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Department of Medicine (K.K., E.E., A.H., H.Y.-J.), Atherosclerosis Research Unit, King Gustaf V Research Institute, Karolinska Institutet, S-17176 Stockholm, Sweden; Finnish Twin Cohort Study (K.H.P., J.K.), Department of Public Health, University of Helsinki, 00014 Helsinki, Finland; Obesity Research Unit (K.H.P., A.R.), Department of Psychiatry, Helsinki University Hospital, 00029 Helsinki, Finland; Department of Medicine (K.H.P., H.Y.-J.), Division of Diabetes, Helsinki University Central Hospital, 00029 Helsinki, Finland; and National Public Health Institute (J.K.), Department of Mental Health and Alcohol Research, 00300 Helsinki, Finland
Address all correspondence and requests for reprints to: Katja Kannisto, Department of Medicine, Atherosclerosis Research Unit, King Gustaf V Research Institute, Karolinska Institutet, S-17176 Stockholm, Sweden. E-mail: katja.kannisto{at}medks.ki.se.
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Abstract |
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overexpression, central obesity, and insulin resistance. It is controversial whether 11ß-HSD-1 or GR expression are increased in human adipose tissue in obesity. We studied effects of acquired obesity on 11ß-HSD-1 gene (real-time PCR) and protein (Western blotting) expression in sc adipose tissue in 17 monozygotic twin pairs aged 24–27 yr with a mean intrapair difference in body mass index (BMI) of 3.8 kg/m2 (range 0.4–10.1 kg/m2). Intrapair correlations were calculated to study effects of acquired obesity on 11ß-HSD-1 expression. Western blot analysis of adipose tissue homogenates identified approximately 50- and approximately 68-kDa proteins specific for 11ß-HSD-1. Both structural forms correlated positively with 11ß-HSD-1 mRNA concentrations. Intrapair differences in 11ß-HSD-1 mRNA, and the 50- and 68-kDa proteins in sc adipose tissue correlated positively with those in BMI (kilograms per square meter) (r = 0.78 for 11ß-HSD-1 mRNA, P = 0.0002; r = 0.87 for the 11ß-HSD-1 50-kDa protein, P = 0.0003; and r = 0.62 for the 11ß-HSD-1 68-kDa protein, P = 0.033), total body fat (percent) (r = 0.65, P = 0.005; r = 0.83, P = 0.001; and r = 0.69, P = 0.013, respectively) and sc fat (cubed centimeters) (r = 0.66, P = 0.004; r = 0.94, P = 0.0001; and r = 0.71, P = 0.009, respectively). Furthermore, 11ß-HSD-1 mRNA and 50-kDa protein expression, but not 68-kDa protein expression, correlated positively with intrapair differences in intraabdominal fat mass (cubed centimeters) (r = 0.62, P = 0.008; r = 0.69, P = 0.013; r = 0.48, P = 0.112) and serum fasting insulin concentration (milliunits per liter) (r = 0.76, P = 0.0004; r = 0.60, P = 0.037; and r = 0.43, P = 0.160, respectively). Intrapair differences in GR expression were significantly inversely correlated with those in BMI and total and sc fat mass. In conclusion, expression of 11ß-HSD-1 in sc adipose tissue is increased in human acquired obesity and is closely related to accumulation of sc and intraabdominal fat and features of insulin resistance.
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Introduction |
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Obesity appears to be associated with increased 11ß-HSD-1 activity in adipose tissue in both rodents and humans (4, 5, 6, 7, 8, 9), but the available data are controversial. Tomlinson et al. (10) recently found no correlation of 11ß-HSD-1 mRNA (by real-time PCR) in human sc or omental adipocytes with obesity. In contrast, Wake et al. (11) showed human adipose tissue 11ß-HSD-1 mRNA levels to be closely correlated with 11ß-HSD-1 enzyme activity, and both gene expression and enzyme activity were positively correlated with indices of obesity and insulin resistance (11). 11ß-HSD-1 activity is also increased in adipose tissue of obese Zucker rats (5). Selective 11ß-HSD-1 inhibitors lower blood glucose levels and improve insulin sensitivity in mouse models of type 2 diabetes (12). Transgenic overexpression of 11ß-HSD-1 under the adipocyte fatty acid binding protein (aP2) promoter enhancer in mice white adipose tissue results in central obesity, dyslipidemia, and insulin resistance (13), whereas homozygous 11ß-HSD-1 knockout mice are protected from obesity and the development of metabolic syndrome (14). These mice also overexpress GR (glucocorticoid receptor) in visceral adipose tissue, which could further enhance glucocorticoid action.
In general, cross-sectional studies of human disease markers rarely permit unequivocal distinction between genetic vs. environmental and lifestyle effects on the investigated variables. However, studies of monozygotic (MZ) twins discordant for markers of disease can help to distinguish between genetic and acquired causes (15). Studying young adults within a narrow age range minimizes early effects of age-related differences on insulin resistance and obesity. In the present study, 17 young adult MZ twin pairs were specifically recruited to exhibit intrapair differences in body mass index (BMI) after screening of 658 MZ twin pairs in the FinnTwin16 cohort for the purpose of investigating whether acquired obesity, independent of genetic factors, is associated with increased 11ß-HSD-1 activity and GR expression in adipose tissue.
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Subjects and Methods |
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The study participants were recruited from FinnTwin16 (16), which is a population-based, longitudinal study including Finnish twins born 1975–1979, identified in the Finnish national population registry. Questionnaires were initially sent to 16-yr-old twins, resulting in 5661 responses (response rates 88 and 93% for boys and girls, respectively). All responding twins were surveyed again at 17 and 18.5 yr of age and as young adults. All 658 MZ twin pairs with information on BMI were screened, yielding only 14 healthy pairs with a BMI difference of at least 4 kg/m2, such that one cotwin had a normal weight (BMI 
25 kg/m2), whereas the other was obese (BMI 30 kg/m2). In the whole sample, intrapair correlation (twin A vs. twin B) in BMI among MZ pairs was 0.78 (n = 658 MZ pairs) (17). Of these extremely discordant MZ pairs, nine pairs (five male and four female pairs) participated in the clinical studies. In addition to these discordant pairs, we studied eight more concordant MZ pairs (four male and four female pairs). These pairs had a BMI difference of less than 2.3 kg/m2. Thus, a total of 17 monozygotic twin pairs (nine male and eight female pairs) having intrapair differences in BMI ranging from 0.4 to 10.1 kg/ m2 were recruited for the present study. Monozygosity was confirmed by genotyping of multiple, informative genetic markers D3S1358 (10 alleles), vWA (12 alleles), FGA (20 alleles), AMEL (two alleles), THO1 (seven alleles), TPOX (eight alleles), CSF1PO (10 alleles), D5S818 (11 alleles), and D13S317 (11 alleles) in the Paternity Testing Laboratory at the National Public Health Institute, Helsinki, Finland. mRNA measurements were performed on all subjects, but due to insufficient adipose tissue material or difficulties in the analytic procedures, protein analyses were available on only 12 pairs. All participants gave their informed consent to participate in the study and were considered healthy (except for obesity) and without medication and were not pregnant. The study was approved by Finnish and Swedish local ethics committees.
Clinical examination
All subjects were studied after an overnight fast starting at 0800 h. Two 18-gauge catheters (Venflon; Viggo-Spectramed, Helsingborg, Sweden) were inserted in the nondominant arm, one in the antecubital vein for infusion of insulin and glucose and another in a retrograde position in a heated dorsal hand vein for withdrawal of arterialized venous blood. After obtaining baseline blood samples for measurement of plasma glucose, serum insulin, lipids, and free fatty acid (FFA) concentrations, a needle aspiration biopsy of sc fat was taken under local anesthesia (18). The fat sample was immediately frozen and stored in liquid nitrogen until analysis. Part of the biopsy was immediately treated with collagenase for 30 min at 37 C. From this sample, the diameter of 200 adipocytes was determined using a microscope. Whole-body insulin sensitivity was thereafter measured using the euglycemic hyperinsulinemic clamp technique. Within the next 24 h, body composition was measured using magnetic resonance imaging (MRI) and dual-energy x-ray absorptiometry (vide infra).
Measures of insulin sensitivity
Whole-body insulin sensitivity of glucose metabolism (M-value) was determined by using the euglycemic hyperinsulinemic clamp technique (19). Insulin (Actrapid Human, Novo Nordisk, Copenhagen, Denmark) was infused in a primed continuous manner at a rate of 1 mU/kg–1·min–1 for 120 min. Normoglycemia was maintained by adjusting the rate of a 20% glucose infusion based on measurements of plasma glucose, which were performed every 5 min from arterialized venous blood. During the insulin infusion, blood samples were taken at 30-min intervals for measurement of serum insulin and FFA concentrations.
Measures of body composition
Determination of sc and intraabdominal fat was performed by series of T1-weighted transaxial MRI scans from region extending from 8 cm above to 8 cm below the fourth and fifth lumbar interspace (16 slices, field of view 375 x 500 mm2, slice thickness 10 mm, breath-hold repetition time 138.9 msec, echo time 4.1 msec). Intraabdominal and sc fat areas were measured using image analysis software (Alice 3.0, Parexel, Waltham, MA) as described (20). The reproducibility of repeated measurements of sc and intraabdominal fat as determined on two separate occasions in our laboratory in nondiabetic subjects is 3 and 5% (n = 10), respectively (21). Body fat was measured by dual-energy x-ray absorptiometry as described (22).
Tissue preparation and real-time quantitative PCR
Frozen fat tissue (50–150 mg) was homogenized in 2 ml RNA STAT-60 (Tel-Test, Friendswood, TX), and total RNA was isolated according to the manufacturer’s instructions. After DNase treatment (RNase-free DNase set, Qiagen, Hilden, Germany), RNA was purified using the RNeasy minikit (Qiagen). RNA concentrations were measured using the RiboGreen fluorescent nucleic acid stain (RNA quantification kit, Molecular Probes, Eugene, OR). The quality of RNA was checked by agarose gel electrophoresis. Average yields of total RNA were 3 ± 1 µg per 100 mg of adipose tissue wet weight and did not correlate with measures of body fat. Isolated RNA was stored at –80 C until quantification of the target mRNAs. A total of 0.1 µg RNA was transcribed into cDNA using Moloney murine leukemia virus reverse transcriptase (Life Technologies, Paisley, UK) and oligo (dT)12–18 primer. Primers and probes for 11ß-HSD-1 and GR used for real-time PCR have been described (11). The primer cDNA specificity was verified in real-time PCR by using both the RNA and cDNA as a template. Three microliters cDNA was amplified with 1x Taqman buffer, 5 mM MgCl2, 200 µM of each dNTP, 200 µM of each primer, 1.25 pM of probe, 0.25 U Amp-Erase Uracil N-Glycosylase, and 1.25 U AmpliTaq Gold (PE Applied Biosystems, Foster City, CA) in real-time quantitative PCR using an ABI PRISM 7000 Sequence detection system instrument and software (PE Applied Biosystems). Expression levels were quantified (in arbitrary units) by generating a six-point serial standard curve. For normalization of gene expression to RNA loading, control samples were run using TATA-box binding protein (Taqman Assays-on-Demand Gene Expression products, PE Applied Biosystems), the expression of which was not related to measures of body fat.
Protein extraction and Western blot analysis
Frozen adipose tissue samples and human primary fibroblasts (150–400 mg) were homogenized in 400 µl ice-cold lysis buffer containing 50 mM Tris (pH 7.5), 1 mM EDTA, 1 mM EGTA, 150 mM NaCl, 5 mM Na4P2O7, 50 mM NaF, 1 mM Na3VO4, 0.1% Triton X-100, 10 mM ß-glycerolphosphate, 500 µM phenylmethylsulfonyl fluoride, 400 µM dithiothreitol, 1 µM Microcystin, and one Complete Mini, EDTA-free tablet (Roche Diagnostics, Mannheim, Germany). Insoluble material was removed by centrifugation (12,000 x g for 10 min at 4 C). Protein was quantitated in the supernatant using the Bradford reagent (Bio-Rad Laboratories, Hercules, CA) and measured spectrophotometrically at 590 nm.
To determine expression of 11ß-HSD-1 and the housekeeping gene, transcription factor IIB (TFIIB) in human sc adipose tissue, 100 and 50 µg of protein, respectively, were loaded on a SDS-polyacrylamide gel and subjected to electrophoresis, which was performed under reducing conditions on 10% polyacrylamide as described by Laemmli (23). The resolved proteins were transferred to a nitrocellulose sheet as detailed by Towbin et al. (24). The nitrocellulose membrane was then incubated with rabbit polyclonal antibodies against 11ß-HSD-1 ( Diagnostic International, San Antonio, TX) and TFIIB (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) at dilutions of 1:5000 and 1:500, respectively. Western blot analysis was also performed using the primary antibody preabsorbed with a 1:500 dilution of the immunizing peptide to demonstrate specificity for 11ß-HSD-1. The blots incubated with the primary antibody were probed with the corresponding secondary antibodies to IgG
(Dakopatts, 
lvsj
, Sweden, 1:50 000 dilution)
conjugated to horseradish peroxidase. The enhanced chemiluminescence Advance Western blot detection system (Amersham Biosciences, Little Chalfont, Buckinghamshire, UK) was used according to the manufacturer’s instructions. The light emitted from the membrane was quantified using the FUJI LAS 1000 (Fuji Photo Film Co., Ltd., Tokyo, Japan). The resulting bands were confirmed by comparing the size of the protein with known molecular markers (Bio-Rad Laboratories). A protein standard was run on each gel to enable comparison of light intensities among different membranes. For normalization of gene expression to protein loading, control samples were run using TFIIB, which is one of the eukaryotic, basal transcription factors that are needed for transcription from protein-coding genes requiring RNA polymerase II, a process that is not sequence specific. TFIIB expression was not related to measures of body fat.
Other measurements
Serum-free insulin concentrations were determined with RIA (Phadeseph Insulin RIA, Pharmacia & Upjohn Diagnostics, Uppsala, Sweden) after precipitation with polyethylene glycol (25). Plasma glucose concentrations were measured in duplicate with the glucose oxidase method using a Glucose analyzer II (Beckman Instruments, Fullerton, CA) (26). Serum total and high-density lipoprotein (HDL) cholesterol, and triglyceride concentrations were measured with enzymatic kits from Roche Diagnostics using an autoanalyzer (Roche Diagnostics Hitachi 917, Hitachi Ltd., Tokyo, Japan). Serum FFAs were measured by a fluorometric assay (27).
Statistical analyses
In the data analysis, most comparisons were based on contrasting the cotwin with the higher BMI (heavier twin) with the cotwin with a smaller BMI (leaner twin) by calculating the correlations between intrapair differences in BMI and other body composition measures with intrapair differences in the other study variables. Paired nonparametric Wilcoxon tests were used to test the significance of intrapair differences for individual variables, whereas Spearman rank-correlations were used to test associations between variables. Mann-Whitney U test was performed to test whether the expression patterns were the same for both genders. Stata software (release 8.0; Stata Corp., College Station, TX) was used for carrying out calculations. Data are shown as mean ± SE. P < 0.05 were considered statistically significant.
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Results |
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Physical and biochemical characteristics including ranges of intrapair differences of the study subjects are shown in Table 1
. By definition, the heavier twins had higher BMIs, and, expectedly, the heavier twins had significantly higher total body fat and total volume of sc and abdominal fat as measured by MRI. The heavier twins were also more insulin resistant than the leaner twins and had lower HDL-cholesterol concentrations.
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In primary human fibroblasts, bands of approximately 34 and 68 kDa were observed in Western blot analysis, consistent with previous data (3). The 34-kDa band was present but very weak in Western blots of human sc adipose tissue biopsies, whereas the 68-kDa band and an additional band of 50 kDa were strongly expressed (Fig. 1). Significantly weaker bands of 34, 50, and 68 kDa were seen when Western blots were carried out after preabsorbing the primary antibodies with the immunizing peptide (dilution 1:500).
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The correlations between intrapair differences in body composition, insulin sensitivity, and 11ß-HSD-1 expression are shown in Table 2. Intrapair differences in 11ß-HSD-1 mRNA and 50- and 68-kDa protein expression were all highly significantly positively correlated with intrapair differences in BMI (Fig. 2 and Table 2). Consequently, heavier twins had higher 11ß-HSD-1 mRNA and protein expression than the leaner twins. Intrapair differences in mRNA and 50- and 68-kDa protein expression of 11ß-HSD-1 all correlated significantly with intrapair differences in total body fat (Table 2). Similarly, intrapair differences in sc 11ß-HSD-1 mRNA and 50- and 68-kDa protein expression correlated with intrapair differences in sc adipose tissue mass (Table 2). Furthermore, intrapair differences in sc 11ß-HSD-1 mRNA and 50-kDa protein expression, but not 68-kDa protein expression, correlated significantly and positively with intrapair differences in intraabdominal fat mass (Table 2).
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). Intrapair differences in serum insulin levels correlated significantly and positively with those of 11ß-HSD-1 mRNA and 50 kDa, but not significantly with intrapair differences in 68-kDa protein expression, whereas intrapair differences in the M-value correlated inversely with those of 11ß-HSD-1 mRNA and tended to do so with the 50- and 68-kDa protein (Table 2).
Intrapair differences in fat cell size also tended to correlate with 11ß-HSD-1 mRNA and correlated significantly with those in 50- and 68-kDa protein (Table 2). Intrapair differences in adipocyte size were highly correlated with those in body composition as follows: BMI (r = 0.79, P = 0.0002), total body fat (r = 0.79, P = 0.0002), sc adipose tissue mass (r = 0.85, P = 0.0001), and intraabdominal fat mass (r = 0.61, P = 0.01). The association between serum insulin and 11ß-HSD-1 mRNA remained significant after adjusting for intrapair differences in BMI (partial r = 0.51, P = 0.040) or intrapair differences in adipocyte size (partial r = 0.69, P = 0.003). Additionally, intrapair differences in adipocyte size were significantly correlated with intrapair differences in the measures of insulin resistance: homeostasis model assessment (HOMA) (r = 0.52, P = 0.034), serum insulin (r = 0.56, P = 0.019), and M-value (r = –0.57, P = 0.017).
Intrapair differences in fasting serum FFA, triglyceride, and HDL-cholesterol concentrations did not correlate with intrapair differences in sc 11ß-HSD-1 mRNA or protein expression. Instead, intrapair differences in FFAs during hyperinsulinemia correlated significantly with those of 11ß-HSD-1 68-kDa protein expression but not with intrapair differences in 11ß-HSD-1 mRNA or 50-kDa protein expression (Table 2). Intrapair differences in mean arterial pressure did not correlate with those of 11ß-HSD-1 expression (Table 2).
Effects of acquired obesity on GR expression
Intrapair differences in GR expression in sc adipose tissue correlated significantly and inversely with intrapair differences in BMI (r = –0.59, P = 0.012), total body fat (r = –0.67, P = 0.003), and sc adipose tissue mass (r = –0.65, P = 0.044). In contrast, correlations between intrapair differences in intraabdominal adipose tissue mass or plasma insulin concentration were not significant (r = –0.30, P = 0.231 and r = –0.20, P = 0.437, respectively). Intrapair differences in GR mRNA expression correlated significantly and inversely with intrapair differences in fat cell size (r = –0.62, P = 0.008). Intrapair differences in GR mRNA expression did not correlate with 11ß-HSD-1 mRNA or 50- and 68-kDa expression (r = –0.33, P = 0.195; r = –0.48, P = 0.112; and r = –0.45, P = 0.138).
11ß-HSD-1 and GR expression in sc adipose tissue
The effect of acquired obesity on 11ß-HSD-1 expression was further tested in those twin pairs who had the largest differences in BMI (greater than the mean of 3.5 kg/m2). In these pairs, one cotwin was obese (BMI 
30 kg/m2) and the other nonobese (BMI < 25 kg/m2). In the heavier cotwins, 11ß-HSD-1 mRNA and 50- and 68-kDa protein expression were 1.8-, 1.8-, and 1.6-fold higher than in the leaner cotwins (P = 0.008; P = 0.028; and P = 0.046, respectively). An example of a Western blot is shown in Fig. 3. Intrapair differences in 11ß-HSD-1 mRNA levels correlated significantly and positively with those in 11ß-HSD-1 50-kDa protein but not with those in 68-kDa protein levels (r = 0.66, P = 0.020; and r = 0.49, P = 0.106, respectively). Concentrations of GR mRNA did not differ between heavier and leaner twins (data not shown). Gender differences were observed in neither intrapair differences in 11ß-HSD-1 mRNA nor 50- and 68-kDa protein or GR mRNA expression (data not shown).
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Discussion |
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The antibody, which has been raised against an N-terminal fragment of mouse 11ß-HSD-1, was first tested in human primary fibroblasts, in which it recognized two previously described 34- and 68-kDa bands (28). The 68-kDa band was originally identified in the human liver and was suggested to represent a dimeric form of 11ß-HSD-1 (3, 29). To the best of our knowledge, 11ß-HSD-1 protein expression has not been previously studied using Western blotting in human or rodent adipose tissue. We identified three bands corresponding to molecular weights of 34, 50, and 68 kDa in human sc adipose tissue. Based on blocking peptide experiments, all of these were specific for the 11ß-HSD-1 protein. Further support for the notion that the 50- and 68-kDa bands were specific for 11ß-HSD-1 was provided by the correlation analyses, which showed both proteins to be positively correlated with 11ß-HSD-1 mRNA levels. In addition, in intrapair correlation analyses, differences in expression of the proteins correlated with differences in multiple measures of obesity and especially the 50-kDa form with serum insulin (Table 2). In contrast, the 34-kDa band was only occasionally visible and too weak to be reliably quantified in human adipose tissue. The 50-kDa 11ß-HSD-1 protein has not been previously reported in other tissues. Whether the 50-kDa subtype of 11ß-HSD-1 is formed by transcriptional splicing or posttranscriptional modification is not known. Two splice variants have been detected for 11ß-HSD-1 (accession no. NM_005525 and NM_181755), but both encode for the same protein. Topological characterization of 11ß-HSD-1 has revealed a tyrosine motif in the transmembrane helix that is also present in a microsomal 50-kDa esterase/N-acetylases, which has been reported to act as an estrogen receptor luminal targeting sequence (30). However, 11ß-HSD-1 is structurally and functionally unrelated to these genes (31). Whether the 50-kDa protein variant has enzyme activity is not studied in the present study.
Data on effects of obesity on human adipose tissue 11ß-HSD-1 expression/activity have been somewhat heterogeneous and seemingly contradictory. In a recent study, sc 11ß-HSD-1 mRNA and protein activity were measured in adipose tissue biopsies and cultured primary adipocytes isolated from the same biopsies taken from a group of nonobese and obese women undergoing elective surgery (10). No significant correlations were found between 11ß-HSD-1 activity or expression measured in omental or sc adipose tissue and BMI, whereas an inverse correlation was found between BMI and 11ß-HSD-1 activity in cultured omental preadipocytes (10). However, our results are in keeping with the majority of previous studies that have shown that 11ß-HSD-1 activity measured either in vitro or in vivo and/or mRNA concentrations in sc adipose tissue are positively correlated with BMI (4, 6, 8, 11, 32). The 11ß-HSD-1 mRNA expression also correlates with 11ß-HSD-1 enzyme activity (11). The reason for the discrepant data are unknown. Possibly, genetic variation in 11ß-HSD-1 may have confounded the relationship between obesity and 11ß-HSD-1 expression. Mutations resulting in low activity in the gene encoding 11ß-HSD-1 have recently been shown to interact with mutations in the enzyme hexose-6-phosphate dehydrogenase, the normal activity of which is essential for 11ß-HSD-1 function, and cause cortisone reductase deficiency (33).
Whether the adipose tissue 11ß-HSD-1 overexpression is a cause or a consequence of obesity is still uncertain. Transgenic mice overexpressing 11ß-HSD-1 in visceral and sc adipose tissues are hyperphagic and prone to gain weight, especially in the visceral depot, during high-fat feeding (13). These data support the notion that increased activity of 11ß-HSD-1 per se may be a mechanism leading to obesity, but equally plausibly our results indicate that weight gain may cause increased activity of 11ß-HSD-1. However, it is important to emphasize that we did not measure 11ß-HSD-1 in visceral fat, and although we found a positive correlation between expression of sc 11ß-HSD-1 and amount of intraabdominal fat, a significant correlation does not prove causality. Thus, a role of 11ß-HSD-1 overexpression in sc or visceral adipose tissue for development of human obesity is far from proven. Unfortunately, it is not for ethical reasons possible to obtain intraabdominal fat biopsies from healthy twins.
Acquired obesity might be due to expansion of sc adipose tissue mass by either hypertrophy of existing adipocytes or differentiation of adipose tissue stromal cells to adipocytes. In transgenic mice overexpressing 11ß-HSD-1, adipocyte hypertrophy occurred in both sc and visceral adipose tissue depots (13). Our results showed a positive correlation between 11ß-HSD-1 expression and adipocyte size in sc adipose tissue. Therefore, overexpression of 11ß-HSD-1 in sc adipose tissue, as evidenced here during acquired obesity, is associated with expansion of adipose tissue due to adipocyte hypertrophy.
The transgenic 11ß-HSD-1 overexpressing mice showed signs of insulin resistance and increased FFA levels (13). Also in humans, glucocorticoids impair glucose uptake and increase lipolysis in insulin-sensitive tissues (34). Our results showing a correlation between 11ß-HSD-1 overexpression in sc adipose tissue and increased plasma FFA concentrations during hyperinsulinemic conditions are in agreement with results from previous studies showing elevation of serum FFAs after a selective increase in serum cortisol in humans (35). 11ß-HSD-1 overexpression might increase both lipolysis in adipose tissue and lipoprotein lipase activity, which could promote triglyceride storage in adipose tissue depots (36). The latter, in turn, could increase insulin resistance.
In mature adipocytes, glucocorticoids regulate several adipocyte-specific genes, including genes of the local renin-angiotensin system, leptin, and peroxisome proliferator-activated receptor-
(37, 38). GR activation might therefore be important in the generation of the downstream metabolic consequences of obesity. Transgenic 11ß-HSD-1 overexpressing mice exhibited higher GR expression in visceral than sc adipose tissue (13). Therefore, GR-mediated effects on obesity might be more significant in visceral adipose tissue. Consistent with this, we did not see any correlation between intrapair differences in 11ß-HSD-1 and GR expression in sc adipose tissue. This, of course, does not exclude the possibility that GR expression is increased in visceral adipose tissue in humans.
In conclusion, 11ß-HSD-1 gene and protein expression in sc adipose tissue are up-regulated already in the early stages of acquired obesity and directly correlated with sc, total, and intraabdominal fat mass as well as with insulin resistance. This was demonstrated in a unique group of young adult MZ twin pairs, which had a wide range of intrapair differences in BMI and enabled accessing effects of acquired weight gain, independent of genetic factors. The acquired differences in 11ß-HSD-1 protein expression were also positively correlated with those in serum FFA concentrations during hyperinsulinemic conditions, suggesting a relationship between 11ß-HSD-1 protein expression and lipolysis. These results together with those of previous studies support the hypothesis that overexpression of 11ß-HSD-1 could be involved in the pathogenesis of central obesity and insulin resistance. The first, potent selective inhibitor against 11ß-HSD-1 has already been developed (39) and will hopefully enable testing of the validity of the 11ß-HSD-1 hypothesis in subjects with insulin resistance.
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Acknowledgments |
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Footnotes |
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K.K. and K.H.P. contributed equally to this work.
Abbreviations: BMI, Body mass index; FFA, free fatty acid; GR, glucocorticoid receptor; HDL, high-density lipoprotein; HOMA, homeostasis model assessment; 11ß-HSD-1, 11ß-hydroxysteroid dehydrogenase type 1; MRI, magnetic resonance imaging; M-value, whole-body insulin sensitivity of glucose metabolism; MZ, monozygotic; TFIIB, transcription factor IIB.
Received January 30, 2004.
Accepted June 2, 2004.
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References |
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cortisol conversion in subjects with central adiposity. J Clin Endocrinol Metab 84:1022–1027This article has been cited by other articles:
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R. A. De Sousa Peixoto, S. Turban, J. H. Battle, K. E. Chapman, J. R. Seckl, and N. M. Morton Preadipocyte 11{beta}-Hydroxysteroid Dehydrogenase Type 1 Is a Keto-Reductase and Contributes to Diet-Induced Visceral Obesity in Vivo Endocrinology, April 1, 2008; 149(4): 1861 - 1868. [Abstract] [Full Text] [PDF]
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S. Boullu-Ciocca, V. Achard, V. Tassistro, A. Dutour, and M. Grino Postnatal Programming of Glucocorticoid Metabolism in Rats Modulates High-Fat Diet-Induced Regulation of Visceral Adipose Tissue Glucocorticoid Exposure and Sensitivity and Adiponectin and Proinflammatory Adipokines Gene Expression in Adulthood Diabetes, March 1, 2008; 57(3): 669 - 677. [Abstract] [Full Text] [PDF]
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 C. E. Perrone, D. A. L. Mattocks, G. Hristopoulos, J. D. Plummer, R. A. Krajcik, and N. Orentreich Methionine restriction effects on 11 -HSD1 activity and lipogenic/lipolytic balance in F344 rat adipose tissue J. Lipid Res., January 1, 2008; 49(1): 12 - 23. [Abstract] [Full Text] [PDF]
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A. M. Nuotio-Antar, D. L. Hachey, and A. H. Hasty Carbenoxolone treatment attenuates symptoms of metabolic syndrome and atherogenesis in obese, hyperlipidemic mice Am J Physiol Endocrinol Metab, December 1, 2007; 293(6): E1517 - E1528. [Abstract] [Full Text] [PDF]
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 B. R Walker Glucocorticoids and Cardiovascular Disease Eur. J. Endocrinol., November 1, 2007; 157(5): 545 - 559. [Abstract] [Full Text] [PDF]
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Z. Michailidou, A. P Coll, C. J Kenyon, N. M Morton, S. O'Rahilly, J. R Seckl, and K. E Chapman Peripheral mechanisms contributing to the glucocorticoid hypersensitivity in proopiomelanocortin null mice treated with corticosterone J. Endocrinol., July 1, 2007; 194(1): 161 - 170. [Abstract] [Full Text] [PDF]
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D. Qi and B. Rodrigues Glucocorticoids produce whole body insulin resistance with changes in cardiac metabolism Am J Physiol Endocrinol Metab, March 1, 2007; 292(3): E654 - E667. [Abstract] [Full Text] [PDF]
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D. J. Wake, N. Z. M. Homer, R. Andrew, and B. R. Walker Acute In Vivo Regulation of 11{beta}-Hydroxysteroid Dehydrogenase Type 1 Activity by Insulin and Intralipid Infusions in Humans J. Clin. Endocrinol. Metab., November 1, 2006; 91(11): 4682 - 4688. [Abstract] [Full Text] [PDF]
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P. Darmon, F. Dadoun, S. Boullu-Ciocca, M. Grino, M.-C. Alessi, and A. Dutour Insulin resistance induced by hydrocortisone is increased in patients with abdominal obesity Am J Physiol Endocrinol Metab, November 1, 2006; 291(5): E995 - E1002. [Abstract] [Full Text] [PDF]
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B. Mariniello, V. Ronconi, S. Rilli, P. Bernante, M. Boscaro, F. Mantero, and G. Giacchetti Adipose tissue 11{beta}-hydroxysteroid dehydrogenase type 1 expression in obesity and Cushing's syndrome Eur. J. Endocrinol., September 1, 2006; 155(3): 435 - 441. [Abstract] [Full Text] [PDF]
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V. S. Densmore, N. M. Morton, J. J. Mullins, and J. R. Seckl 11{beta}-Hydroxysteroid Dehydrogenase Type 1 Induction in the Arcuate Nucleus by High-Fat Feeding: A Novel Constraint to Hyperphagia? Endocrinology, September 1, 2006; 147(9): 4486 - 4495. [Abstract] [Full Text] [PDF]
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 P. F. Pilch and N. Bergenhem Pharmacological Targeting of Adipocytes/Fat Metabolism for Treatment of Obesity and Diabetes Mol. Pharmacol., September 1, 2006; 70(3): 779 - 785. [Abstract] [Full Text] [PDF]
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K. H. Pietilainen, K. Kannisto, E. Korsheninnikova, A. Rissanen, J. Kaprio, E. Ehrenborg, A. Hamsten, and H. Yki-Jarvinen Acquired Obesity Increases CD68 and Tumor Necrosis Factor-{alpha} and Decreases Adiponectin Gene Expression in Adipose Tissue: A Study in Monozygotic Twins J. Clin. Endocrinol. Metab., July 1, 2006; 91(7): 2776 - 2781. [Abstract] [Full Text] [PDF]
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 M. Grino Prenatal nutritional programming of central obesity and the metabolic syndrome: role of adipose tissue glucocorticoid metabolism Am J Physiol Regulatory Integrative Comp Physiol, November 1, 2005; 289(5): R1233 - R1235. [Full Text] [PDF]
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 S. M. Grundy, J. I. Cleeman, S. R. Daniels, K. A. Donato, R. H. Eckel, B. A. Franklin, D. J. Gordon, R. M. Krauss, P. J. Savage, S. C. Smith Jr, et al. Diagnosis and Management of the Metabolic Syndrome: An American Heart Association/National Heart, Lung, and Blood Institute Scientific Statement Circulation, October 25, 2005; 112(17): 2735 - 2752. [Full Text] [PDF]
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J. M. Paterson, J. R. Seckl, and J. J. Mullins Genetic manipulation of 11{beta}-hydroxysteroid dehydrogenases in mice Am J Physiol Regulatory Integrative Comp Physiol, September 1, 2005; 289(3): R642 - R652. [Abstract] [Full Text] [PDF]
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 S. Arampatzis, B. Kadereit, D. Schuster, Z. Balazs, R. A S Schweizer, F. J Frey, T. Langer, and A. Odermatt Comparative enzymology of 11{beta}-hydroxysteroid dehydrogenase type 1 from six species J. Mol. Endocrinol., August 1, 2005; 35(1): 89 - 101. [Abstract] [Full Text] [PDF]
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R. Basu, R. J. Singh, A. Basu, E. G. Chittilapilly, M. C. Johnson, G. Toffolo, C. Cobelli, and R. A. Rizza Obesity and Type 2 Diabetes Do Not Alter Splanchnic Cortisol Production in Humans J. Clin. Endocrinol. Metab., July 1, 2005; 90(7): 3919 - 3926. [Abstract] [Full Text] [PDF]
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R. Andrew, J. Westerbacka, J. Wahren, H. Yki-Jarvinen, and B. R. Walker The Contribution of Visceral Adipose Tissue to Splanchnic Cortisol Production in Healthy Humans Diabetes, May 1, 2005; 54(5): 1364 - 1370. [Abstract] [Full Text] [PDF]
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G. Apostolova, R. A. S. Schweizer, Z. Balazs, R. M. Kostadinova, and A. Odermatt Dehydroepiandrosterone inhibits the amplification of glucocorticoid action in adipose tissue Am J Physiol Endocrinol Metab, May 1, 2005; 288(5): E957 - E964. [Abstract] [Full Text] [PDF]
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E. E. Kershaw, N. M. Morton, H. Dhillon, L. Ramage, J. R. Seckl, and J. S. Flier Adipocyte-Specific Glucocorticoid Inactivation Protects Against Diet-Induced Obesity Diabetes, April 1, 2005; 54(4): 1023 - 1031. [Abstract] [Full Text] [PDF]
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T. C. Sandeep, R. Andrew, N. Z.M. Homer, R. C. Andrews, K. Smith, and B. R. Walker Increased In Vivo Regeneration of Cortisol in Adipose Tissue in Human Obesity and Effects of the 11{beta}-Hydroxysteroid Dehydrogenase Type 1 Inhibitor Carbenoxolone Diabetes, March 1, 2005; 54(3): 872 - 879. [Abstract] [Full Text] [PDF]
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) in mRNA concentration (n = 17 pairs) (A), 50-kDa protein concentration (n = 12 pairs) (B), and 68-kDa protein concentration (n = 12 pairs) (C) of 11ß-HSD-1 in sc adipose tissue and intrapair differences (