative sections of rat uterine epithelia. Paraffin embeded uteri were sectioned (5 μm) and stained with H&E to measure epithelial cell height. Treatment groups are: (A) OVX-vehicle (50% ethanol), (B) 17β-estradiol (200 μg/kg of bw), (C) 2-OHE1 (200 μg/kg of bw), (D) 16α-OHE1 (200 μg/kg of bw).

Download figure to PowerPoint

PCNA labeling was significantly increased in the mammary glands excised from the 17β-estradiol–treated animals but not the 16α-OHE1 or 2-OHE1 treatment groups compared with the OVX-vehicle group. PCNA labeling (mean ± SE) in the OVX-vehicle group (1.4 ± 0.5%) was not significantly different from the intact group (4.2 ± 2.1%; p = 0.15) because of the estrous cycle phase-associated variability in the intact animals (range of labeled cells 0–13%). Figure 2 provides representative H&E-stained photomicrographs (×100) of mammary gland sections from the OVX, 17β-estradiol, 2-OHE1, and 16α-OHE1–treated rats. The 17β-estradiol group had stimulated mammary glands with increased lobular and alveolar development. Mammary glands from the OVX-vehicle and 2-OHE1 animals were regressed, displaying little lobular/alveolar development and resembling sexually immature virgin rats.30 Mammary glands from the 16α-OHE1 group were intermediate in appearance between OVX and 17β-estradiol groups.

thumbnail image

Figure FIG. 2. Photomicrographs (×100) of representative rat mammary gland sections. Lymph node regions were removed, fixed, and stained with H&E as described in the Materials and Methods section. Treatment groups are: (A) OVX-vehicle (50% ethanol), (B) 17β-estradiol (200 μg/kg of bw), (C) 2-OHE1 (200 μg/kg of bw), (D) 16α-OHE1(200 μg/kg of bw).

Download figure to PowerPoint

Cortical bone histomorphometry data are shown in Table 3. There were no significant differences in static bone measurements, including cross-sectional area, medullary area, or cortical bone area among the intact, OVX, and the three estrogen-treated groups. There was an age-related change in all five groups relative to the baseline animals in the cross-sectional and cortical bone area and periosteal perimeter expansion. OVX resulted in an increase in the periosteal bone formation rate and MAR, which was prevented by 17β-estradiol treatment. 16α-OHE1 treatment also resulted in prevention of the increased bone formation and MAR, comparable to the 17β-estradiol treatment. 2-OHE1 had no effect on the OVX-induced increase in bone formation or MAR.

Table Table 3. Effects of OVX, 17β-Estradiol, 2-OHE1, and 16α-OHE1 on Cortical Bone Histomorphometry
Thumbnail image of

The high rate of longitudinal bone growth prevented measurement of cancellous bone remodeling in the proximal tibial metaphysis. In contrast, the epiphysis is skeletally mature bone. Although OVX does not generally result in loss of cancellous bone from the epiphysis, bone turnover is greatly increased at that site.31 Cancellous bone histomorphometry data from the tibial epiphysis are presented in Table 4. There were no significant differences among the five groups in cancellous bone area or bone perimeter in this 21-day study. In contrast, bone formation rate, whether expressed relative to total tissue volume, bone volume, or bone surface, was significantly increased with OVX. OVX-induced changes were prevented by treatment with 17β-estradiol or 16α-OHE1. Treatment with 2-OHE1 had no effect on OVX-associated increase in bone formation. 16α-OHE1 treatment resulted in bone measurements that did not differ from those of the 17β-estradiol–treated group. MAR was slightly but significantly increased after OVX. 17β-estradiol treatment prevented this increase; 16α-OHE1 was less effective at mitigating the increase in MAR associated with OVX. The surface of bone covered by double labels was significantly increased with OVX and was prevented by treatment with 17β-estradiol or 16α-OHE1.

Table Table 4. Effects of OVX, 17β-Estradiol, 2-OHE1, and 16α-OHE1 on Cancellous Bone Histomorphometry
Thumbnail image of

DISCUSSION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. References

OVX resulted in the expected uterine and mammary gland atrophy while increasing serum cholesterol and bw gain. Regarding bone, longitudinal and radial bone growth were accelerated and cancellous bone turnover was elevated. These findings have been previously documented by numerous studies.27,31–33

Treatment of OVX animals with 17β-estradiol also resulted in the expected changes.27,32,34 Uterine weight was increased and epithelial height was significantly greater than that of the ovary-intact animals. 17β-estradiol treatment prevented the OVX-associated increases in bone growth and turnover; bone formation and MARs were significantly less than the OVX-vehicle animals and were comparable to, or less than, those of the intact animals. 17β-estradiol treatment failed to result in an attenuation of the OVX-induced increase in serum cholesterol. Previous studies have indicated that high doses of estrogen (and estrogen agonists) do not necessarily result in cholesterol lowering and may result in an increase in serum cholesterol.35,36

Treatment of OVX animals with 2-OHE1 had no effect on any of the estrogen target tissues evaluated. Serum measurements for 2-OHE1 confirmed that it was present at high concentrations in the circulation (637 ± 79 pg/ml), in comparison with the intact (52 ± 30 pg/ml) and OVX animals (12 ± 12 pg/ml), even 24 h after the final subcutaneous injection. Therefore, rapid clearance or methylation of 2-OHE1 to 2-methyl OHE1 cannot explain the lack of estrogen activity. The data do confirm previous reports in estrogen-receptor-positive cell culture lines that 2-OHE1 appears to have no estrogen agonistic activity.4,7 It is not possible to determine whether 2-OHE1 has estrogen antagonistic effects within the present experimental design; this is currently under investigation. It should be noted that ascorbic acid was added in the preparation of 2-OHE1 to inhibit oxidation of the metabolite. Ascorbic acid has been reported to affect collagen synthesis and mineralization in cell37 and organ cultures38 systems with results varying depending on dose, timing, and cell stage. We speculate that any effect of this addition of ascorbic acid would be minimal, although to definitively substantiate this the use of a second control group (vehicle + ascorbic acid) would be warranted.

Overall, the data indicate that 16α-OHE1 is acting as a full estrogen agonist on the bone and a partial estrogen agonist in the mammary gland and uterus. To our knowledge, this is the first report of the effects of these naturally occurring estrogen metabolites on the skeleton and the first evidence of tissue specificity of an endogenous estrogen.

Previous short-term studies (3 days) of weanling rats suggested that 16α-OHE1 is a complete estrogen agonist on uterine wet weight.6 Longer term treatment (3 weeks) of sexually mature OVX rats with the 16α-OHE1 metabolite resulted in no significant activity on PCNA labeling of the mammary gland, partial agonistic activity on uterine weight, uterine epithelial cell height and bw, and potent agonistic activity on serum cholesterol and bone measurements. These short- and long-term responses are remarkably similar to those induced by triphenylethylene and benzothiophene antiestrogens.35,36,39–41

Dose–response studies are required to determine the potency of 16α-OHE1. However, the potency of this estrogen metabolite is similar or greater than tamoxifen, droloxifene, and raloxifene.35,36,39,40 Skeletally mature rats are needed to determine whether 16α-OHE1, protects against OVX-induced bone loss. The purpose of this initial study was first to determine whether there was any effect of the estrogen metabolites on the skeleton and second to be able to measure these potential effects on all aspects of bone physiology, i.e., longitudinal and radial growth, modeling, and remodeling. This necessitated the use of young growing animals.

17β-estradiol, as well as pharmacological estrogen agonists/antagonists, elicit very similar effects in the skeletons of rats as they do in humans.25,26,42–44 Given the similarity between species, the present results, which demonstrate no effect of 2-OHE1 and a potent estrogen agonism by 16α-OHE1 on the OVX rat skeleton, suggest that women who metabolize a greater proportion of their estradiol/estrone through the C-2 pathway would be at greater risk for rapid bone loss, osteoporosis, and/or fractures. In this regard, recent studies demonstrate negative relationships between 2-OHE117,18 and BMD, and positive association between 16α-OHE1 and BMD17 in postmenopausal women. Conversely, women with more 16α-hydroxylation might be at lower risk for osteoporosis. It may be hypothesized that women who metabolize estrogens more through 16-hydroxylation have a greater “estrogenic environment” or more “estrogen exposure.” This environment would foster greater bone mass and/or attenuation of bone lost at menopause, thus resulting in reduced propensity for osteoporotic fracture. However, the increased estrogenic environment may favor mitogenic activity in the breast and/or other reproductive tissues, thus resulting in increased risk for cancer development. Women with greater bone density are at elevated risk for developing breast cancer.20,21

In summary, this is the first study to evaluate the effect of 2-OHE1 and 16α-OHE1 on the skeleton. 2-OHE1 had no activity on the skeleton of OVX rats. In contrast, 16α-OHE1 appears to be a naturally produced partial estrogen agonist with tissue specificity. The data suggest that the effects of antiestrogens, such as tamoxifen and raloxifene, may be mimicking normal biological pathways. The potential to modulate the hydroxylation pathways through pharmacological means and/or lifestyle changes may provide avenues by which women can modify their risk for developing osteoporosis, breast cancer, coronary heart disease, and/or other estrogen-associated diseases. Further investigation into the area of estrogen metabolism and osteoporosis risk is clearly implicated.

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. References

The authors acknowledge Dr. Robert Strange for his scientific input on mammary gland biology. Ms. Lori M. Rolbiecki is gratefully acknowledged for her secretarial assistance. These studies were supported in part by the AMC Cancer Research Center and by National Institutes of Health grant AR41418.

References

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. References
  • 1
    Yen SSC, Jaffe RB 1991 Reproductive Endocrinology, 3rd ed, W.B. Saunders, Philadelphia, PA, U.S.A.
  • 2
    Schneider J, Kinne D, Fracchia A, Pierce V, Anderson KE, Bradlow HL, Fishman J 1982 Abnormal oxidative metabolism of estradiol in women with breast cancer Proc Natl Acad Sci USA 79:30473051.
  • 3
    Fishman J, Martucci C 1980 Biological properties of 16α-hydroxyestrone: Implications in estrogen physiology and pathophysiology J Clin Endocrinol Metab 51:611615.
  • 4
    Schneider J, Huh MM, Bradlow HL, Fishman J 1984 Antiestrogen action of 2-hydroxyestrone on MCF-7 human breast cancer cells J Biol Chem 259:48404845.
  • 5
    Swaneck GE, Fishman J 1988 Covalent binding of the endogenous estrogen 16α-hydroxyestrone to estradiol receptor in human breast cancer cells: Characterization and intranuclear localization Proc Natl Acad Sci USA 85:78317835.
  • 6
    Martucci CP, Fishman J 1979 Impact of continuously adminstered catechol estrogens on uterine growth and luteinizing hormone secretion Endocrinology 105:12881292.
  • 7
    Vandewalle B, Lefebvre J 1989 Opposite effects of estrogen and catecholestrogen on hormone-sensitive breast cancer cell growth and differentiation Mol Cell Endocrinol 61:239246.
  • 8
    Bradlow HL, Michnovicz JJ 1989 A new approach to the prevention of breast cancer Proc Royal Soc Edinburgh 95B:77.
  • 9
    Sepkovic DW, Bradlow HL, Ho G, Hankinson SE, Gong L, Osborne MP, Fishman J 1995 Estrogen metabolite ratios and risk assessment of hormone-related cancers: Assay validation and prediction of cervical cancer risk Ann NY Acad Sci 768:312316.
  • 10
    Osborne MP, Bradlow HL, Wong GYC, Telang NT 1993 Upregulation of estradiol C16α-hydroxylation in human breast tissue: A potential biomarker of breast cancer risk J Natl Cancer Inst 85:19171920.
  • 11
    Kabat GC, Chang CJ, Sparano JA, Sepkovic DW, Hu XP, Khalil A, Rosenblatt R, Bradlow HL 1997 Urinary estrogen metabolites and breast cancer: A case-control study Cancer Epidemiol Biomarkers Prev 6:505509.
  • 12
    Bradlow HL, Hershcopf RJ, Martucci CP, Fishman J 1985 Estradiol 16α-hydroxylation in the mouse correlates with mammary tumor incidence and presence of murine mammary tumor virus: A possible model for the hormonal etiology of breast cancer in humans Proc Natl Acad Sci USA 82:62956299.
  • 13
    Coker AL, Crane MM, Sticca RP, Sepkovic DW 1997 Re: Ethnic differences in estrogen metabolism in healthy women J Natl Cancer Inst 89:8990.
  • 14
    Adlercreutz H, Fotsis T, Hockerstedt K, Hamalainen E, Bannwart C, Bloigu S, Valtonen A, Ollus A 1989 Diet and urinary estrogen profile in premenopausal omnivorous and vegetarian women and in premenopausal women with breast cancer J Steroid Biochem 34:527530.
  • 15
    Adlercreutz H, Gorbach SL, Goldin BR, Woods MN, Dwyer JT, Hamalainen E 1994 Estrogen metabolism and excretion in Oriental and Caucasian women J Natl Cancer Inst 86:10761082.
  • 16
    Bradlow HL, Telang NT, Sepkovic DW, Osborne MP 1996 2-Hydroxyestrone: The “good” estrogen J Endocrinol 150:S259S265.
  • 17
    Lim SK, Won YJ, Lee JH, Kwon SH, Lee EJ, Kim KR, Lee HC, Hah KB, Chung BC 1997 Altered hydroxylation of estrogen in patients with postmenopausal osteopenia J Clin Endocrinol Metab 82:10011006.
  • 18
    Hodge J, Roodman-Weiss J, Lyss C, Wagner D, Klug T, Civitelli R 1995 Increased inactive estrogen metabolites in urine of early postmenopausal women with low bone density J Bone Miner Res 10:S444.
  • 19
    Leelawattana R, Ziambaras K, Lyss C, Roodman-Weiss J, Wagner D, Klug T, Civitelli R 1997 Estrogen metabolites in urine of early postmenopausal women correlate with bone density and rate of bone loss J Bone Miner Res 12:S132.
  • 20
    Cauley JA, Lucas FL, Kuller LH, Vogt MT, Browner WS, Cummings SR 1996 Bone mineral density and risk of breast cancer in older women J Am Med Assoc 276:14041408.
  • 21
    Zhang Y, Kiel DP, Kreger BE, Cupples LA, Ellison RC, Dorgan JF, Schatzkin A, Levy D, Felson DT 1997 Bone mass and the risk of breast cancer among postmenopausal women N Engl J Med 336:611617.
  • 22
    Westerlind KC, Sarkar G, Bolander ME, Turner RT 1995 Estrogen receptor mRNA is expressed in vivo in rat calvarial periosteum Steroids 60(8):484487.
  • 23
    Turner RT, Backup P, Sherman PH, Hill E, Evans GL, Spelsberg TC 1992 Mechanism of action of estrogen on intramembranous bone formation: Regulation of osteoblast differentiation and activity Endocrinology 131:883889.
  • 24
    Hoyland JA, Mee AP, Baird P, Braidman IP, Mawer EB, Freemont AJ 1997 Demonstration of estrogen receptor mRNA in bone using in situ reverse-transcriptase polymerase chain reaction Bone 20:8792.
  • 25
    Turner RT, Riggs BL, Spelsberg TC 1994 Skeletal effects of estrogen Endocr Rev 15:275300.
  • 26
    Gallagher JC 1996 Estrogen: prevention and treatment. In: MarcusR, FeldmanD, and KelseyJ (eds.) Osteoporosis, Academic Press, New York, NY, U.S.A.
  • 27
    Westerlind KC, Wakley GK, Evans GL, Turner RT 1993 Estrogen does not increase bone formation in growing rats Endocrinology 133:29242934.
  • 28
    Parfitt AM, Drezner MK, Glorieux FH, Kanis JA, Malluche H, Meunier PJ, Ott SM, Recker RR 1987 Bone histomorphometry: Standardization of nomenclature, symbols, and units J Bone Miner Res 2:595610.
  • 29
    Klug TL, Bradlow HL, Sepkovic DW 1994 Monoclonal antibody-based enzyme immunoassay for simultaneous quantitation of 2- and 16α-hydroxyestrone in urine Steroids 59:648655.
  • 30
    Pitelka DR 1988 The mammary gland. In: WeissL (ed.) Cell and Tissue Biology. Urban & Schwarzenberg, Baltimore, MD, U.S.A., pp. 877898.
  • 31
    Wronski TJ, Cintron M, Dann LM 1988 Temporal relationship between bone loss and increased bone turnover in ovariectomized rats Calcif Tissue Res 43:179183.
  • 32
    Turner RT, Vandersteenhoven JJ, Bell NH 1987 The effects of ovariectomy and 17β-estradiol on cortical bone histomorphometry in growing rats J Bone Miner Res 2:6166.
  • 33
    Wronski TJ, Walsh CC, Iganszewski LA 1986 Histological evidence for osteopenia and increased bone turnover in ovariectomized rats Bone 7:119123.
  • 34
    Wronski TJ, Dann LM, Scott KS, Crooke LR 1989 Endocrine and pharmacological suppressors of bone turnover protect against osteopenia in ovariectomized rats Endocrinology 125:810816.
  • 35
    Evans GL, Bryant HU, Magee DE, Turner RT 1996 Raloxifene inhibits bone turnover and prevents further cancellous bone loss in adult ovariectomized rats with established osteopenia Endocrinology 137:41394144.
  • 36
    Ke HZ, Simmons HA, Pirie CM, Crawford DT, Thompson DD 1995 Droloxifene, a new estrogen antagonist/agonist, prevents bone loss in ovariectomized rats Endocrinology 136:24352441.
  • 37
    Denis I, Pointillart A, Lieberherr M 1994 Cell stage-dependent effects of ascorbic acid on cultured porcine bone cells Bone Miner 25:149161.
  • 38
    Ganta DR, McCarthy MB, Gronowicz GA 1997 Ascorbic acid alters collagen integrins in bone culture Endocrinology 138:36063612.
  • 39
    Turner RT, Wakley GK, Hannon KS, Bell NH 1988 Tamoxifen inhibits osteoclast-mediated resorption of trabecular bone in ovarian hormone-deficient rats Endocrinology 122:11461150.
  • 40
    Evans GL, Bryant HU, Magee D, Sato M, Turner RT 1994 The effects of raloxifene on tibia histomorphometry in ovariectomized rats Endocrinology 134:22832288.
  • 41
    Jimenez MA, Magee DE, Bryant HU, Turner RT 1997 Clomiphene prevents cancellous bone loss from tibia of ovariectomized rats Endocrinology 138:17941800.
  • 42
    Love RR, Mazess RB, Tormey DC, Barden HS, Newcomb PA, Jordan VC 1988 Bone mineral density in women with breast cancer treated with adjuvant tamoxifen for at least two years Breast Cancer Res Treat 12:297301.
  • 43
    Fornander T, Rutqvist LE, Sjoberg HE, Blomqvist L, Mattsson A, Glas U 1990 Long-term adjuvant tamoxifen in early breast cancer: Effect on bone mineral density in postmenopausal women J Clin Oncol 8:10191024.
  • 44
    Mack TM, Ross RK 1989 Risks and benefits of long-term treatment with estrogens Schweiz Med Wochenschr 119:18111820.