OBJECTIVE: To clarify histologic localization of estrone sulfatase in normal uterine endometrium and adenomyotic tissue and to confirm that estrone sulfatase is one of the enzymes that supplies estrogen to adenomyotic tissue.
METHODS: Specimens from 21 patients who had undergone hysterectomy were obtained from uteri with histopathologically proven adenomyosis. Specimens from 28 patients who had undergone hysterectomy for a disease of the uterine cervix were used as control specimens of normal uterine endometrium. Cases of hormone‐dependent disease, such as leiomyoma, adenomyosis, and endometrial neoplasm, were excluded from cases of normal endometrium. The myometrium in patients with adenomyosis was examined. These tissues were examined by immunohistochemistry using anti‐estrone sulfatase monoclonal antibodies. Power analysis was performed. With α = 0.05, 1 − β = 0.8, P1= 25%, and P2 = 75%, 14 specimens from each group were sufficient to detect significant differences among them. The Fisher exact test, sign test, and McNemar test were used for statistical analysis.
RESULTS: In normal endometrial tissue, immunostaining for estrone sulfatase was observed only on the glandular epithelial cells of the basilar layer of the endometrium. However, all functional layers of the endometria were negative for staining for estrone sulfatase. In adenomyotic tissue, glandular epithelial cells showed immunostaining for estrone sulfatase. Rates of immunostaining in adenomyotic tissue were higher than those in the basilar layer of normal uterine endometrium (76% and 43%, respectively, P = .02). The myometrium was not stained.
CONCLUSION: Estrone sulfatase may be one of the enzymes supplying estrogen for growth of adenomyosis.
Adenomyosis is considered an estrogen‐dependent disease similar to breast cancer, endometriosis, endometrial cancer, and uterine leiomyoma.1–4 Gonadotropin‐releasing hormone (GnRH) agonist, which suppresses serum estrogen levels, is used to treat adenomyosis. It is clinically known that suppression of serum estrogen levels in patients with adenomyosis retards the progress of this disease. Estrogen is thus closely associated with the growth and development of adenomyosis.
There are two sources of estrogen in estrogen‐dependent disease. One is secretion by the ovaries and delivery to target tissues through the bloodstream, and the other is biosynthesis in local tissue catalyzed by enzymes such as aromatase and estrone sulfatase. Estrone sulfatase catalyzes the conversion of estrone sulfate to estrone, which is further transformed to the potent estrogen estradiol (E2) by the enzyme 17‐hydroxysteroid dehydrogenase (17β‐HSD). Estrone sulfate is the most abundant estrogen in plasma, with serum levels of this conjugated steroid reported to be two to ten times higher than levels of estrone.5,6 Estrone sulfatase appears to be a major source of estrogen for growth of estrogen‐dependent disease.7,8
The levels of estrone sulfatase activity in normal endometrium and adenomyotic tissue have been reported previously.4 However, the histologic localization of estrone sulfatase is not known, because no anti‐estrone sulfatase monoclonal antibodies have been available for use in immunohistochemistry.
Recently, anti‐estrone sulfatase monoclonal antibodies (KM1049) originating from human placenta have been produced for immunohistochemistry.9 In the present study, we demonstrate the histologic localization of estrone sulfatase in both normal uterine endometria and adenomyotic tissue in order to confirm that estrone sulfatase in adenomyosis is a source of estrogen.
MATERIALS AND METHODS
The Institutional Review Board at The Jikei University School of Medicine, Tokyo, Japan, approved this study. Informed consent was obtained from all patients undergoing surgical treatment.
Surgical specimens obtained from 21 patients who had undergone hysterectomy were histopathologically diagnosed with adenomyosis. Normal endometrial tissue was taken from the normal portion of the uteri of 28 patients with cervical cancer stage I or carcinoma in situ. Patients with normal endometrium were 29–49 years old, and patients with adenomyosis 27–48 years old. Normal endometrium was not obtained from patients with estrogen‐dependent diseases such as uterine leiomyoma, adenomyosis, endometriosis, or endometrial neoplasm, because we had concerns that estrogen expression from such estrogen‐dependent diseases could affect normal endometrium. No patients with adenomyosis received hormonal treatment for 6 months before hysterectomy. All patients had regular menstrual cycles. Cases were divided into two groups, the proliferative phase and the secretory phase, by examining endometria histologically. The myometrium in patients with adenomyosis was also studied.
All patients who were treated for adenomyosis and carcinoma in situ stage I uterine cervical cancer by hysterectomy at Jikei University Hospital from January 1996 to December 1998 were included in this study. However, we excluded patients who had received the hormonal therapy or had estrogen‐dependent disease.
To determine moderately group‐specific sample size, we performed power analysis as follows. We assumed that adenomyotic tissue exhibited staining and endometrium did not. Statistically, under the condition, α = 0.05, 1 − β = 0.8, P1= 25%, P2 = 75%, 14 specimens in each group were sufficient to detect significant differences between them. McNemar test was used to compare the presence of immunostaining between functional and basilar layers in each specimen. The sign test was used to compare the degree of immunostaining between functional and basilar layers. Fisher exact test was used to compare the presence of immunostaining between adenomyotic tissue and normal endometrium, because patients with adenomyosis were different from those with normal endometrium. P values less than .05 were considered statistically significant.
Immunohistochemical staining was performed as follows. The tissues were fixed immediately with buffered 10% formalin for 24–72 hours, dehydrated, and embedded in paraffin. Before use, 4‐μm sections were cut and deparaffinized in ethanol and xylene. Immunohistochemical staining was performed with the use of an immunoperoxidase avidin‐biotin conjugate system, with diaminobenzidine and hydrogen peroxide as the substrate and hematoxylin as the counterstain. Slides were rinsed in a phosphate buffer (pH 7.4) for examination. The sections were incubated with primary monoclonal antibodies (KM1049). All primary antibodies were titrated by dilution (1:500 with anti‐estrone sulfatase monoclonal antibodies) to obtain optimal intensity of specific staining with minimal nonspecific background reactivity. The secondary or link antibody was a biotinylated horse anti‐mouse immunoglobulin (Vector Laboratories, Inc., Burlingame, CA) for use with primary murine monoclonal antibodies. Negative controls initially consisted of tissue processed without inclusion of the primary antibody.
We examined the slides by light microscopy. All slides were evaluated according to the intensity of immunostaining. We classified the intensity of the immunostaining into four levels, ranging from 0 = negative staining, 1 = lower intensity, 2 = moderate intensity, and 3 = the highest intensity in all slides. To make the results simple, we divided these levels into two groups, positive staining and negative staining. Cases classified as 0 or 1 were defined as negative staining, and cases classified as 2 or 3 were defined as positive staining. Among the cases classified as 2 or 3, if the frequency of immunostained cells was under 50%, they were defined as negative staining.
Figure 1 shows immunohistochemical staining with anti‐estrone sulfatase monoclonal antibodies of normal endometrium in the proliferative phase. Staining for estrone sulfatase was marked in the basilar layer of the endometrium, but no staining was found in the functional layer of endometrium (Figure 1A). Staining was detected in the cytoplasm of glandular cells, whereas no staining was observed in the stromal cells (Figure 1B).
Figure 2 shows immunohistochemical staining of normal endometrium in the secretory phase. Similarly, in the proliferative phase, staining was observed mainly in the basilar layer of the endometrium (Figure 2A). Only glandular epithelium was stained for estrone sulfatase (Figure 2B). There were no difference in expression of estrone sulfatase between the proliferative phase and the secretory phase.
In adenomyotic tissue, estrone sulfatase expression was observed. Figure 3 shows the adenomyotic tissue composed of glandular epithelium and stroma within the myometrium. Similarly, in normal endometrium, staining was detected only in the cytoplasm of glandular cells. The myometrium did not exhibit staining. In adenomyotic tissues, there was no difference in the rate of staining or localization of estrone sulfatase between the proliferative and secretory phases.
Incidences of specimens exhibiting staining are given in Table 1. The rate of immunostaining in adenomyotic tissue was significantly higher than that in normal endometrium (P = .02, Fisher exact test). The degree of immunostaining in the functional layer was significantly lower than that in the basilar layer (P = .001, sign test), and the rate of immunostaining in the functional layer also was significantly lower than that in the basilar layer (P = .001, McNemar's test)
At least seven different human sulfatases that hydrolyze sulfate ester groups (either sulfolipids or glycosaminoglycans) have been described.10 Steroid sulfatase is known to be a membrane‐bound, microsomal enzyme. Substantial similarities were found in the primary structures of four human arylsulfatases (arylsulfatases A, B, C, and D).11–13 Arylsulfatase C corresponds to steroid sulfatase. Therefore, all other anti‐steroid sulfatase monoclonal antibodies should be ruled out as a cross‐reaction with arylsulfatase.
The anti‐estrone sulfatase monoclonal antibody (KM1049) used in this study has high specificity and has no cross‐reaction with arylsulfatase A or B.9 Consequently, this monoclonal antibody can be used for immunohistochemical analysis of estrone sulfatase.
It has been thought that adenomyosis develops as a result of direct invasion of the myometrium by eutopic endometrial tissue; this tissue implants in the myometrium and grows there.4 In this study, estrone sulfatase expression was observed on both the basilar layer of endometrium and adenomyotic tissues. This suggests that both the basilar layer of the endometrium and adenomyotic tissue could supply estrogen resulting from catalysis by estrone sulfatase. The basilar layer of the endometrium and adenomyosis are similar histologically. Moreover, there may also be similarities between them in estrogen metabolism via estrone sulfatase.
Yamamoto et al4 found that estrone sulfatase activity in adenomyosis is higher than that in myometrium. However, estrone sulfatase activity is significantly lower in adenomyosis than in the endometrium. In the present study, estrone sulfatase expression was found in the glandular epithelium of adenomyosis but was not detected in the myometrium. The contrary results of the study by Yamamoto et al might have resulted from their inclusion of excessive myometrial tissue to obtain adenomyotic tissue. Homogenization of adenomyotic tissue with myometrial tissue could have been responsible for their findings. The present study histologically demonstrates the presence of estrone sulfatase in adenomyosis. Although the mere presence of estrone sulfatase is not definite proof of adenomyotic tissue to synthesize estrogen, those findings suggest that estrone sulfatase might be one of the enzymes supplying estrogen for growth of adenomyosis.
Danazol is one of the agents used to treat adenomyosis. Because danazol is an inhibitor of estrone sulfatase, it can inhibit biosynthesis of estrogen in adenomyotic tissue.4 In addition, several potent estrone sulfatase inhibitors have been reported to inhibit the growth of estrogen‐dependent breast cancer cells.14–17 Because estrone sulfatase inhibitors can specifically inhibit the enzymatic activity of estrone sulfatase, they could open new possibilities in the treatment of adenomyosis.
The mechanism of the regulation of estrone sulfatase is not completely understood. Currently, there is much interest in the effects that cytokines and growth factors might have in the development of estrogen‐dependent disease. Tumor necrosis factor‐alpha, insulin‐like growth factor‐1, and basic fibroblast growth factor are reported to stimulate sulfatase activity in a dose‐dependent manner in breast cancer.18,19 Further investigations to clarify the mechanism of regulation of estrone sulfatase will be needed to develop new treatments for adenomyosis.
1. Turunen A, Timonen S, Procope B. On the aetiology of endometriosis. Acta Obstet Gynaecol Scand 1961;40:206–22.
2. Yamamoto T, Takamori K, Okada H. Estrogen biosynthesis in leiomyoma and myometrium of the uterus. Horm Metabol Res 1984;16:678–9.
3. Tseng L, Mazella J, Mann WJ, Chumas J. Estrogen synthesis in normal and malignant human endometrium. J Clin Endocrinol Metab 1982;55:1029–31.
4. Yamamoto T, Noguchi T, Tamura T, Kitawaki J, Okada H. Evidence for estrogen synthesis in adenomyotic tissues. Am J Obstet Gynecol 1993;169:734–8.
5. Pasqualini JR, Gelly C, Nguyen BL, Vella C. Importance of estrogen sulfates in breast cancer. J Steroid Biochem 1989;34:155–63.
6. Santen RJ, Leszczynski D, Tilson-Mallet N, Feil PD, Wright C, Manni A, et al. Enzymatic control of estrogen production in human breast cancer: Relative significance of aromatase versus sulfatase pathways. Ann N Y Acad Sci 1986;464:126–37.
7. Yamamoto T, Kitawaki J, Urabe M, Honjo H, Tamura T, Noguchi T, et al. Estrogen productivity of endometrium and endometrial cancer tissue: Influence of aromatase on proliferation of endometrial cancer cells. J Steroid Biochem Mol Biol 1993;44:463–8.
8. Naitoh K, Honjo H, Yamamoto T, Urabe M, Ogino Y, Yasumura T, et al. Estrone sulfate and sulfatase activity in human breast cancer and endometrial cancer. J Steroid Biochem 1989;33:1049–54.
9. Watanabe T, Kishino Y, Watanabe A, Sasaki H, Tanaka T, Yokoyama K, et al. Anti-estrone sulfatase monoclonal antibodies available for immunohistochemistry. Acta Obstet Gynaecol Jpn 1999;51(Suppl):S362.
10. Parenti G, Meroni G, Ballabio A. The sulfatase gene family. Curr Opin Genet Dev 1997;7:386–91.
11. McKusick VA, Neufeld EF. The mucopolysaccharide storage diseases. In: Stanbury JB, ed. The metabolic basis of inherited disease. New York: McGraw-Hill, 1983:751–77.
12. Kolodny EH, Moser HW. Sulfatide lipidosis: Metachromatic leukodystrophy. In: Stanbury JB, ed. The metabolic basis of inherited disease. New York: McGraw-Hill, 1983: 881–905.
13. Sapiro LJ. Steroid sulfatase deficiency and X-linked ichthyosis. In: Stanbury, JB, ed. The metabolic basis of inherited disease. New York: McGraw-Hill, 1983:1027–39.
14. Pasqualini JR, Chetrite G, Nestour EL. Control and expression of oestrone sulphatase activities in human breast cancer. J Endocrinol 1996;150(Suppl):S99–S105.
15. Reed MJ, Purohit A, Duncan LJ, Singh A, Roberts CJ, Williams GJ, et al. The role of cytokines and sulphatase inhibitors in regulating oestrogen synthesis in breast tumors. J Steroid Biochem Mol Biol 1995;53:413–20.
16. Li PK, Pillai R, Young BL, Bender WH, Martino DM, Lin FT. Synthesis and biochemical studies of estrone sulfatase inhibitors. Steroids 1993;58:106–11.
17. Li PK, Pillai R, Dibbelt L. Estrone sulfate analogs as estrone sulfatase inhibitors. Steroids 1995;60:299–306.
18. Purohit A, Wang DY, Ghilchik MW, Reed MJ. Regulation of aromatase and sulphatase in breast tumor cells. J Steroid Biochem Mol Biol 1992;41:563–6.
19. Purohit A, Chapman O, Duncan L, Reed MJ. Modulation of oestrone sulphatase activity in breast cancer cell lines by growth factors. J Steroid Biochem Mol Biol 1992;41:563–6.
© 2001 by The American College of Obstetricians and Gynecologists.