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Nocturnal free fatty acids are uniquely elevated in the longitudinal development of diet-induced insulin resistance and hyperinsulinemia

Department of Physiology and Biophysics, Keck School of Medicine of the University of Southern California, Los Angeles, California

Submitted 7 December 2006 ; accepted in final form 29 January 2007

	ABSTRACT

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Obesity is strongly associated with hyperinsulinemia and insulin resistance, both primary risk factors for type 2 diabetes. It has been thought that increased fasting free fatty acids (FFA) may be responsible for the development of insulin resistance during obesity, causing an increase in plasma glucose levels, which would then signal for compensatory hyperinsulinemia. But when obesity is induced by fat feeding in the dog model, there is development of insulin resistance and a marked increase in fasting insulin despite constant fasting FFA and glucose. We examined the 24-h plasma profiles of FFA, glucose, and other hormones to observe any potential longitudinal postprandial or nocturnal alterations that could lead to both insulin resistance and compensatory hyperinsulinemia induced by a high-fat diet in eight normal dogs. We found that after 6 wk of a high-fat, hypercaloric diet, there was development of significant insulin resistance and hyperinsulinemia as well as accumulation of both subcutaneous and visceral fat without a change in either fasting glucose or postprandial glucose. Moreover, although there was no change in fasting FFA, there was a highly significant increase in the nocturnal levels of FFA that occurred as a result of fat feeding. Thus enhanced nocturnal FFA, but not glucose, may be responsible for development of insulin resistance and fasting hyperinsulinemia in the fat-fed dog model.

obesity; diurnal

	MATERIALS AND METHODS

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Animals

Male mongrel dogs (n = 8, 29.8 ± 1.4 kg) were housed in the Keck School of Medicine of University of Southern California (USC) vivarium under controlled kennel conditions (12:12-h light-dark cycle). Animals were accepted into the study following physical examination and a comprehensive blood panel. Chronic catheters were surgically implanted 7–10 days before the beginning of the study: one was inserted in the jugular vein and advanced to the right atrium for sampling of central venous blood, and a second catheter was inserted in the femoral vein and advanced to the vena cava for tracer, insulin, and somatostatin infusion. All catheters were led to the neck subcutaneously and exteriorized. Catheters were flushed with heparinized saline (10 U/ml) at least twice a week, and the exteriorization site was cleaned with hydrogen peroxide (4%). Dogs were accustomed to laboratory procedures and were used for experiments only if judged to be in good health as determined by visual observation, body temperature, and hematocrit. On the morning of each experiment, 19-gauge angiocatheters (Allegiance Healthcare, Ontario, CA) were inserted percutaneously into the saphenous vein for glucose infusion. The experimental protocol was approved by the USC Institutional Animal Care and Use Committee.

Diet

Dogs were fed a weight-maintaining standard diet of one can of Hill's Prescription Diet (10% carbohydrate, 9% protein, 8% fat, 0.3% fiber, and 73% moisture; Hill's Pet Nutrition, Topeka, KS) and 825 g of dry chow (36.6% carbohydrate, 26.4% protein, 14.7% fat, and 2.9% fiber; Wayne dog food; Allied Mills, Chicago, IL) for a period of 2–3 wk before any experiments were conducted to ensure weight stabilization. This standard diet consisted of 3,885 kcal/day: 37.9% from carbohydrates, 26.3% from protein, and 35.8% from fat. Following weight stabilization (week 0), dogs were maintained on a hypercaloric, high-fat diet for a period of 6 wk in which the standard diet was supplemented with 6 g/kg prediet body weight of cooked bacon grease supplied by the Keck School of Medicine cafeteria. This hypercaloric, high-fat diet consisted of a total of 5,392 kcal/day: 27.4% carbohydrates, 19% protein, and 53.6% fat. To acclimate the animals to the feeding protocol that would be used during the 24-h plasma profiling experiments (see below), throughout the study dogs were presented with their meal at 9:00 AM and given 1 h to eat, after which the meal was removed.

Magnetic Resonance Imaging

During weeks 0 and 6 of the high-fat diet, magnetic resonance imaging (MRI) scans were performed on the dogs as previously described (25). Thirty 1-cm axial abdominal images (T1 slices, TR 500, TE:14) were obtained using a General Electric 1.5-Tesla Horizon magnet (version 5.7 software). Of the 30 images obtained, 20 of these images were used for analysis of total trunk body fat, depending on the relative torso length of the animal. Images were analyzed using Scion Image (Windows 2000 version Beta 4.0.2; Scion, Frederick, MD), which quantifies fat tissue (pixel value 121–254) and other tissue (20–120) in each slice. Fat volume was calculated by dividing the number of pixels counted as fat by the ratio of the total number of pixels (256 x 256) and known area (34.9 x 34.9 cm) for a 1-cm image. Total trunk fat and tissue were estimated as the integrated fat or tissue across all 20 slices. Percent fat was calculated as the total trunk fat divided by the total trunk tissue. Omental fat was defined as fat within the peritoneal cavity in an 11-cm region of the thorax, using the slice at the level where the left renal artery branches from the abdominal aorta as a midpoint landmark. Percent omental fat was calculated as the omental fat divided by the total tissue area in these same slices.

Euglycemic Hyperinsulinemic Clamps

The euglycemic hyperinsulinemic clamps were performed as previously described (25) during weeks 0 and 6 of the high-fat diet. Animals were familiarized with the Pavlov sling at least 1 wk before the first experiment. At 7:00 AM on the day of the clamp, animals were brought to the laboratory and placed in the Pavlov sling. A 19-gauge angiocatheter was placed in a saphenous vein and secured. Approximately 30 min later (t = –120 min), a primed continuous infusion of high-performance liquid chromatography-purified [3-³H]glucose (25 µCi + 0.25 µCi/min infusion; DuPont-NEN, Boston, MA) was started. After tracer equilibration, basal samples were taken at –30, –20, –10, and –1 min. At time t = 0 min, a somatostatin infusion (1.0 µg·min^–1·kg^–1; Bachem California, Torrance, CA) was started to suppress endogenous insulin and glucagon secretion and was continued for the duration of the experiment. Porcine insulin was infused (0.75 mU·kg^–1·min^–1; Eli Lilly, Indianapolis, IN) into the femoral vein to attain hyperinsulinemia. Glucose was clamped at basal by a variable glucose infusion labeled with D-[3-³H]glucose (2.0 µCi/g) to minimize fluctuations in plasma specific activity. Blood samples were drawn from the jugular catheter every 10 min from –30 to 60 min, every 15 min from 60 to 120 min, and then every 10 min from 120 to 180 min.

Twenty-Four-Hour Plasma Profiling

To obtain a plasma profile over a 24-h period, we utilized the following protocol during both week 0 and week 6 of the study. The precise profiling protocol was identical at week 0 and week 6. At 5:00 AM on the day of each experiment, the animal was brought into the laboratory. The jugular vein catheter was exposed and secured at the neck to allow blood sampling. The animal was housed individually in a kennel (5 x 5 ft) for the duration of the experiment. Animals were unrestrained in the kennel throughout the experimental protocol. Starting at 6:00 AM, blood samples were drawn at 1-h intervals for the 24-h period until 6:00 AM the following morning. The dogs were presented with the standard diet meal (see Diet) at 9:00 AM and given until 10:00 AM to eat, whereupon the meal was removed. All uneaten food was weighed and recorded, and the exact same meal by weight and composition was given during the week 6 experiments to negate any acute effects of differences in food consumption on the experimental outcome. The animals consumed 40% of the meal during the week 0 experiment, and there was no significant difference in percent meal consumption during the week 6 experiment.

Sample Collection and Storage

Samples for assay of insulin, D-[3-³H]glucose, cortisol, and growth hormone were taken in tubes precoated with lithium fluoride and heparin (Brinkmann Instruments, Westbury, NY). The tubes for insulin and glucose also contained 50 µl of EDTA. Samples for determination of C-peptide and glucagon were collected in precoated lithium fluoride/heparin tubes containing 25 µl of EDTA and 50 µl of Trasylol (10,000 KIU/ml; Serological Proteins, Kankakee, IL). GLP-1 samples were taken in tubes precoated with lithium fluoride and heparin containing 50 µl of EDTA and 25 µl of dipeptidyl peptidase IV inhibitor (Linco Research, St. Charles, MO). Samples for FFA, glycerol, and triglyceride assays were taken in tubes with EDTA and paraoxon to inhibit lipase activity. All samples were immediately centrifuged, and plasma was separated and stored at –80°C for further analysis.

Assays

Glucose was measured with a YSI 2300 autoanalyzer (Yellow Springs Instruments, Yellow Springs, OH). FFA (NEFA C; Wako Pure Chemical Industries, Richmond, VA), glycerol, and triglycerides (serum triglyceride/glycerol determination kit; Sigma Chemical, St. Louis, MO) were measured using colorimetric methods, utilizing commercially available kits. Insulin was measured using an ELISA originally developed for human serum or plasma (Linco Research) and adapted for dog plasma by using a dog standard kindly provided by Novo-Nordisk. The method is based on two murine monoclonal antibodies that bind to different epitopes of insulin but that do not bind to proinsulin. Cortisol, glucagon, growth hormone, and canine C-peptide were measured using a radioimmunoassay kit, and active GLP-1 was measured using ELISA (cortisol RIA kit from Diagnostic Products, Los Angeles, CA; all other kits from Linco Research).

Samples for [³H]glucose tracer assay were deproteinized using barium hydroxide and zinc sulfate. The supernatants were then evaporated in a vacuum, reconstituted in water, and counted in Ready Safe scintillation fluid (Beckman liquid scintillation fluid; Beckman Instruments, Fullerton, CA). Tracer infusates were processed identically to plasma samples.

Calculations

The time courses of endogenous glucose production and glucose disappearance during the euglycemic hyperinsulinemic clamp were calculated using Steele's model with a labeled glucose infusion as previously described (17). Derivatives of all time course data were calculated with OOPSEG (10). Basal was defined as the average of four samples taken every 10 min from t = –30 to 0 min, and steady state was defined as the average of four samples taken from t = 150 to 180 min. Insulin sensitivity (S_I) was calculated using the equation

where G_inf is the difference in glucose infusion rate at steady state from basal, I is the difference in plasma insulin at steady state from basal, and G is the steady-state plasma glucose concentration.

During the 24-h protocol, the total integrated area under the curve (AUC) was calculated using the trapezoidal rule. Insulin secretion rates were estimated using measured C-peptide levels and a two-compartment model for C-peptide distribution, as described previously (33). C-peptide sampling occurs from the central compartment, from which it diffuses into a peripheral compartment. Estimates of the kinetic parameters associated with our analysis were acquired from previously published data for canine C-peptide distribution kinetics (33, 35, 36).

Statistical Analyses

All experimental data are expressed as means ± SE. Repeated-measures ANOVA with Bonferroni posttests was used to compare all time course data before and after fat feeding. Paired Student's t-tests were used to identify the significantly different time point pairs and to compare all fasting metabolic parameters, 24-h averages, and 24-h AUC between weeks 0 and 6. The t-tests were performed using Microsoft Excel XP, and all ANOVAs were performed using GraphPad InStat 3.0 (GraphPad Software, San Diego, CA).

	RESULTS

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Body Composition

As expected, body weight increased in response to the increased caloric content starting at week 1 (Fig. 1A). By week 6, animals had gained an average of 3 kg (from 27.6 ± 1.4 to 30.3 ± 2.0 kg, P < 0.05). Weight increase was reflected in a substantial increase in body fat as assessed by MRI. Total trunk fat increased from 1,081 ± 133 cm³ at week 0 to 1,907 ± 304 cm³ at week 6 (P = 0.006), an approximate 76% increase in total trunk adiposity over the 6-wk period.

View larger version (36K): [in this window] [in a new window]   Fig. 1. A: average body weight. Consistent with the increase in body fat, body weight increased in response to the increased caloric content starting at week 1 (dashed line represents the average of individual body weights for weeks –2 to 0). The increase in body weight continued throughout the fat-feeding regimen such that body weight had increased by 2.9 kg by week 6 of the study. *P < 0.05 vs. week 0, paired t-test. B: omental (open bars) and subcutaneous fat (hatched bars) calculated as cm3 from the sum of 11 axial slices. **P = 0.006, paired t-test. C: axial MRI images from 4 representative dogs at weeks 0 and 6 of fat feeding. Each image was taken at the level of the left renal artery branching from the abdominal aorta, which was used as a midpoint landmark for the 11-cm3 region used in quantifying omental and subcutaneous fat. Images are inverse T1 weighted; adipose tissue appears as yellow, and all other tissue appears red. By week 6 of the study, adipose tissue showed accumulation in both the omental and subcutaneous compartments.

Fasting Metabolic Parameters

Despite an impressive 76% increase in total trunk fat (see Body Composition) and increased body weight, neither fasting free fatty acids (week 0: 0.50 ± 0.06 vs. week 6: 0.58 ± 0.04 mM; P = 0.24) nor fasting glucose (week 0: 99 ± 2 vs. week 6: 95 ± 1 mg/dl; P = 0.12) was changed by the high-fat diet. This lack of a change in fasting levels of FFA or glucose recapitulates previous results with a lesser fat diet (25, 31). However, despite unchanged fasting levels of FFA, there was a significant increase in insulin resistance after 6 wk of a hypercaloric, high-fat diet such that whole body insulin sensitivity decreased by 30% (week 0: 7.7 ± 0.8 vs. week 6: 5.5 ± 0.3 x 10^–4 dl·pM^–1·mg^–1·kg^–1; P = 0.008) in association with the increased adiposity. This decrease in insulin sensitivity was due to a significant reduction in insulin's ability to stimulate glucose uptake and a tendency for a decrease in insulin's ability to suppress endogenous glucose output (Fig. 2, A and B). The suppression of FFA during the hyperinsulinemic conditions of the glucose clamp was profound but similar before and after fat feeding (Fig. 2C).

View larger version (16K): [in this window] [in a new window]   Fig. 2. Time course data of glucose uptake (A), glucose production (B), and free fatty acids (FFA; C) during the clamp at week 0 () and week 6 () of the high-fat diet. *P < 0.05 vs. week 0, paired t-test.

Twenty-Four-Hour Plasma Profile

Insulin. We considered the possibility that increases in FFA, glucose, or other related factors at alternative times of the day might have played a role in the development of insulin resistance and compensatory hyperinsulinemia. The insulin response to a meal (10:00 AM to 6:00 PM) was profoundly enhanced after fat feeding (Fig. 3A). The total AUC for plasma insulin for the 8-h period following the 9:00 AM meal increased by 223 ± 44% after 6 wk of fat feeding (P = 0.01). This increased insulin response following the meal resulted in a significant increase in both the average insulin concentration and the total AUC over the 24-h period (Table 1) such that average insulin was increased by 70.1 ± 25.8% (P < 0.05) and the total AUC was increased by 69.8 ± 26.1% (P < 0.05) compared with week 0.

View larger version (23K): [in this window] [in a new window]   Fig. 3. Twenty-four-hour plasma profile of insulin (A) and C-peptide (B) at week 0 () and week 6 (). C: insulin secretion as calculated by C-peptide deconvolution analysis at week 0 (solid line) and week 6 (hatched line). Vertical dashed lines represent times of meal presentation and meal removal. *P < 0.05 vs. week 0, paired t-test.

Insulin secretion rates, calculated from deconvolution of plasma C-peptide concentrations, were unchanged after 6 wk of high-fat diet (Fig. 3C), suggesting that a decrease in insulin clearance was a major contributor to the compensatory increase in plasma insulin after fat feeding. This finding confirms previous results for the fat-fed dog model, which showed that decreases in insulin clearance as well as increases in insulin secretion can act to contribute to hyperinsulinemia (25, 31).

Glucose. Recapitulating fasting glucose levels, there was absolutely no discernible increase in glucose throughout the 24-h observation period after a 6-wk high-fat diet (Fig. 4A). The average glucose for the 24-h period before fat feeding was unchanged, as was the total AUC (Table 1), indicating that increased glucose could not be a signal for either insulin resistance or compensatory hyperinsulinemia in the fat-fed dog model.

View larger version (20K): [in this window] [in a new window]   Fig. 4. Twenty-four hour plasma profiles at week 0 () and week 6 (). A: plasma glucose concentrations did not show a significant increase from basal throughout the 24-h period, and there was no significant difference in plasma glucose levels at any time point before and after fat feeding. Plasma FFA (B) and plasma glycerol levels (C) over the 24-h period showed a similar increase at week 6 of the study that became significant at 5:00 PM and continued until the morning, whereas triglyceride levels (D) were unchanged by fat feeding. Vertical dashed lines represent times of meal presentation and meal removal. *P < 0.05 vs. week 0, paired t-test.

It was of interest to ask whether this return to the control diet only on the day of the experiment was responsible for the overall increase in FFA observed during the late postprandial and nocturnal periods. To discount this possibility, we presented a fat-supplemented meal to a separate set of 6-wk fat-fed dogs (n = 9) on the day of nocturnal measurements. To confirm the phenomenon of increased nocturnal FFA, we assessed FFA levels from 6:00 to 8:00 PM (when the rise begins) and from 2:00 to 4:00 AM (when FFA are maximal). There was a very similar increase in FFA levels after 6 wk of fat feeding whether or not we returned to the control diet for 1 day (Fig. 5). In the present study, FFA levels between 6:00 and 8:00 PM increased from 0.37 ± 0.07 mM at week 0 to 0.59 ± 0.08 mM at week 6. Similarly, when dogs were fed the fat-supplemented meal on the day of the week 6 experiments, FFA levels also increased from 0.25 ± 0.04 mM at week 0 to 0.59 ± 0.03 at week 6 (P < 0.001). The changes in overnight FFA also showed remarkably similar results. In the present study, when dogs were fed the control diet on the day of the week 6 experiments, average FFA levels from 2:00 to 4:00 AM increased from 0.45 ± 0.04 mM at week 0 to 0.80 ± 0.05 mM at week 6. When dogs were fed the fat-supplemented meal on the day of the week 6 experiments, FFA levels were still increased during this time frame from 2:00 to 4:00 AM (week 0: 0.47 ± 0.07 vs. week 6: 0.76 ± 0.04 mM; P < 0.001). It was clearly not the switch to the control diet for a single day that was responsible for the large increase in FFA at night.

View larger version (22K): [in this window] [in a new window]   Fig. 5. Average plasma FFA concentrations at week 0 (open bars) and week 6 (hatched bars) at basal, between 6:00 and 8:00 PM, and between 2:00 and 4:00 AM, when the control meal was given on the day of the experiment (A) and when the fat-supplemented meal was given on the day of the experiment (B). *P < 0.05 vs. week 0, paired t-test.

Cortisol, growth hormone, and glucagon. Cortisol, growth hormone, or glucagon did not exhibited a change in overall plasma concentrations over the 24-h period after 6 wk of fat feeding. (Fig. 6, A–C). In addition, the secretory pattern of growth hormone as derived from the 15-min sampling period between 12:00 and 3:00 AM was unchanged (data not shown).

View larger version (21K): [in this window] [in a new window]   Fig. 6. Twenty-four-hour plasma profiles of cortisol (A), growth hormone (B), glucagon (C), and glucagon-like peptide-1 (GLP-1; D) at week 0 () and week 6 (). Vertical dashed lines represent times of meal presentation and meal removal. *P < 0.05 vs. week 0, paired t-test.

	DISCUSSION

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Of the many possible factors influencing development of the metabolic syndrome, FFA have been implicated as an important component in the development of insulin resistance, particularly in obesity (8, 14). Elevated levels of fasting FFA levels are often thought of as a characteristic marker for insulin resistance in obesity, and it has been suggested that these FFA may be responsible for reduced insulin action at both skeletal muscle and liver. Ensuing insulin resistance would then result in elevated glucose, which signals for a compensatory increase in circulating insulin concentrations. Thus it has long been assumed that modest fasting and/or postprandial plasma glucose is responsible for increases in plasma insulin in the insulin-resistant state. As an example, in Pima Indians, a population highly at risk for diabetes, glucose increases longitudinally with insulin resistance (46). In contrast, other populations at risk for diabetes are able to maintain normal glucose levels despite insulin resistance (16, 22), indicating that elevated fasting glucose may not be the primary signal responsible for hyperinsulinemia with insulin resistance but, rather, that glucose only becomes elevated when hyperinsulinemia is unable to fully compensate for insulin resistance. And although fasting levels of FFA can be elevated in obesity and insulin resistance, this is not always the case. Studies conducted in humans as well as animals have found relatively normal levels of both FFA and glucose at fasting despite obesity, insulin resistance, and hyperinsulinemia (16, 20, 22, 38). The latter data lead to the question of whether these metabolites are responsible for insulin resistance and hyperinsulinemia during obesity. Studies conducted in our own laboratory (25, 31) using the fat-fed dog model have shown significant insulin resistance and increases of 90–150% in basal insulin with no measurable change in fasting FFA or glucose. In the present study, we sought to examine whether there were any alterations in FFA or glucose over the 24-h day as well as other factors, including cortisol, growth hormone, and GLP-1, that could potentially serve as contributors to the development of insulin resistance and compensatory hyperinsulinemia.

In the current study, eight normal dogs were fed a hypercaloric, high-fat diet with an increase in fat content of 20% for a period of 6 wk. There was significant accumulation of total trunk body fat due to increases in both visceral and subcutaneous fat depots with a concomitant increase in body weight. The dogs exhibited a decrease in insulin sensitivity and developed fasting hyperinsulinemia. However, despite increased adiposity, insulin resistance, and fasting hyperinsulinemia, the animals did not develop an elevation in fasting FFA or fasting hyperglycemia. Neither fasting FFA nor glucose appears to be the cause for development of insulin resistance and compensatory hyperinsulinemia in this obese dog model. More impressive was virtually exact reproducibility of the 24-h glucose pattern in lean vs. fat-fed animals in contrast to the elevation in 24-h insulin profile after fat feeding. Similarly, although circadian rhythms were present, there were no changes in the 24-h profiles of growth hormone, cortisol, or even GLP-1 before compared with after fat feeding. In contrast to all other variables measured, there was a profound and highly significant increase in the 24-h profile of FFA after fat diet, due almost entirely to the nocturnal rise in FFA, which began at 5:00 PM and continued throughout the night into the early morning. These results nominate nocturnal elevation of FFA to be potentially responsible for development of insulin resistance and subsequent increase in plasma insulin availability during diet-induced obesity in the dog model.

There has been much evidence supporting the existence of differences between the diurnal metabolic profiles of obese and nonobese individuals as well as the discrepancy between the diurnal patterns of healthy vs. insulin-resistant subjects. It has been reported that a diurnal variation in FFA exists (32) as well as a fall in hepatic glucose output which occurs during sleep that is highly synchronized with a decrease in FFA levels in normal weight individuals (13). It has also been shown that the rate of appearance of FFA at night is increased in type 2 diabetics (30) as well as an overnight elevation in plasma FFA that is correlated with the overnight increase in hepatic glucose output in type 2 diabetes (44). In addition, it has been demonstrated that a reduction in overnight FFA levels by acute pharmacological blockade of lipolysis results in a reduction of insulin levels (1). This suggests that the elevated nocturnal FFA observed with fat feeding may be pivotal in the development of hepatic insulin resistance in addition to being a potential signal for insulin upregulation, even when fasting FFA are unchanged.

The data of the current study linking elevated nocturnal FFA with insulin resistance and compensatory hyperinsulinemia are indeed correlative in nature. Nevertheless, the suggestion that FFA themselves may play an important role in the metabolic syndrome is supported by a wealth of data showing that chronic infusion of FFA decreases insulin sensitivity and that FFA can upregulate insulin levels by both increasing insulin secretion and decreasing hepatic insulin extraction. Acute elevation of FFA by intravenous infusion enhanced both basal and glucose-stimulated insulin secretion, whereas long-term infusion of FFA has been shown to attenuate pancreatic insulin secretion (9, 12). Others have shown that an acute elevation of FFA can cause hyperinsulinemia without a change in insulin secretion (5). In addition, FFA impair the hepatic clearance of insulin, which would increase the fraction of secreted insulin delivered to the systemic circulation (42, 47). In this study, there was development of hyperinsulinemia without a similar change in C-peptide concentrations, suggesting that hepatic clearance of insulin was indeed responsible for basal hyperinsulinemia and the upregulated insulin levels in this model of insulin resistance. Although the majority of the difference in plasma insulin levels occurs when FFA are in fact similar, it is possible that the chronic effect of elevated FFA at night may impair hepatic insulin extraction by means of increased triglyceride deposition in the liver, resulting in subsequent hyperinsulinemia (43). Our group has shown previously that dogs fed a diet with an elevated fat content exhibited a 45% increase in liver triglyceride (24, 25). In addition, because we did not measure the time course of changes in nocturnal FFA during fat feeding, we cannot discount the possibility that potential earlier elevations in nocturnal FFA led to temporal augmentation of insulin secretion by direct stimulation of the pancreatic -cells. Previous studies completed in our laboratory have shown that in dogs fed a lesser fat diet, compensation for insulin resistance occurs in a longitudinal manner that includes both enhanced insulin secretion and decreased insulin clearance (31), suggesting that there may be a temporal effect of FFA in mediating hyperinsulinemia via insulin secretion vs. insulin clearance. Studies examining the longitudinal time course for the development of elevated nocturnal FFA and insulin resistance during high-fat feeding in conjunction with the direct examination of insulin secretion vs. hepatic insulin extraction are currently under investigation in our laboratory.

Although the present studies were not designed to determine the origin of the highly significant increase in the nocturnal levels of FFA, we can speculate as to potential mechanisms. Lipolysis of endogenous FFA stored as triglycerides (mainly adipose tissue) is a major determinant of increased FFA in plasma (4). However, in the present study, likely lipolytic candidates such as growth hormone and cortisol as well as glucagon were ruled out. It is also possible that the sensitivity to lipolytic factors is altered in the obese, insulin-resistant animal. So-called "adipokines," which have been implicated in the development of insulin resistance and hyperinsulinemia (34), may also be responsible for increased lipolytic activity during obesity. In particular, IL-6 has been shown to be elevated in obesity and has also been shown to display a diurnal pattern of secretion, with IL-6 levels peaking at 1:00 AM or later (39). Moreover, it has been demonstrated that IL-6 can elicit lipolytic effects in adipose tissue both in vitro and in vivo (27, 28), implying that IL-6 may be a mechanism involved in modulating FFA levels. Low adiponectin levels have also been associated with insulin resistance and obesity (3) and have been implicated as a possible regulatory factor involved in FFA release (45). In addition, adiponectin was recently shown to exhibit ultradian pulsatility as well as a diurnal variation that declines significantly at night (19) and is absent in obesity and insulin resistance (11), suggesting that decreased levels of adiponectin may also be involved in mediating the nocturnal elevation of FFA observed in this study. Another possible mechanism for increased FFA is catecholamine-stimulated lipolysis. Efferent signals from the brain via sympathetic nervous system innervation of adipose tissue have been shown to be of great significance in the mobilization of FFA stores from adipose tissue (6). Moreover, Landsberg (26) found increases in sympathetic activity in obesity in humans, and it has been suggested that sympathetic activity is increased at night in at-risk patients (33a). Our group (23) recently presented evidence for cyclic lipolysis that can be blocked by the 3 receptor antagonist bupranolol, indicating an important role for the central nervous system in regulating the provision of lipid fuels. It is tempting to suggest that the central nervous system plays an intermediate role in the development of insulin resistance and subsequent upregulation of plasma insulin in response to fat feeding by modulating sympathetic nervous system control of FFA levels at night. In addition to increased lipolytic release of intracellular FFA stores, a proportion of fatty acids generated by intravascular triglyceride hydrolysis also contributes to plasma FFA (18, 29), and this can be increased in obesity (37). In this study, although FFA levels showed a similar suppression following the meal before and after fat feeding, FFA levels at week 6 began to increase during the late postprandial period despite unchanged triglyceride levels. This late postprandial rise in FFA levels suggests that there may have been an increase in the proportion of fatty acids released from triglyceride-rich lipoproteins that further contributed to the nocturnal increase in FFA. Further studies utilizing tracer techniques are needed to address this issue.

It is also possible that the sensitivity to lipolytic factors is altered in the obese, insulin-resistant animal. Although lipolytic hormones such as growth hormone, cortisol, and glucagon were unchanged by fat feeding during the 24-h experiments, we observed elevated FFA levels the morning following the experiment (i.e., 6:00 AM on day 2). Although it is possible that the experimental paradigm per se may have contributed to the elevated FFA observed on day 2 of the experiment, it is of note that this effect was only observed after 6 wk of fat feeding.

It is generally agreed that the defect in insulin sensitivity as well as a defect in pancreatic function in combination are responsible for the chronic hyperglycemia seen in diabetes. To effectively prevent the onset of diabetes, it is crucial to understand the underlying mechanisms responsible for development of insulin resistance and subsequent hyperinsulinemic compensation in normal individuals and how these mechanisms might fail in those at risk for diabetes. Our results indicate that during obesity induced by a high-fat diet, there is development of insulin resistance and a compensatory increase in both fasting and postprandial insulin levels, most likely due to a decrease in hepatic insulin clearance. The present study appears to rule out several putative factors known to be involved in the development of insulin resistance and insulin upregulation, including glucose itself, cortisol, growth hormone, and glucagon. FFA remain as a candidate. We have not proven in the present study that FFA are the signal, because we did not examine the effects of suppressing lipolysis at night. However, this present study suggests that in obesity, there is significant development of insulin resistance and hyperinsulinemia that can occur even when basal levels of FFA or glucose are unchanged and that nocturnal FFA levels may be a key factor in the development of the metabolic syndrome.

	GRANTS

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This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-27619 and DK-29867 (to R. N. Bergman). K. J. Catalano is supported by National Institutes of Aging Predoctoral Training Grant T32 AG-00093.

	ACKNOWLEDGMENTS

 
We thank Edward Zuniga for assistance with animal care and handling. We also thank Antonios Panteleon for help with C-peptide deconvolution analysis.

	FOOTNOTES

 

Address for reprint requests and other correspondence: R. N. Bergman, Dept. of Physiology and Biophysics, Keck School of Medicine of the Univ. of Southern California, 1333 San Pablo St. MMR 626, Los Angeles, CA 90033 (e-mail: rbergman{at}usc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

	REFERENCES

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 

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