![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
3 Department of Biochemistry, Faculty of Medicine, 4 Faculty of Nutrition and Food Sciences, 5 Department of Anatomy, Faculty of Medicine, and 6 Laboratory for Molecular Cell Biology, Faculty of Medicine and Institute for Molecular and Cell Biology, University of Porto, 4200-319 Porto, Portugal
* To whom correspondence should be addressed. E-mail: rosariom{at}med.up.pt.
 |
ABSTRACT |
|---|
 TOP ABSTRACT IntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
|---|
|
Introduction |
|---|
|
TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
|---|
Estrogens can be produced in the AT from aromatization of circulating androgens by the action of aromatase (7) and its expression is higher in scAT in comparison to vAT (8). Evidence from estrogen deficiency models shows that estrogen signaling contributes to weight regulation and its absence is related to v obesity, insulin resistance, and glucose intolerance along with other features of the metabolic syndrome (9,10). In men, aromatase mutations that give rise to lack of enzyme function are related with undetectable estrogen concentration and normal or high testosterone and gonadotropins associated with excess weight or obesity. Body fat accumulation in the v region and alterations of lipid metabolism are present as well (11). In women, aromatase deficiency also results in developmental and metabolic alterations. Surprisingly, these disturbances are ameliorated by estrogen treatment (12). Furthermore, alterations in AT distribution and inflammatory status are common after estrogen production ceases in menopause and may also be reversed by hormone replacement treatment (13). vAT has a relative estrogen insufficiency due to its lower expression of aromatase compared with scAT (7). Although estrogens influence body fat accumulation by altering appetite and energy expenditure, direct AT effects are possible, including lipoprotein lipase inhibition and hormone-sensitive lipase stimulation, as well as preadipocyte proliferation and differentiation alterations (14,15). This hormone is thus likely to induce its antiobesity effects locally, as its tissue concentration may be 
10 times higher than circulating concentrations (16) and because AT accounts for most of the estrogen production in males and postmenopausal females (10).
The investigation of the antiobesity properties of food components is a popular field of research, because it may lead to the discovery of new naturally occurring agents to prevent or treat obesity. There are studies on green tea (GT) and GT components, especially catechins, demonstrating their effects on energy expenditure, fat oxidation, blood lipids, and preadipocyte differentiation and adipocyte apoptosis (17,18). Yet, to date, little is known about its ability to interfere with estrogen synthesis in vivo, particularly in the AT. Aromatase activity inhibition by some GT components has been demonstrated on placental microsomes (19) and choriocarcinoma-derived JAR cells (20), but in vivo long-term studies are lacking. We have also recently found that red wine, a beverage containing compounds able to inhibit aromatase activity in cellular lines, could induce an increase in aromatase expression in animal models after prolonged treatment (21). Taking this information into account, together with the known effects of locally produced estrogens, it is possible that GT components may modulate aromatase in the AT and that, as a consequence, alterations in tissue dynamics may occur. For this reason, we intended to determine the effect of chronic consumption of GT on the expression of aromatase in the AT from different anatomical locations. We also sought to determine the effect of GT intake on adipocyte size, AT apoptosis, and proliferation. To approach this subject, we treated adult male Wistar rats with GT, performed evaluations in scAT and vAT, and measured circulating 17β-estradiol and testosterone concentrations.
|
Materials and Methods |
|---|
|
TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
|---|
-tocopherol equivalents). Bottles containing the beverages were protected from light to avoid oxidation of light-sensitive components and beverages were renewed every 2–3 d with water (controls) or freshly prepared GT. We monitored ad libitum consumption of both food and fluid every other day and recorded rat weight every week. To prepare GT infusion, 1 L of boiling water (100°C) was dispensed over 3 tea bags (1.3 g/bag) for 5 min. The infusion was allowed to cool to room temperature before being transferred to the bottles and supplied to the rats. The tea used during the treatments was from the same batch and the composition of the final beverage was determined after 3 randomly chosen infusion preparations. Tea composition in catechins was determined using HPLC (22) and was as follows: (–)-epigallocatechin-3-gallate (EGCG), 1.21 ± 0.256 mmol/L; (–)-epicatechin (EC), 0.84 ± 0.071 mmol/L; EC-3-gallate, 0.29 ± 0.067 mmol/L; (+)-gallocatechin-3-gallate, 0.07 ± 0.001 mmol/L. Mean caffeine content was 0.52 ± 0.022 mmol/L, measured using GC-ion trap MS, as described (23). Animal procedures were according to the European Community guidelines (86/609/EEC) and the Portuguese Act (129/92) for the use of experimental animals. Tissue collection and preparation. At the end of treatment, fed rats were anesthetized (intraperitoneal sodium pentobarbital; 80 mg/kg body weight) and perfused with saline at 4°C. All rats were killed between 1000 and 1200. Samples from sc (inguinal) and v (mesenteric) AT were removed and frozen in liquid nitrogen for protein extraction. A fraction of each sample was immersed in 10% formalin solution for paraffin embedding and 4-µm-thick tissue sections were made for histological analysis, apoptosis determination, and immunohistochemistry.
Plasma 17β-estradiol and testosterone measurement. Before perfusion, blood was drawn from the left ventricle into heparinized tubes and plasma fractions were frozen at –80°C until analysis. 17β-Estradiol and testosterone were measured with RIA using antiserum against 17β-estradiol and 125I-17β-estradiol (Coat-a-Count TKE2, Diagnostic Products) or antiserum against testosterone and 125I-testosterone (Testo-RIA-CT, Biosource), respectively. The commercial kits used to measure 17β-estradiol and testosterone were specific for these hormones and had very low reactivity with estrone or androstenedione. The method for measuring 17β-estradiol had a sensitivity of 29 pmol/L; the detection limit was 174 nmol/L for the test measuring testosterone.
Measurement of adipocyte size. Hematoxylin-eosin–stained tissue sections were observed and photographed under specimen identity occultation. We estimated adipocyte mean diameter using NIS Elements BR software (Nikon). Fifty to 100 adipocytes from 5 randomly selected different fields were measured for each sample.
Apoptosis assay. Terminal deoxynucleotidyl transferase-mediated deoxyuridine triphosphate nick end-labeling was used according to the manufacturer (Roche) in paraffin-embedded tissue sections. Cell nuclei were counterstained with 4',6-diamidine-2'-phenylindole dihydrochloride (DAPI; Roche). Slides were visualized under a fluorescence microscope (Nikon 50i) at 200x magnification. Apoptosis was determined as the percentage of positive over total counted cells. We counted 200 nuclei from different section fields for each sample.
Determination of proliferation. Proliferation was measured through immunohistochemical labeling of proliferation-characteristic nuclear protein Ki67. This cellular marker is expressed in all active phases of the cell cycle (G1, S, G2, and mitosis) and is absent in quiescent (G0) cells (24). Anti-Ki67 primary and fluorescein isothiocyanate-conjugated secondary antibodies (1:50 and 1:200 in 4% bovine serum albumin, respectively; Santa Cruz Biotechnology) were used. Total nuclei were counterstained with DAPI and visualized at 200x magnification. Nuclei stained with anti-Ki67 antibody were counted in 5 randomly selected section fields. Total DAPI-stained nuclei were counted for the same optical fields. Proliferation is given as a percentage of proliferating cells over total cells. We counted 400–600 nuclei in scAT and 200–300 in vAT from different section fields for each sample.
Western blotting. AT samples (0.8–1 g) were incubated at 4°C for 2 h in 1.5 mL of detergent protein extraction buffer (50 mmol/L Tris-HCl, pH 7.4, 1% Triton X-100, 0.2% sodium deoxycholate, 0.2% sodium dodecylsulfate, 1 mmol/L EDTA, 1 mmol/L phenylmethylsulfonyl fluoride) containing 25 µL protease inhibitor cocktail (Sigma) as described (25). Each sample was incubated at 4°C with agitation for 2 h in the buffer and subsequently homogenized in a Teflon-glass homogenizer and centrifuged at 10,000 x g for 10 min. The infranatant protein solution was collected, quantified by bicinchoninic acid protein assay (Pierce), and stored at –80°C. Before SDS-PAGE separation, proteins were dissolved 1:1 in loading buffer (Bio-Rad Laboratories) containing 2% mercaptoethanol, denatured for 90 s at 95°C, and 20 µg of protein was loaded per well. Anti-aromatase polyclonal antibody (1:200, Santa Cruz Biotechnology) and horseradish peroxidase-conjugated polyclonal antibody (1:1000) were used followed by chemiluminescent detection. β-Actin primary antibody (1:1000, Lab Vision) and secondary antibody hybridizations were conducted using the same procedure. Band intensity was determined using Gel Pro Analyzer (Media Cybernetics) and aromatase was normalized to β-actin expression (expressed in arbitrary units). Determinations were made in membranes containing both control and GT-treated rat proteins in 2 separate experiments.
Statistical analyses. Results are expressed as means ± SEM. Mixed effects model with random intercept was used to assess the progression of body weight with time (26 repeated measures of the body weight) adjusted for the liquid and food intake. This model allows the incorporation of both fixed (population-specific) and random (subject-specific) variables and assumes that the vector of repeated measures on each subject follows a linear regression model. Using the random intercept, it is tested if variability in subject-specific slopes can be ascribed to treatment. The differences between 2 groups were assessed by unpaired Student's t-test or Mann Whitney U-test when variance of measurements was significantly different. We used Fisher's exact test to determine the difference between groups in 17β-estradiol measurements, comparing the number of detectable and undetectable measurements in controls and treated animals. Association between outcome variables was determined with the data pooled from the 2 groups using Pearson or Spearman correlations for data with normal or nonnormal distributions, respectively, and AT measurements were compared within the same tissue. The normality of the distribution for each outcome variable was tested using Shapiro's test. Statistical analyses were performed using Graph Pad Prism software (Graph Pad Software version 3.0) and SPSS (SPSS 12.0 for Windows). Differences were considered significant when P < 0.05.
|
Results |
|---|
|
TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
|---|
|
|
|
Aromatase expression.
Relative aromatase expression in homogenates from scAT was 213% higher than that of vAT (P < 0.05) in control rats (Fig. 2). Interestingly, GT treatment induced a marked increase in aromatase expression in both AT locations to 318.5 ± 60.6% of control in scAT (P < 0.05) and 285.5 ± 82.9% of control in vAT (P < 0.01).
|
|
|
|
|
Discussion |
|---|
|
TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
|---|
59% of the total of catechins), (–)-epigallocatechin, EC-3-gallate, and EC (26). After oral administration of tea, catechins can be found in the plasma and widely distributed in rat tissues (26). From modulation of food intake and energy expenditure, to antiadipogenic and apoptosis-promoting activities, several relevant effects have been described for tea in the past few years; catechins, present in great amounts in GT, have been shown to be involved in most of the actions described (27–29). Caffeine, ranging from 0.51–0.86 mmol/L (26), may also account for some of the effects attributed to GT, as it increases thermogenesis and acts synergistically with other tea components such as the catechins (30). It must be stressed, however, that some studies did not support a GT effect on energetic metabolism (31,32). As shown in the present study, prolonged GT intake by adult male Wistar rats resulted in a significantly lower weight gain compared with controls. In association with lower weight gain, increased aromatase expression was observed in both scAT and vAT of GT-treated rats. Furthermore, this increase in aromatase expression reflected upon higher plasma concentration of 17β-estradiol. We used adult rats to avoid the variations in body weight due to changes in lean mass that are associated with growth in younger rats.
Although decreased body weight gain has already been reported both in humans and animals after GT treatment (27,33,34), increased estrogen production has not yet been related to GT capacity to regulate body composition. Previous studies reported that GT catechins were able to inhibit aromatase activity in the placenta (19,20). The beverage, when placed in contact with the same cells, reduced aromatase activity by 50%. However, those studies investigated acute effects. It seems likely that chronic inhibition of aromatase activity could result in increased aromatase expression as a feedback mechanism (35–37). Furthermore, catechins may also interfere with estrogen signaling through interaction with estrogen receptor (ER) or ERβ (38–40), although the actions reported are not consistent between cell lines, possibly due to the presence of distinct costimulators or corepressors, making it difficult to predict the effects on the AT. Xanthines could also be involved in the effect of GT upon aromatase by increasing cellular cAMP, a known inducer of aromatase expression through the AT aromatase promoter I.3 (7).
Increased aromatase expression reflected upon plasma 17β-estradiol as this hormone, undetectable in controls, reached measurable concentrations in GT-drinking rats. Indeed, there was a significant positive association between plasma 17β-estradiol and aromatase expression in scAT and vAT. In parallel, testosterone concentration significantly decreased in the same rats. Plasma concentrations of estradiol and testosterone were negatively associated. This decrease in circulating testosterone may be at least in part due to its aromatization to 17β-estradiol or estrone (7). It has been proposed that the relative amount of estrogens and androgens or of their receptors (10,14) will determine the response of the AT to sex steroids. Furthermore, those ratios are thought to be closely related to the sexual dimorphism in body fat distribution and to the differential risk of men and pre- and postmenopausal women to develop metabolic diseases (41).
Estrogens have profound effects on energy metabolism as demonstrated by the amount of fat accumulation in estrogen insufficiency models (9,10), with reversion of the effect by estrogen treatment. On the other hand, estrogen therapy ameliorates the plasma lipid profile in healthy males, although no effect has been reported on body weight (42). Mice lacking ER develop the same metabolic abnormalities as aromatase knockout mice, showing that ER mediates the protective actions of estrogens (14). These alterations were absent in ERβ knockouts (14), although recently it has been reported that mice lacking ERβ, when fed a high-fat diet, had improved insulin sensitivity and glucose tolerance compared with wild-type mice (43).
As first described by Vague (44), the 2 main AT depots display major differences in the metabolic consequences they induce. Unfavorable cardiovascular profiles and the risk of developing type 2 diabetes are greater in individuals whose AT is located within the abdominal cavity than in those having mainly sc fat (2), although there is still no exact knowledge of the reasons for these differences. Sex steroids contribute to determine the location of fat distribution and the ability of different AT depots to produce and respond to these hormones varies tremendously (45). Confirming previous findings (8), we found the expression of aromatase in scAT was higher than that in vAT. The differential aromatase expression may contribute to the metabolic differences encountered between the accumulation of fat in the 2 locations. In fact, 2 key proteins involved in lipid deposition, lipoprotein lipase and leptin, are transcriptionally regulated by 17β-estradiol, being down- and upregulated, respectively, by this hormone (46,47). Interestingly, with GT intake, vAT expression of aromatase rose to levels comparable to those found in scAT from controls.
scAT is the main AT depot responsible for estrogen production. Because plasma 17β-estradiol was undetectable in control rats, vAT would be under little influence of this hormone due to its low endogenous capacity to produce estrogens. On the other hand, in GT-treated rats, both scAT and vAT may be influenced by estrogens and the increase in plasma concentration of 17β-estradiol may also indicate that distant tissues or organs may be affected. Food intake was not influenced, as rats from both groups ingested similar amounts of food. In fact, in models of estrogen insufficiency, increases in body fat mass are not associated with hyperphagia (48). Estrogens have, however, been shown to increase voluntary energy expenditure (48) and leptin and decrease insulin sensitivities in the hypothalamus (49).
Either induced by the elevation of 17β-estradiol production or directly by the presence of GT components, e.g. catechins or xanthines, other interesting effects were found on the AT. First, measurement of adipocyte diameter revealed a decrease in adipocyte size in both AT depots following GT treatment. It has been pointed out that obesity and its metabolic consequences result from both increased number of adipose cells (hyperplasia) and adipocyte size (hypertrophy) (50). However, adipocyte size seems more critical for predicting morbidities related to AT excess. Inflammatory events originating in AT play a key role in obesity-associated diseases, the infiltration of macrophages in the AT being a central event (4). Cinty et al. (51) have shown that macrophages in the AT surround dead adipocytes. We have previously demonstrated that, for simple physical reasons, larger adipocytes are more prone to rupture than small ones (52), particularly when they are contained in the abdominal cavity where sudden variations of pressure frequently take place (53,54). The inflammatory events related with large adipocytes are further supported by the recent report that adiponectin release from adipocytes is inversely correlated with adipocyte size in obese patients (50). Conversely, tumor necrosis factor-, interleukin-6, and high sensitivity C-reactive protein circulating concentrations were positively associated with the same variable (50). Thus, by reducing adipocyte size, chronic GT treatment might have a great impact on obesity-driven inflammation. Both estrogens and GT catechins have properties that can account for reduced adipocyte size, some of which are similar. One example is the stimulation of catecholamine-induced lipolysis toward increased β-oxidation of fatty acids (33,55). It was very interesting to find that plasma 17β-estradiol concentrations were inversely correlated with adipocyte size in scAT and vAT. Adipocyte size was also positively correlated to body weight.
Together with smaller adipocyte size, we found increased apoptosis in vAT and increased proliferation in both AT depots, suggesting that AT is undergoing remodeling. Epidemiological studies have shown that usual GT drinkers have lower waist circumferences (34), which is in agreement with the observation of a high apoptotic rate found in the vAT of GT-treated rats. Modification of the AT profile through selective apoptosis of mature adipocytes in the vAT would avoid the rupture of large lipid-laden adipocytes and the generation of an inflammatory response. Furthermore, the presence of higher apoptosis in vAT was positively associated with 17β-estradiol and inversely correlated with testosterone plasma concentrations, in conformity with the sexual dimorphism of vAT accumulation.
AT remodeling extended also to cell proliferation. Preadipocytes and mature adipocytes are the major cell types that compose AT, but vascular endothelial and smooth muscle cells are also documented (14). Proliferation was identified mainly in the AT stromal fraction where preadipocytes and blood vessels are predominant. Although we do not know which cell types were Ki67 immunostained, proliferation of either type would reveal greater capacity for lipid trafficking between mature and maturing adipocytes and between locations of AT deposition. Both aromatase expression and plasma 17β-estradiol concentrations had a strong positive correlation with the percentage of proliferating cells in scAT and vAT. Thus, it is very likely that 17β-estradiol production in the AT may be responsible for the changes observed in proliferation either in cells from the adipose lineage or others. For example, endothelial, smooth muscle, and mesenchymal cells may be induced to proliferate in the presence of 17β-estradiol (56). Previous studies reported that GT catechins, particularly EGCG, have antimitogenic effects on adipose cells (29). This apparent contradiction may have arisen from the fact that the antimitogenic action was reported in vitro in 3T3-L1 preadipocytes. In that experimental setting, the effect of GT or GT catechins may not be mediated through estrogens, because these cells either lack or have residual aromatase expression. Furthermore, as mentioned previously, several other types of cells compose AT and many of them are known targets for estrogens actions. Also worth noting is the inverse correlation between plasma testosterone and the percentage of proliferating cells. This, along with the known antiadipogenic effect of testosterone (41), implicates this hormone in the reduction of AT plasticity in response to energy excess that predisposes men to metabolic disorders.
The fact that total AT or body composition was not measured is an important limitation of this study. In addition, determination of metabolic parameters and baseline and time course measurements of hormones would strengthen the conclusions. Even so, relevant alterations were observed after GT treatment, which deserve attention and further study.
In conclusion, GT intake induced profound remodeling of AT. The causal role of the GT-induced increase in aromatase expression and circulating concentration of 17β-estradiol needs to be explored, although the strong correlations of 17β-estradiol and aromatase measurements with adipocyte size, apoptosis, and proliferation are in favor of the existence of a causal relationship. We also propose that the presence of a higher precursor adipose cell pool together with the proadipogenic effect of estrogens and attenuation of the antiadipogenic effect of testosterone would facilitate differentiation and the distribution of fat in an increased number of cells, resulting in reduced cell size. This could then contribute to the redistribution of body AT and decrease obesity-related inflammation.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
2 Author disclosures: R. Monteiro, M. Assunção, J. P. Andrade, D. Neves, C. Calhau, and I. Azevedo, no conflicts of interest.
7 Abbreviations used: AT, adipose tissue; DAPI, 4',6-diamidine-2'-phenylindole dihydrochloride; EC, (–)-epicatechin; EGCG, (–)-epigallocatechin-3-gallate; ER, estrogen receptor; GT, green tea; sc, subcutaneous; v, visceral.
Manuscript received 7 May 2008. Initial review completed 28 May 2008. Revision accepted 7 August 2008.
|
LITERATURE CITED |
|---|
|
TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
|---|
1. Hotamisligil GS. Inflammation and metabolic disorders. Nature. 2006;444:860–7.[Medline]
2. Eckel RH, Grundy SM, Zimmet PZ. The metabolic syndrome. Lancet. 2005;365:1415–28.[Medline]
3. Lafontan M, Berlan M. Do regional differences in adipocyte biology provide new pathophysiological insights? Trends Pharmacol Sci. 2003;24:276–83.[Medline]
4. Xu H, Barnes GT, Yang Q, Tan G, Yang D, Chou CJ, Sole J, Nichols A, Ross JS, et al. Chronic inflammation in fat plays a crucial role in the development of obesity-related insulin resistance. J Clin Invest. 2003;112:1821–30.[Medline]
5. Kershaw EE, Flier JS. Adipose tissue as an endocrine organ. J Clin Endocrinol Metab. 2004;89:2548–56.
6. Linder K, Arner P, Flores-Morales A, Tollet-Egnell P, Norstedt G. Differentially expressed genes in visceral or subcutaneous adipose tissue of obese men and women. J Lipid Res. 2004;45:148–54.
7. Simpson ER, Clyne C, Rubin G, Boon WC, Robertson K, Britt K, Speed C, Jones M. Aromatase: a brief overview. Annu Rev Physiol. 2002;64:93–127.[Medline]
8. Rink JD, Simpson ER, Barnard JJ, Bulun SE. Cellular characterization of adipose tissue from various body sites of women. J Clin Endocrinol Metab. 1996;81:2443–7.[Abstract]
9. Heine PA, Taylor JA, Iwamoto GA, Lubahn DB, Cooke PS. Increased adipose tissue in male and female estrogen receptor-alpha knockout mice. Proc Natl Acad Sci USA. 2000;97:12729–34.
10. Simpson ER, Misso M, Hewitt KN, Hill RA, Boon WC, Jones ME, Kovacic A, Zhou J, Clyne CD. Estrogen - the good, the bad, and the unexpected. Endocr Rev. 2005;26:322–30.
11. Jones ME, Boon WC, Proietto J, Simpson ER. Of mice and men: the evolving phenotype of aromatase deficiency. Trends Endocrinol Metab. 2006;17:55–64.[Medline]
12. Jones ME, McInnes KJ, Boon WC, Simpson ER. Estrogen and adiposity: utilizing models of aromatase deficiency to explore the relationship. J Steroid Biochem Mol Biol. 2007;106:3–7.[Medline]
13. Sites CK, L'Hommedieu GD, Toth MJ, Brochu M, Cooper BC, Fairhurst PA. The effect of hormone replacement therapy on body composition, body fat distribution, and insulin sensitivity in menopausal women: a randomized, double-blind, placebo-controlled trial. J Clin Endocrinol Metab. 2005;90:2701–7.
14. Cooke PS, Naaz A. Role of estrogens in adipocyte development and function. Exp Biol Med (Maywood). 2004;229:1127–35.
15. Mayes JS, Watson GH. Direct effects of sex steroid hormones on adipose tissues and obesity. Obes Rev. 2004;5:197–216.[Medline]
16. Yager JD, Davidson NE. Estrogen carcinogenesis in breast cancer. N Engl J Med. 2006;354:270–82.
17. Moon HS, Lee HG, Choi YJ, Kim TG, Cho CS. Proposed mechanisms of (–)-epigallocatechin-3-gallate for anti-obesity. Chem Biol Interact. 2007;167:85–98.[Medline]
18. Wolfram S, Wang Y, Thielecke F. Anti-obesity effects of green tea: from bedside to bench. Mol Nutr Food Res. 2006;50:176–87.[Medline]
19. Satoh K, Sakamoto Y, Ogata A, Nagai F, Mikuriya H, Numazawa M, Yamada K, Aoki N. Inhibition of aromatase activity by green tea extract catechins and their endocrinological effects of oral administration in rats. Food Chem Toxicol. 2002;40:925–33.[Medline]
20. Monteiro R, Azevedo I, Calhau C. Modulation of aromatase activity by diet polyphenolic compounds. J Agric Food Chem. 2006;54:3535–40.[Medline]
21. Monteiro R, Faria A, Mateus N, Calhau C, Azevedo I. Red wine interferes with oestrogen signalling in rat hippocampus. J Steroid Biochem Mol Biol. 2008;111:74–9.[Medline]
22. de Freitas VAP, Glories Y. Concentration and compositional changes of procyanidins in grape seeds and skin of white Vitis vinifera varieties. J Sci Food Agric. 1999;79:1601–6.
23. Zou J, Li N. Simple and environmental friendly procedure for the gas chromatographic-mass spectrometric determination of caffeine in beverages. J Chromatogr A. 2006;1136:106–10.[Medline]
24. Scholzen T, Gerdes J. The Ki-67 protein: from the known and the unknown. J Cell Physiol. 2000;182:311–22.[Medline]
25. Higami Y, Barger JL, Page GP, Allison DB, Smith SR, Prolla TA, Weindruch R. Energy restriction lowers the expression of genes linked to inflammation, the cytoskeleton, the extracellular matrix, and angiogenesis in mouse adipose tissue. J Nutr. 2006;136:343–52.
26. Cabrera C, Artacho R, Gimenez R. Beneficial effects of green tea: a review. J Am Coll Nutr. 2006;25:79–99.
27. Kao YH, Hiipakka RA, Liao S. Modulation of endocrine systems and food intake by green tea epigallocatechin gallate. Endocrinology. 2000;141:980–7.
28. Lin J, Della-Fera MA, Baile CA. Green tea polyphenol epigallocatechin gallate inhibits adipogenesis and induces apoptosis in 3T3–L1 adipocytes. Obes Res. 2005;13:982–90.[Medline]
29. Hung PF, Wu BT, Chen HC, Chen YH, Chen CL, Wu MH, Liu HC, Lee MJ, Kao YH. Antimitogenic effect of green tea (-)-epigallocatechin gallate on 3T3–L1 preadipocytes depends on the ERK and Cdk2 pathways. Am J Physiol Cell Physiol. 2005;288:C1094–108.
30. Dulloo AG, Seydoux J, Girardier L, Chantre P, Vandermander J. Green tea and thermogenesis: interactions between catechin-polyphenols, caffeine and sympathetic activity. Int J Obes Relat Metab Disord. 2000;24:252–8.[Medline]
31. Diepvens K, Kovacs EM, Nijs IM, Vogels N, Westerterp-Plantenga MS. Effect of green tea on resting energy expenditure and substrate oxidation during weight loss in overweight females. Br J Nutr. 2005;94:1026–34.[Medline]
32. Hsu CH, Tsai TH, Kao YH, Hwang KC, Tseng TY, Chou P. Effect of green tea extract on obese women: a randomized, double-blind, placebo-controlled clinical trial. Clin Nutr. 2008;27:363–70.[Medline]
33. Dulloo AG, Duret C, Rohrer D, Girardier L, Mensi N, Fathi M, Chantre P, Vandermander J. Efficacy of a green tea extract rich in catechin polyphenols and caffeine in increasing 24-h energy expenditure and fat oxidation in humans. Am J Clin Nutr. 1999;70:1040–5.
34. Wu CH, Lu FH, Chang CS, Chang TC, Wang RH, Chang CJ. Relationship among habitual tea consumption, percent body fat, and body fat distribution. Obes Res. 2003;11:1088–95.[Medline]
35. Harada N, Hatano O. Inhibitors of aromatase prevent degradation of the enzyme in cultured human tumour cells. Br J Cancer. 1998;77:567–72.[Medline]
36. Kao YC, Okubo T, Sun XZ, Chen S. Induction of aromatase expression by aminoglutethimide, an aromatase inhibitor that is used to treat breast cancer in postmenopausal women. Anticancer Res. 1999;19:2049–56.[Medline]
37. Yue W, Brodie AM. Mechanisms of the actions of aromatase inhibitors 4-hydroxyandrostenedione, fadrozole, and aminoglutethimide on aromatase in JEG-3 cell culture. J Steroid Biochem Mol Biol. 1997;63:317–28.[Medline]
38. Belguise K, Guo S, Sonenshein GE. Activation of FOXO3a by the green tea polyphenol epigallocatechin-3-gallate induces estrogen receptor alpha expression reversing invasive phenotype of breast cancer cells. Cancer Res. 2007;67:5763–70.
39. Farabegoli F, Barbi C, Lambertini E, Piva R. (-)-Epigallocatechin-3-gallate downregulates estrogen receptor alpha function in MCF-7 breast carcinoma cells. Cancer Detect Prev. 2007;31:499–504.[Medline]
40. Kuruto-Niwa R, Inoue S, Ogawa S, Muramatsu M, Nozawa R. Effects of tea catechins on the ERE-regulated estrogenic activity. J Agric Food Chem. 2000;48:6355–61.[Medline]
41. Blouin K, Boivin A, Tchernof A. Androgens and body fat distribution. J Steroid Biochem Mol Biol. 2008;108:272–80.[Medline]
42. Sayed Y, Taxel P. The use of estrogen therapy in men. Curr Opin Pharmacol. 2003;3:650–4.[Medline]
43. Foryst-Ludwig A, Clemenz M, Hohmann S, Hartge M, Sprang C, Frost N, Krikov M, Bhanot S, Barros R, et al. Metabolic actions of estrogen receptor beta (ERbeta) are mediated by a negative cross-talk with PPARgamma. PLoS Genet. 2008;4:e1000108.[Medline]
44. Vague J. Importance of the measurement of fat distribution in pathology. Bull Mem Soc Med Hop Paris. 1950;66:1572–4.[Medline]
45. Belanger C, Luu-The V, Dupont P, Tchernof A. Adipose tissue intracrinology: potential importance of local androgen/estrogen metabolism in the regulation of adiposity. Horm Metab Res. 2002;34:737–45.[Medline]
46. Homma H, Kurachi H, Nishio Y, Takeda T, Yamamoto T, Adachi K, Morishige K, Ohmichi M, Matsuzawa Y, et al. Estrogen suppresses transcription of lipoprotein lipase gene. Existence of a unique estrogen response element on the lipoprotein lipase promoter. J Biol Chem. 2000;275:11404–11.
47. Machinal-Quelin F, Dieudonne MN, Pecquery R, Leneveu MC, Giudicelli Y. Direct in vitro effects of androgens and estrogens on ob gene expression and leptin secretion in human adipose tissue. Endocrine. 2002;18:179–84.[Medline]
48. Jones ME, Thorburn AW, Britt KL, Hewitt KN, Wreford NG, Proietto J, Oz OK, Leury BJ, Robertson KM, et al. Aromatase-deficient (ArKO) mice have a phenotype of increased adiposity. Proc Natl Acad Sci USA. 2000;97:12735–40.
49. Clegg DJ, Brown LM, Woods SC, Benoit SC. Gonadal hormones determine sensitivity to central leptin and insulin. Diabetes. 2006;55:978–87.
50. Bahceci M, Gokalp D, Bahceci S, Tuzcu A, Atmaca S, Arikan S. The correlation between adiposity and adiponectin, tumor necrosis factor alpha, interleukin-6 and high sensitivity C-reactive protein levels. Is adipocyte size associated with inflammation in adults? J Endocrinol Invest. 2007;30:210–4.[Medline]
51. Cinti S, Mitchell G, Barbatelli G, Murano I, Ceresi E, Faloia E, Wang S, Fortier M, Greenberg AS, et al. Adipocyte death defines macrophage localization and function in adipose tissue of obese mice and humans. J Lipid Res. 2005;46:2347–55.
52. Monteiro R, de Castro PM, Calhau C, Azevedo I. Adipocyte size and liability to cell death. Obes Surg. 2006;16:804–6.[Medline]
53. Cobb WS, Burns JM, Kercher KW, Matthews BD, James Norton H, Todd Heniford B. Normal intraabdominal pressure in healthy adults. J Surg Res. 2005;129:231–5.[Medline]
54. Lambert DM, Marceau S, Forse RA. Intra-abdominal pressure in the morbidly obese. Obes Surg. 2005;15:1225–32.[Medline]
55. D'Eon TM, Souza SC, Aronovitz M, Obin MS, Fried SK, Greenberg AS. Estrogen regulation of adiposity and fuel partitioning. Evidence of genomic and non-genomic regulation of lipogenic and oxidative pathways. J Biol Chem. 2005;280:35983–91.
56. Zhao X, Huang L, Yin Y, Fang Y, Zhao J, Chen J. Estrogen induces endothelial progenitor cells proliferation and migration by estrogen receptors and PI3K-dependent pathways. Microvasc Res. 2008;75:45–52.[Medline]
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
