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Original Studies |
Departments of Internal Medicine (J.S.F., R.C.H., S.B.R., E.F.B., W.M.K.) and Pediatrics (M.L.), Washington University School of Medicine, St. Louis, Missouri 63110
Address all correspondence and requests for reprints to: Dr. Wendy M. Kohrt, Department of Internal Medicine, Box 8113, Washington University School of Medicine, 660 South Euclid Avenue, St. Louis, Missouri 63110. E-mail: wkohrt{at}imgate.wustl.edu
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Abstract |
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 Top Abstract IntroductionSubjects and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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Saad et al. have recently shown a positive relationship between maximal insulin-stimulated glucose disposal and the degree of increase in plasma leptin during hyperinsulinemia (8). This suggests that individual differences in insulin responsiveness could help explain the variable findings regarding leptinemic responses to hyperinsulinemia mentioned above (2, 3, 4, 5, 6, 7, 9). If insulin is acting to influence the plasma leptin concentration by specifically altering leptin production by adipocytes, insulin-induced changes in leptinemia should be more closely associated with insulin action on adipocytes per se than with the whole body insulin action studied by Saad et al. (8). We therefore examined the relationships between indexes of global and regional lipolysis (a relatively specific marker of adipocyte metabolism) and changes in plasma leptin during physiological hyperinsulinemia. Whole body lipolysis was assessed by stable isotope kinetics, and lipolysis in abdominal and femoral sc adipose tissue was evaluated by microdialysis.
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Subjects and Methods |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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Fifteen premenopausal (mean ± SE, 33 ± 2 yr) women with no history of diabetes or cardiovascular disease participated in the study. Mean body weight was 70 ± 5 kg (range, 50–118 kg), and body mass index was 26 ± 2 kg/m2. Body fat was 32 ± 4% (range, 14–59%). All subjects had normal glucose tolerance and had had normal monthly menses during the previous year. Subjects gave informed consent before participating, and all procedures were approved by the institutional review board at Washington University School of Medicine.
Oral glucose tolerance test
Subjects reported after an overnight fast. After a baseline blood sample was taken, subjects consumed 75 g glucose. Blood samples were taken 30, 60, 90, and 120 min thereafter. Plasma glucose was determined by the glucose oxidase method (Beckman Coulter, Inc., Fullerton, CA). Normal glucose tolerance was defined as no blood glucose value more than 200 mg/dL and a 120 min value less than 140 mg/dL (10).
Body composition
Body fat was measured by dual energy x-ray absorptiometry (QDR-1000/w, Hologic, Inc., Waltham, MA), using version 5.64 of the enhanced whole body analysis program (11).
Microdialysis
Inlet (30-cm) and outlet (15-cm) tubings of microdialysis probes
(DL-3, Bioanalytical Systems, Inc., West Lafayette, IN)
were separated by 3-cm of polyacrylonitrile dialysis membrane (id, 0.25
mm; od, 0.35 mm). After ethylene oxide gas sterilization, probes were
rinsed in 20% ethanol for 20 min, then soaked overnight (
16 h) in
5% ethanol while being perfused with Ringer’s solution to remove
glycerol from the dialysis membrane.
Microdialysis probes were inserted with sterile technique into sc
adipose tissue under local anesthesia. A 14-gauge catheter was inserted
in the adipose tissue parallel to the skin, entering and exiting the
skin about 6 cm apart. The microdialysis probe was threaded through the
catheter, and the catheter was removed, leaving the probe embedded in
the adipose tissue. Probes were placed in abdominal adipose tissue (two
probes, 3 cm on each side of the umbilicus) and femoral adipose
tissue (two probes at the midthigh, 3 cm apart) in each subject.
Microdialysis probes were perfused (CMA model 102 microdialysis pump, Stockholm, Sweden) at 2.0 µL/min with Ringer’s solution containing 2.5 mmol/L glucose and 5 mmol/L ethanol. The glucose was added to the perfusate to minimize loss of glucose from the interstitial fluid. Ethanol was included in the perfusate to permit detection of changes in blood flow in the region surrounding the probe (12, 13). Probes were allowed to equilibrate for 1 h before collection of microdialysate samples to allow the trauma from probe insertion to subside (14, 15). Dialysate was collected every 10 min for baseline measures (20 µL) and every 15 min during insulin infusions (30 µL). Dialysates were stored at 4 C and analyzed within 48 h for glycerol (16) and ethanol (14).
We have found that the variation for in vitro glycerol recovery is approximately 2% from probe to probe and from lot to lot. In vivo variability in recovery is dependent on both the individual probe and the site to site physiological differences within a given adipose tissue depot. In the current study, the mean within-subject, within-depot coefficients of variation for dialysate glycerol concentrations were 16.7 ± 3.9% (basal), 15.3 ± 2.9% (low dose insulin), and 13.5 ± 2.6% (moderate dose insulin).
Recovery is optimized at low flow rates and approaches 100% at flow rates below 0.3 µL/min. At these low flow rates, the time required to accumulate enough sample for assays would be prohibitive. For these experiments, we perfused the probes only at 2.0 µL/min. In previous experiments, we found that dialysate glycerol concentrations when probes were perfused at 2.0 µL/min were highly correlated (r = 0.84; n = 10; P = 0.002) with calculated interstitial glycerol concentrations based on the no net flux method (data from Ref. 17). We therefore believe that dialysate glycerol concentrations are a good marker of interstitial glycerol concentrations.
We have previously found glycerol recovery by the probes used in this study to be about 60% in abdominal sc adipose tissue of premenopausal women (17). However, we did not measure recovery in the current study, and we have previously found that glycerol recovery is reduced as skinfold thickness increases (unpublished data). To corroborate findings for unadjusted data, we also performed statistics on dialysate glycerol concentrations that were adjusted for differences in recovery. Recovery was computed with a regression equation that we developed from a group of women similar to that in the current study (17): recovery, -0.863 x (mm skinfold thickness) + 67.97% (unpublished equation).
The observation that probe recovery is affected by skinfold thickness has been made in two research laboratories other than our own (18, 19). Several factors that are known to affect recovery, such as tortuosity (dependent in part on interstitial water content), blood flow, or effective membrane surface area, can change with increasing adipose tissue thickness. Therefore, factors related to tissue thickness could affect recovery over a probe located entirely within adipose tissue.
The ethanol microdialysis technique for assessment of adipose tissue blood flow in humans has been validated with the xenon 133 clearance technique (20). The concentration of ethanol infused has been found to have no effect on lipolysis in sc adipose tissue (12). The ratio of ethanol concentration recovered in the dialysate to that in the perfusate (outflow/inflow ratio) is inversely proportional to blood flow in the tissue surrounding the probe (13).
Hyperinsulinemic-euglycemic clamp
Clamp studies were performed at the Washington University General Clinical Research Center after an overnight fast, as previously described (21). Baseline blood and dialysate samples were obtained 30, 20, 10, and 0 min before the start of insulin infusion. The hyperinsulinemic-euglycemic clamp consisted of two 90-min stages of primed, constant insulin infusion. In leaner subjects (<32% body fat), insulin was infused at 10 and 20 mU/m2·min, whereas in more obese subjects (>35% body fat) insulin infusion rates were 6 and 12 mU/m2·min. We anticipated that these infusion rates would result in similar insulin concentrations in lean and obese subjects (22).
Blood samples were taken every 5 min during insulin infusions for determination of blood glucose. Blood glucose was maintained at approximately 5 mmol/L by modulating the infusion rate of 20% dextrose. Blood samples were obtained at 15-min intervals during the clamp for determination of substrate and hormone concentrations. Samples were stored at -80 C for determination of glycerol (16), insulin (23), and catecholamines (24) and at -20 C for determination of leptin (25).
Glycerol kinetics
A primed (1.5 µmol/kg), constant (0.1 µmol/kg·min)
infusion of 2H5-glycerol
(99%; Tracer Technologies, Newton, MA) was started 90 min before the
start of the first insulin infusion and continued until the end of the
clamp. A blood sample taken before the start of the isotope infusion
was used to determine background isotope enrichment. The actual isotope
delivery rate was determined for each infusion by assay of the infusate
enrichment.
Analysis of 2H5-glycerol was performed using a modification of the negative ion chemical ionization (NCI) gas chromatography-mass spectrometry method (26). Plasma (50 µL) was added to 500 µL 3 mol/L perchloric acid, incubated at 4 C for 20 min, and centrifuged for 10 min at 2000 x g. The supernatant was evaporated under nitrogen and reconstituted with 150 µL heptafluorobutyric acid-ethyl acetate (3:1) to form tris-heptafluorobutyryl ester derivatives, incubated at 70 C for 10 min, and dried under nitrogen. After reconstitution in 100 µL ethyl acetate, the NCI spectrum of the heptafluorobutyric acid derivative was obtained (gas chromatography-mass spectrometry, model 5988A, Hewlett-Packard Co., Palo Alto, CA). A DB-17 column (30 m; id, 0.25 mm; 0.25-µm film thickness; J and W Scientific, Folsom, CA) was used with a helium flow rate of 0.5 mL/min and a split ratio of 20:1. The column temperature was set at 100 C for 1 min and was increased 45 C/min to 280 C. The mass spectrometer conditions were: source temperature, 120 C; injector port and detector temperature, 250 C; emission, 300 µA; and ion source pressure, 0.5–0.6 torr. Methane was the Cl- reactant gas, and the concentration of M- ions (nominal mass, 680 and 685 for unlabeled and labeled glycerol, respectively) was monitored. The rate of appearance (Ra) of glycerol over the last 30 min of each clamp stage was calculated using nonsteady state equations (27), assuming a volume of distribution for glycerol of 270 mL/kg (28). The coefficient of variation for day to day reproducibility of isotopic enrichment was 7.1 ± 2.2% (n = 6).
Leptin analysis
The concentration of leptin in plasma samples was determined by RIA (25) with reagents from Linco Research, Inc. (St. Charles, MO). The antibody does not detect human insulin, proinsulin, glucagon, pancreatic polypeptide, or somatostatin, and intra- and interassay coefficients of variation are less than 8% (25).
Statistics
Differences between baseline measures and measures during
insulin infusions were evaluated by repeated measures ANOVA followed by
paired t tests with Bonferroni adjustments to compare means
at the different stages. Basal measures for insulin, glycerol, and
ethanol outflow/inflow ratio were computed as the mean of the four
baseline measurements from the half-hour before the beginning of the
insulin infusions (-30, -20, -10, and 0 min). Values for the low and
moderate insulin clamp stages were computed as the mean value for the
final half-hour of each stage. Changes in leptin during the clamp
stages were determined from the 0, 90, and 180 min values and expressed
as percentages of baseline measures. Correlations between variables
were determined by regression analysis. For all statistical procedures,
was 0.05. Data are presented as the mean ±
SE.
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Results |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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Plasma leptin did not change during either stage of the
hyperinsulinemic clamp (Table 1
). Femoral
dialysate glycerol fell to about 80% and 60% of basal level during
the first (P < 0.05 vs. basal) and second
(P < 0.05 vs. both basal and low dose)
insulin infusions, respectively (Table 1). Abdominal dialysate glycerol
concentration was suppressed to approximately 65% and 50% of baseline
during the two clamp stages (P < 0.05, basal
vs. low and moderate, low vs. moderate). Glycerol
Ra was suppressed to approximately 50% and 40%
of the basal rate during the clamp (P < 0.05, basal
vs. both insulin stages). There were no differences in
plasma catecholamine concentrations between the basal state and the
insulin infusions.
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There were slight decreases in ethanol outflow/inflow ratios
during the insulin infusions, indicative of an increased blood flow
(Table 1
). The abdominal outflow/inflow ratio did not change from
baseline to the first clamp stage, but in the second stage, it was
reduced compared to baseline and first stage values (P
< 0.05). Femoral outflow/inflow decreased from baseline during the
first stage (P < 0.05) and was further reduced in the
second stage (P < 0.05).
Correlations
The change in leptin during insulin infusions was inversely
related (r = -0.44; P < 0.05) to the abdominal
dialysate glycerol concentration for both clamp stages combined
(changes in plasma leptin during a stage are plotted against the
abdominal dialysate glycerol concentration for that stage; Fig. 1A). Changes in leptin were correlated
with abdominal dialysate glycerol during the first clamp stage (r
= -0.55; P < 0.05), but the correlation was not
statistically significant in the during the second stage (r =
-0.39). Changes in leptin were unrelated to femoral dialysate glycerol
(Fig. 1B), glycerol Ra (Fig. 2), or plasma insulin (Fig. 3).
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) were correlated with
abdominal dialysate glycerol concentrations from both the first clamp
stage (r = -0.57; P < 0.05) and the second clamp
stage (r = -0.57; P < 0.05).
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The reported findings that skinfold thickness affects recovery (18, 19) do not invalidate the conclusions of this manuscript. The mathematical correction for changes in dialysate concentration based on skinfold thickness does add additional variability to the data. However, the lack of substantial difference in the correlation coefficients for the relationships using uncorrected (r = -0.44) and corrected (r = -0.45) abdominal dialysate concentrations suggests that the uncorrected data are robust.
We analyzed the relationships between changes in leptin and changes in abdominal dialysate glycerol. We found a significant relationship for the change in abdominal dialysate glycerol during the low dose infusion vs. the change in leptin over the entire clamp (r = -0.60; P < 0.05), but not for changes in dialysate glycerol and plasma leptin within stages.
We have plasma leptin values for nine of the subjects taken 90 min before the start of the insulin infusions. The decline in leptin during this 90-min baseline period (-6.8 ± 2.7%) was not related to the baseline abdominal adipose tissue dialysate glycerol concentration (r = 0.02; P > 0.96).
There was no relationship between the relative changes in ethanol outflow/inflow ratios and relative changes in dialysate glycerol concentrations for either abdominal (r = -0.03) or femoral (r = -0.004) adipose tissue.
Although the baseline plasma leptin concentration was highly correlated with body fat mass (r = 0.97; P < 0.05), changes in leptin during the complete clamp were not significantly related to body fat mass (r = -0.44; P > 0.10).
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Discussion |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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We found considerable variability in the leptinemic response to physiological insulin infusions. Plasma leptin rose in some subjects during insulin infusions, whereas leptin levels in others continued to fall, as if they were fasting. Plasma leptin levels tended to increase the most in those subjects whose abdominal sc adipose tissue was the most sensitive to the antilipolytic effects of insulin.
We found that the leptinemic response to insulin infusion was inversely related to a measure of lipolysis (dialysate glycerol concentration) in abdominal sc adipose tissue, but not to measures of femoral or whole body lipolysis. It has been reported that human abdominal sc fat secretes 2–3 times more leptin in vitro and contains more leptin messenger ribonucleic acid than omental fat (29). Furthermore, the fasting plasma leptin concentration has been found to be related to total sc fat and abdominal sc fat, but not to visceral fat (30). It appears that sc fat exerts more influence on plasma leptin expression than visceral fat, and our data suggest that insulin-associated changes in plasma leptin in women may be more related to the lipolytic state of abdominal sc fat than other (i.e. femoral) sc fat depots.
The apparent inverse association of the level of lipolysis in abdominal sc fat with changes in plasma leptin levels is consistent with the hypothesis that leptin is a lipostatic messenger. We propose that when adipocytes are in the lipogenic state and lipolysis is low, leptin production is highest. Conversely, when adipocytes are most lipolytic, leptin production is lowest. In vivo and in vitro evidence supports this hypothesis. For example, isoprenaline, a ß-adrenergic agonist that stimulates lipolysis, reduced leptin production by 20% after 90 min of iv infusion in humans (31). Furthermore, the isoprenaline-induced reduction in plasma leptin was reversed within 15 min of the cessation of isoprenaline infusion (31). Similarly, when lipolysis was stimulated by cAMP in human visceral adipocytes in vitro, leptin secretion was decreased (32). In addition, incubation of rat adipocytes with dibutyryl cAMP or the ß-agonists isoproterenol and isoprenaline led to decreased leptin secretion (33, 34). On the other hand, maximal suppression of lipolysis by acipimox, a nicotinic acid analog, caused an approximately 5% increase in plasma leptin when infused for 3 h in humans, whereas, in contrast, leptin fell by about 7% during the saline control infusion (35). The current data and previous studies (31, 32, 33, 34, 35) suggest that leptin secretion may be acutely regulated at least partially by the same pathways that regulate lipolysis in adipocytes.
It may be that the absolute levels of lipolysis in adipose tissue are more related to regulation of leptin levels than are changes in lipolysis. For example, if lipolysis and leptin secretion are regulated by the same pathways, it may be that there is a threshold level of signal necessary to both reduce lipolysis to a given level and stimulate leptin production. During fasting, when lipolysis levels are high in all subjects, the signals inhibiting lipolysis may be too low to have an effect on leptin production.
The plasma leptin concentration follows a well known, diurnal pattern, peaking a few hours after the evening meal and reaching its lowest level before refeeding after an overnight fast (36). The fasting plasma leptin concentration has consistently been shown to be highly related to body fat content in humans (2, 30). Citing the high fasting leptin levels in obese subjects, some investigators have suggested that resistance to the anorectic activity of leptin develops during obesity (5). However, it is also possible that an attenuated rise in plasma leptin throughout the day, not the baseline leptin level, underlies the apparent disruption of leptin signaling in obese subjects. For example, it has been shown that the leptinemic response to insulin was lowest in the most obese subjects, who were insulin resistant (8). Saad et al. (8) suggested that the rise in plasma leptin in response to insulin may be attenuated or absent in insulin-resistant individuals. Fasting leptin levels have been found to be nearly 4 times higher (30.5 vs. 8.2 ng/mL) in obese subjects than those in normal weight subjects (37), but when the subjects were fed, there was only a 2-fold difference (34.6 vs. 15.8). Leptin almost doubled from nadir to peak in normal weight subjects, compared to only about a 10% change in obese subjects (37). It may be possible that a reduced effect of insulin on the plasma leptin concentration in obese subjects is responsible for an inappropriate rise in plasma leptin during the day.
Although Saad et al. found an inverse relationship between
adiposity and the rise in plasma leptin during hyperinsulinemia (8), we
found no such relationship. It is likely that our low correlation
between body composition and changes in leptin is based on
methodological differences between our study and that of Saad et
al. Although we examined subjects with a broader range of body fat
than Saad et al. (7–69 kg fat in women vs.
5–48 kg in men), we did not infuse insulin for nearly as long (3
vs. 8 h). In addition, we compared leptin levels to
preinfusion levels, whereas Saad et al. compared leptin
levels at the end of insulin infusions to leptin levels after a saline
infusion. All of the current subjects had normal glucose tolerance, so
any relationship between adiposity and the change in leptin was
probably blunted by a restricted range of insulin resistance. However,
there was a relationship between insulin action on lipolysis
(i.e. absolute glycerol concentration) in abdominal sc
adipose tissue and the change in leptin during insulin infusions. The
results of the current study and others (8, 37) imply that
interventions, such as dietary restriction or exercise training, that
would increase insulin action would also augment the rise in leptin
throughout the day or during hyperinsulinemia. Support for this
conclusion was provided by the report of a 25% rise in the plasma
leptin level in obese women during a 3-h euglycemic-hyperinsulinemic
clamp after, but not before, a 6-day fast (3).
One research group has reported gender differences in the response of leptin to hyperinsulinemia (2). For example, after 3 h of insulin infusion, women, but not men, had a 20% increase in plasma leptin levels (2). It is unknown whether this is a real gender-based difference or whether the difference was based on sample variability, because other studies have found 1) no increase in plasma leptin during hyperinsulinemia in women (3, 4) 2) an increase in plasma leptin during hyperinsulinemia in men (8), or 3) similar increases in men and women (7). Although it is known that the relationship between body fat and fasting plasma leptin is steeper in women than in men (2), we believe that it is premature to suggest that women have a greater leptinemic response to insulin than men. Our subjects were not studied during a standardized time of the menstrual cycle, but without solid evidence for gender differences in the leptin response to insulin (above) we have no reason to believe that this would affect the results. Furthermore, in a previous study, we found that estrogen replacement in postmenopausal women had no effect on fasting leptin levels (38). However, we must acknowledge that our findings for women may not be generalizable to men.
Without a saline infusion control, we were only able to compare plasma leptin levels during insulin infusions to preinfusion levels. Over 3 h, leptin levels would be expected to fall about 20% in fasting subjects infused with saline (7). Thus, it is probable that even though leptin levels fell during insulin infusion in some subjects, leptin levels would have declined further during an insulin-free infusion. We would expect that correction of individual changes in leptin for saline control values would increase the statistical power to examine relationships between leptin and lipolysis, but we cannot be sure. The addition of a saline control in future studies would allow a more accurate examination of the relationship between lipolysis and leptin.
Our findings suggest that the changes in plasma leptin during hyperinsulinemia may be related to the relative lipolytic or lipogenic state of certain sc adipose tissue depots. It appears that mechanisms for regulation of leptin expression in sc abdominal adipose tissue may share some of the pathways that regulate lipolysis.
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Acknowledgments |
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Footnotes |
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2 Current address: Human Performance Laboratory, East Carolina
University, Greenville, North Carolina 27858.
Received February 12, 1999.
Revised May 26, 1999.
Accepted June 28, 1999.
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References |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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-2 adrenoceptors in regulating lipid
mobilization from human adipose tissue. J Clin Invest. 91:1997–2003.
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