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Weight Loss Increases 11ß-Hydroxysteroid Dehydrogenase Type 1 Expression in Human Adipose Tissue

Division of Medical Sciences, Queen Elizabeth Hospital, University of Birmingham (J.W.T., J.S.M., P.M.S.), Birmingham, United Kingdom B15 2TH; and Regional Endocrine Laboratory, Department of Clinical Biochemistry, University Hospital Birmingham National Health Service Trust (P.M.S.C., G.H., L.S.), Birmingham, United Kingdom B29 6JD

Address all correspondence and requests for reprints to: Dr. Paul M. Stewart, Division of Medical Sciences, University of Birmingham, Queen Elizabeth Hospital, Birmingham, United Kingdom B15 2TH. E-mail: p.m.stewart{at}bham.ac.uk.

	Abstract

Top Abstract Introduction Subjects and Methods Results Discussion References

 
The global epidemic of obesity has heightened the need to understand the mechanisms that underpin its pathogenesis. Clinical observations in patients with Cushing’s syndrome have highlighted the link between cortisol and central obesity. However, although circulating cortisol levels are normal or reduced in obesity, local regeneration of cortisol, from inactive cortisone, by 11ß-hydroxysteroid dehydrogenase type 1 (11ßHSD1) has been postulated as a pathogenic mechanism. Although levels of expression of 11ßHSD1 in adipose tissue in human obesity are debated in the literature, global inhibition of 11ßHSD1 improves insulin sensitivity. We have determined the effects of significant weight loss on cortisol metabolism and adipose tissue 11ßHSD1 expression after 10-wk ingestion of a very low calorie diet in 12 obese patients (six men and six women; body mass index, 35.9 ± 0.9 kg/m²; mean ± SE). All patients achieved significant weight loss (14.1 ± 1.3% of initial body weight). Total fat mass fell from 41.8 ± 1.9 to 32.0 ± 1.7 kg (P < 0.0001). In addition, fat-free mass decreased (64.4 ± 3.4 to 58.9 ± 2.9 kg; P < 0.0001) and systolic blood pressure and total cholesterol also fell [systolic blood pressure, 135 ± 5 to 121 ± 5 mm Hg (P < 0.01); total cholesterol, 5.4 ± 0.2 to 4.8 ± 0.2 mmol/liter (P < 0.05)]. The serum cortisol/cortisone ratio increased after weight loss (P < 0.01). 11ßHSD1 mRNA expression in isolated adipocytes increased 3.4-fold (P < 0.05). Decreased 11ßHSD1 activity and expression in obesity may act as a compensatory mechanism to enhance insulin sensitivity through a reduction in tissue-specific cortisol concentrations. Inhibition of 11ßHSD1 may therefore be a novel, therapeutic strategy for insulin sensitization.

	Introduction

Top Abstract Introduction Subjects and Methods Results Discussion References

 
THE ROLE OF cortisol metabolism in the pathogenesis of obesity is still unclear. There are striking phenotypic similarities among patients with circulating cortisol excess, those with Cushing’s syndrome who develop reversible central obesity, and those patients with simple obesity who have normal (or slightly reduced) circulating cortisol levels (1). The hypothalamic-pituitary-adrenal axis is not normal in obesity. The metabolic clearance rate for cortisol is increased, with a secondary increase in cortisol secretion driven by ACTH (2, 3). It remains unclear whether these observations are a cause or a consequence of the obese phenotype. Their reversibility with weight loss has not been studied.

Within adipose tissue, the enzyme 11ß-hydroxysteroid dehydrogenase type 1 (11ßHSD1) interconverts inactive glucocorticoid cortisone and cortisol. In vivo it is the reductase activity that is believed to predominate, generating cortisol in an autocrine/paracrine manner within the adipocyte microenvironment. Cortisol promotes preadipocyte differentiation, and inhibition of 11ßHSD1 prevents cortisone-induced preadipocyte differentiation (4). Furthermore, mice overexpressing 11ßHSD1 under the adipocyte-specific aP2 promoter develop central obesity as a result of increased adipocyte size (5). However, in human obesity, generation of cortisol from an oral dose of cortisone acetate (believed to largely reflect hepatic 11ßHSD1 activity) is impaired (6, 7, 8). The excretion of the urinary tetrahydrometabolites of cortisol and cortisone [tetrahydrocortisol (THF) plus alloTHF/tetrahydrocortisone (THE) ratio] in the setting of normal urinary free cortisol and cortisone excretion is also believed to reflect global 11ßHSD1 activity (9). The results have been more variable. Some studies have described decreased ratios consistent with decreased 11ßHSD1 reductase activity with increasing body mass index (BMI) in simple obesity (7, 8). Other studies have failed to show this relationship (1, 10, 11, 12), and indeed, positive correlations have been described (6, 13, 14). The explanation for this discrepancy is not clear. A reduction in hepatic 11ßHSD1 expression may not extend to expression in adipose tissue, and a theory has evolved suggesting adipose tissue specific overexpression of 11ßHSD1 in human obesity (8, 15, 16). We have been unable to endorse this; in our studies preadipocyte 11ßHSD1 expression was shown to be lowest in the most obese patients, and we have hypothesized that this may have an important effect to reduce cortisol availability to the glucocorticoid receptor and enhance preadipocyte proliferation (17). The global inhibition of 11ßHSD1 in human obesity and, therefore, the lack of ability to regenerate cortisol within tissues are likely to be responsible for the observed increased cortisol metabolic clearance rate and cortisol secretion rates.

The aim of this study was to perform a detailed characterization of the global and adipose tissue-specific expression and activity of 11ßHSD1 with significant weight loss in human obesity.

	Subjects and Methods

Top Abstract Introduction Subjects and Methods Results Discussion References

 
The study had the approval of the local research ethics committee, and all patients gave their full, informed, written consent. Twelve obese patients (six men and six women) were recruited from local advertisement (median age, 49 yr; range 23–58; mean BMI, 35.9 ± 0.9 kg/m²). Patients had no significant past medical history and were taking no regular medications. Fasting screening bloods were drawn at 0900 h, and all patients had normal blood counts, glucose levels, and renal, liver, and thyroid functions. Patients with impaired fasting glucose (>6.0 mmol/liter) or diabetes mellitus were excluded from the study. All patients had normal resting electrocardiograms.

Once enrolled in the study, all patients had further fasting blood samples drawn at 0900 h for measurement of total cholesterol, triglycerides, cortisol, cortisone, glucose, and insulin. Measurements of BMI, waist (measured supine, at the level of the umbilicus) and hip (at the level of the greater trochanter) circumference, and blood pressure [average of three readings, measured supine after 10 min of rest using Dynamap (Critikon, Tampa, FL)] were also taken. In addition, all patients performed a 24-h urine collection for corticosteroid metabolite analysis using gas chromatography/mass spectrometry as previously described (9). Measurement of the ratio of total cortisol metabolites Fm (total cortisol metabolites = cortisol + THF + 5THF + -cortol + ß-cortol + 20-dihydrocortisol + 20ß-dihydrocortisol + 6ß-hydroxycortisol) to total cortisone metabolites Em (total cortisone metabolites = cortisone + THE + -cortolone + ß-cortolone + 20DHE + 20ßDHE) provides an assessment of the global set-point of cortisol to cortisone conversion. More specifically, the ratio of tetrahydrometabolites of cortisol (THF + 5THF) to those of cortisone (THE) provides a reflection of 11ßHSD1 activity, providing that the ratio of urinary free cortisol to cortisone (reflecting renal 11ßHSD2 activity) is unaltered. The activities of 5- and 5ß-reductases can be inferred from measurement of the THF/5THF ratio.

To provide a more accurate reflection of hepatic 11ßHSD1 activity, a cortisol generation curve was performed for all patients as previously described (18). Briefly, 1 mg dexamethasone was taken orally at 2400 h the night preceding the test. At 0845 h the patients were cannulated in an antecubital fossa vein, blood samples were drawn at 0900 h, and then an oral dose of cortisone acetate (25 mg) was given. The subsequent generation of cortisol in serum was then followed by sampling at 30, 60, 90, 12, 150, 180, and 240 min.

Body composition analysis was performed using dual energy x-ray absorptiometry with a total body scanner (Lunar DPX-L, Lunar Corp., Madison, WI). Coefficients of variation for multiple scans were less than 3%. Regional fat mass (trunk and leg) was analyzed as previously described (18).

In addition, all patients had an sc buttock biopsy performed under local anesthetic to obtain approximately 1–2 g adipose tissue. The sample was divided into two parts. Half was used for adipocyte isolation and subsequent total RNA extraction and preadipocyte culture, and the remaining half was snap-frozen in liquid nitrogen and stored at –70 C for whole tissue total RNA extraction (see below). All samples were processed within 30 min after the biopsy.

After all investigations had been completed, patients were entered into a weight loss program using total meal replacement, very low calorie diet (Lipotrim, Howard Foundation, Cambridge, UK). This diet provides 425 (women) and 559 (men) kcal/d. The median duration of dietary intervention was 10 wk (range, 8–14 wk). After significant weight loss (>10% initial body weight), all subjects returned to a normal diet, and once refeeding had been commenced for at least 1 wk, all of the investigations described above were repeated. Investigations were not repeated sooner so as to avoid the confounding effect that the stress of the hypocaloric diet may have had on the hypothalamic-pituitary-adrenal axis.

Adipocyte isolation and preadipocyte culture

Adipocytes and preadipocytes were isolated as previously reported (19, 20). Briefly, adipose tissue biopsies were washed in PBS containing 50,000 U/liter penicillin and 50,000 µg/liter streptomycin (Life Technologies, Inc., Paisley, UK). The tissue was then prepared and digested with collagenase class 1 (2 mg/ml; Worthington Biochemical Corp., Reading, UK) in 1x Hanks’ Balanced Salt Solution (Life Technologies, Inc.) for 45 min at 37 C. Samples were centrifuged at 90 x g for 1 min, the intact adipocyte layer was removed, and RNA was extracted as described below. Samples were then centrifuged at 90 x g for 5 min, the pellet containing preadipocytes was removed, and cells were washed with DMEM/Nutrient Mixture F-12 (Life Technologies, Inc.) containing 15% fetal calf serum (Life Technologies, Inc.) and seeded on 96-well plates (Costar, Cambridge, MA). Cells were left overnight and washed the following day with 1x Hanks’ Balanced Salt Solution. Proliferation assays were performed on d 1, 4, and 7 of culture (see below).

RNA extraction and RT

Total RNA was extracted using a single step extraction method [Tri-Reagent (Sigma-Aldrich, Poole, UK; adipocytes) or Genelute total mammalian RNA extraction kit (Sigma-Aldrich; whole adipose tissue)]. RNA integrity was assessed by electrophoresis on 1% agarose gels, and quantity was determined spectrophotometrically at OD₂₆₀. One microgram of total RNA was initially denatured by heating to 70 C for 5 min. Thirty units of avian myeloblastosis virus, 200 ng random primers, 20 U ribonuclease inhibitor, and 40 nmol deoxy-NTPs with 5x reaction buffer were added to the RNA, and the reverse transcriptase reaction was carried out at 37 C for 1 h. The reaction was terminated by heating the cDNA to 95 C for 5 min.

Real-time PCR

11ßHSD1 mRNA levels were analyzed using an ABI 7700 sequence detection system (PerkinElmer Biosystems, Warrington, UK), which employs TaqMan chemistry for highly accurate quantification of mRNA levels as previously described (21). RT-PCR was performed in 25-µl volumes on 96-well plates in reaction buffer containing TaqMan universal PCR master mix (PerkinElmer), 3 mM Mn(Oac)₂, 200 µM deoxy-NTPs, 1.25 U AmpliTaq Gold polymerase (PerkinElmer), 1.25 U AmpErase UNG (PerkinElmer), 100–200 nmol TaqMan probe, 900 nmol primers, and 25–50 ng cDNA. All reactions were multiplexed with the housekeeping gene (18S), provided as a preoptimized control probe (PerkinElmer), enabling data to be expressed in relation to an internal reference to allow for differences in RT efficiency. Data were obtained as ct values according to the manufacturer’s guidelines (the cycle number at which logarithmic PCR plots cross a calculated threshold line) and were used to determine ct values (ct of the target gene minus ct of the housekeeping gene). Fold changes in expression were calculated according to the transformation: fold increase = 2^{– difference in ct}. All target gene probes were labeled with the fluorescent label 6-carboxyfluorescein, and the housekeeping gene was labeled with the fluorescent label VIC. Reactions were as follows: 50 C for 2 min, 95 C for 10 min, and then 44 cycles of 95 C for 15 sec and 60 C for 1 min. To exclude potential bias caused by averaging data that had been transformed through the equation 2^–ct, all statistics were performed at the ct stage.

Oligonucleotide primers and a TaqMan probe for 11ßHSD1 were as follows: forward, AGGAAAGCTCATGGGAGGACTAG; reverse, ATGGTGAATATCATCATGAAAAAGATTC; and probe, CATGCTCATTCT CAACCACATCACCAACA.

Preadipocyte proliferation

Preadipocyte proliferation was assessed using a commercially available colorimetric assay for determining the number of viable cells (Promega, Madison, WI) according to the manufacturer’s guidelines with appropriate controls (no cells). Cells were incubated with reagents for 1 h at 37 C in a humidified 5% CO₂ atmosphere. The absorbance at 490 nm reflects the number of living cells and was measured using a 96-well plate reader. Readings were performed in triplicate, and the mean no-cell control reading was subtracted from the mean of the cell-containing wells.

Biochemical assays

Cortisol was assayed using a chemiluminescent immunoassay (Bayer Advia Centaur, Bayer Diagnostics, Newbury, UK) with interassay coefficients of variation of 10.2% at 76 nmol/liter, 7.7% at 528 nmol/liter, and 7.4% at 882 nmol/liter. Cortisone was assayed after extraction from serum, followed by RIA of the extract with [¹²⁵I]cortisone and Sac-Cel (IDS Ltd., Tyne and Weir, UK) second antibody separation. The coefficient of variation for 10 consecutive assays was less than 15% for values between 50 and 100 nmol/liter and less than 10% for values greater than 100 nmol/liter. Serum insulin was measured using an immunoenzymometric assay with no significant cross-reactivity with proinsulin(s), calibrated against IRP 66/304 (Medgenix Insulin-EASIA, BioSource, Camarillo, CA). Interassay coefficients of variation were less than 10% over the range 95–1038 pmol/liter.

Urea, creatinine, electrolytes, cholesterol, and triglycerides were measured using standard laboratory methods (Roche Modular System, Roche, Lewes, UK). Glucose was assayed using the hexokinase method (Instrumentation Laboratory, Warrington, UK), with interassay coefficients of variation of 2.0% at 4.7 mmol/liter and 1.7% at 33.4 mmol/liter.

Insulin sensitivity was derived from fasting glucose and insulin data, using the homeostasis model assessment (HOMA) mathematical model for specific insulin assays based on the method previously described (22) (HOMA-R values greater than 1 suggested insulin resistance).

Statistical analysis

The data are presented as the mean ± SE unless otherwise stated. For comparison of outcomes before and after weight loss, paired t tests were used. Where multiple comparisons were made, repeated measure ANOVA was used. For real-time PCR data, statistical analysis was performed on ct values, rather than fold changes.

	Results

Top Abstract Introduction Subjects and Methods Results Discussion References

 
Body composition

Treatment with a very low calorie diet resulted in significant weight loss in all subjects. Mean BMI fell from 35.9 ± 0.9 to 30.7 ± 0.7 kg/m² (P < 0.0001). There were significant reductions in total and regional fat mass as well as smaller reductions in total and regional lean mass. The mean reduction in total fat mass was 9.7 ± 1.5 kg, and the mean reduction in total lean mass was 5.6 ± 0.7 kg (Table 1). In addition, alterations in body fat distribution were observed. The trunk/leg fat mass ratio fell (1.60 ± 0.15 to 1.34 ± 0.11; P < 0.05), indicative of selective central fat loss. The waist/hip ration also fell, although this did not reach statistical significance (Table 1).

Despite significant alterations in body composition, fasting glucose did not change after weight loss. However, fasting insulin levels decreased significantly (P < 0.01), and insulin resistance decreased, as measured by the glucose/insulin ratio (P < 0.05) and HOMA analysis (P < 0.01). Fasting total cholesterol fell, although triglyceride concentrations remained unaltered (Table 2).

Systolic blood pressure fell from 135 ± 5 to 121 ± 5 mm Hg (P < 0.01) after weight loss. Although diastolic blood pressure decreased, this failed to reach statistical significance (76 ± 3 vs.72 ± 1 mm Hg; P = 0.20).

Corticosteroid metabolism

Circulating cortisol and cortisone concentrations did not change with weight loss [0900 h cortisol, 287 ± 24 vs. 355 ± 36 nmol/liter (P = 0.1); 0900 h cortisone, 60 ± 3 vs. 60 ± 5 nmol/liter (P = 0.7)]. However, the 0900 h cortisol/cortisone ratio increased, indicative of a shift in set-point toward cortisol generation consistent with increased 11ßHSD1 activity (Table 3). Urinary corticosteroid metabolites analysis using gas chromatography/mass spectrometry demonstrated a significant decrease in total 24-h production of cortisone metabolites, again consistent with increased 11ßHSD1 reductase activity. In addition, we observed a borderline significant decrease in the 24-h urinary cortisol secretion rate (P = 0.06) and THE (P = 0.06). Other corticosteroid metabolites did not change with weight loss (Table 3).

View larger version (15K): [in this window] [in a new window]   FIG. 1. Cortisol generation curves before and after significant weight loss in 12 obese subjects. After overnight dexamethasone suppression (1 mg), generation of cortisol in the serum after oral cortisone acetate (25 mg) is measured by repeated serum sampling. Cortisol generation does not differ before or after weight loss at any time point.

Successful RNA extraction and RT were performed in 11 patients. Whole adipose tissue expression of 11ßHSD1 did not change with weight loss (13.5 ± 0.2 vs. 14.1 ± 0.3 ct; P = 0.11, before vs. after). However, 11ßHSD1 expression increased in 10 of the 11 patients. Adipocyte 11ßHSD1 expression increased 3.4-fold (11.6 ± 0.4 vs. 9.9 ± 0.6 ct; P < 0.05, before vs. after; Fig. 2).

View larger version (14K): [in this window] [in a new window]   FIG. 2. A, 11ßHSD1 expression in human adipocytes before and after weight loss in 11 individuals as measured using real-time PCR (PerkinElmer; data are expressed as ct, with high values representing low expression). B, Mean whole adipose tissue 11ßHSD1 expression does not alter with weight loss. However, mean adipocyte-specific expression increases 3.4-fold.

View larger version (11K): [in this window] [in a new window]   FIG. 3. Preadipocyte cell number estimation using colorimetric cell proliferation assay (CellTitre96 One Solution Proliferation assay, Promega) in sc buttock biopsies from obese individuals before and after weight loss. Cell number estimations were performed on d 1, 4, and 7 of culture in serum-containing medium. The OD490nm reflects the number of intact viable cells. Cell number does not differ at any time point.

	Discussion

Top Abstract Introduction Subjects and Methods Results Discussion References

The role of 11ßHSD1 in adipocyte biology and in human obesity is still unclear. It is more highly expressed in omental compared with sc preadipocytes (20). We have postulated that in preadipocytes it may act to limit proliferation (17), and in adipocytes it may promote differentiation (4). To support this, in vitro experiments have shown that inhibition of 11ßHSD1 with glycerrhetinic acid blocks the prodifferentiative and antiproliferative actions of cortisone (4, 17). Further evidence for its differentiative role is derived from mice overexpressing 11ßHSD1 under the aP2 promoter, which develop obesity as a result of increased adipocyte size due to a tissue-specific increase in corticosterone concentrations (5). In this study we were unable to demonstrate significant alterations in the preadipocyte proliferation rate after weight loss.

Much of the current literature has discussed the abnormalities of cortisol metabolism as possible causative factors. However, it is equally possible that they arise as a consequence of the obese phenotype. Detailed studies of cortisol metabolism before and after weight loss have not been performed, but circulating levels do not appear to change (23). Cortisol-binding globulin (CBG) decreases with weight loss (24). Although not measured in this study, a decrease in CBG would cause a decrease in the cortisol/cortisone ratio, with cortisone being essentially unbound to CBG. Alterations in CBG are therefore unlikely to explain the observed changes in circulating glucocorticoids.

Although this study was not designed to determine levels of 11ßHSD1 in obesity, the global inhibition of 11ßHSD1 that has previously been reported in obesity may represent a compensatory mechanism by which tissue-specific cortisol concentrations are reduced in an attempt to improve insulin sensitivity in obese, insulin-resistant individuals (7). With significant weight loss, this regulatory mechanism is no longer required. Evidence for the role of 11ßHSD1 in the control of insulin sensitivity comes from a wide variety of rodent and clinical studies. Healthy, nonobese, volunteers treated with carbenoxolone to inhibit 11ßHSD activity display improved insulin sensitivity (25). In addition, in glucose-intolerant individuals, 11ßHSD1 activity, as measured by cortisol generation curves, is impaired (26). One interpretation of these data is that down-regulation of 11ßHSD1 activity may represent a mechanism to improve insulin sensitivity in a glucose-intolerant individual. This hypothesis is also supported by rodent models. The 11ßHSD1 knockout mouse displays relative insulin sensitivity (27), and in other models of rodent obesity there is a compensatory decrease in hepatic (28) and adipose tissue (29) 11ßHSD1 expression.

Patients with apparent cortisone reductase deficiency are unable to activate oral cortisone acetate to cortisol (30) and represent the putative human 11ßHSD1 knockout (31, 32, 33, 34). We have recently defined the molecular basis for the disease caused by intronic mutations within the HSD11B1 gene that decrease transcription in combination with mutations in hexose-6-phosphate dehydrogenase, an enzyme believed to provide NADPH for 11ßHSD1, which is essential for reductase activity (35). If our hypothesis is correct, these patients should be relatively insulin sensitive due to decreased tissue-specific cortisol concentrations; however, such studies have not been performed.

Selective 11ßHSD1 inhibition remains an exciting therapeutic prospect. These drugs are not available as yet for studies in humans. However, recently a novel class of agents has been described (arylsulfonamidothiazoles) that, unlike the liquorice derivatives, has a greater than 200-fold selectivity for inhibition of 11ßHSD1 rather than 11ßHSD2. Rodents treated with these drugs show significant improvements in insulin sensitivity (36). These drugs, therefore, have considerable potential as adjunctive insulin sensitizers in the treatment of type 2 diabetes mellitus, and their use may extend to patients with the metabolic syndrome. Clinical studies, using gold standard techniques, including hyperinsulinemic, euglycemic clamps, specifically and carefully designed to test the hypothesis that modulation of 11ßHSD1 activity may impact upon tissue specific insulin sensitivity are now warranted. Although the results from this study may point to this role, we must be careful not to overinterpret the data. The aim of this study was to characterize the changes in 11ßHSD1 activity and expression that occur with weight loss, rather than to investigate the functional consequences of this change in expression profile. The control of insulin sensitivity is a complex multifactorial process. The degree to which insulin sensitivity may be improved with 11ßHSD1 inhibition in human studies is not known. A further important consideration when interpreting the data from this study is that all fat depots are not identical; they contribute differently to mortality and morbidity (15, 37). In this study we have used sc gluteal adipose tissue. Depot-specific patterns of gene expression as well as differing patterns of differentiation in cell culture are well described (38, 39).

The mechanism that underpins this regulation of 11ßHSD1 with obesity and weight loss is not clear. 11ßHSD1 is regulated by a wide variety of growth factors, hormones, and cytokines in a tissue-specific manner (40). With the recognition of adipose tissue as an endocrine organ and with the rapidly increasing list of adipokines, it is possible that a factor produced locally within the adipocyte microenvironment may have an important regulatory role. Large population studies have failed to show associations between intronic microsatellite markers and body mass index (41), and in smaller studies, no coding sequence mutations of the HSD11B1 gene have been identified in patients with abdominal obesity (42). The true mechanism that determines 11ßHSD1 activity in human obesity remains to be defined.

This study has broadened our understanding of the role of prereceptor cortisol metabolism within adipose tissue. Tissue-specific cortisol excess has never been demonstrated in simple human obesity. Indeed, with weight loss there may be a rise in adipose tissue cortisol concentrations reflecting increased adipocyte expression of 11ßHSD1. Although we and others have speculated that the role of 11ßHSD1 may be to control preadipocyte proliferation and adipocyte differentiation, it may have an important role in determining tissue-specific insulin sensitivity.

	Footnotes

 
Abbreviations: BMI, Body mass index; CBG, cortisol-binding globulin; HOMA, homeostasis model assessment; 11ßHSD1, 11ß-hydroxysteroid dehydrogenase type 1; THE, tetrahydrocortisone; THF, tetrahydrocortisol.

Received August 6, 2003.

Accepted March 9, 2004.

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