1 Department of Pediatrics, University of Colorado Health Sciences Center, Denver 80262; 2 Department of Food Science and Human Nutrition, Colorado State University, Fort Collins, Colorado 80525; and 3 Department of Exercise Science and Physical Education, Arizona State University, Tempe, Arizona 85287
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ABSTRACT |
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Defects in fat
metabolism may contribute to the development of obesity, but what these
defects are and where they occur in the feeding/fasting cycle are
unknown. In the present study, basal fat metabolism was characterized
using a high-fat diet (HFD)-induced model of obesity development. Male
rats consumed a HFD (45% fat, 35% carbohydrate) ad libitum for either
1 or 5 wk (HFD1 or HFD5). After 1 wk on the HFD, rats were separated on
the basis of body weight gain into obesity-prone (OP, 48 g) or
obesity-resistant (OR,
40 g) groups. Twenty-four-hour-fasted rats
were studied either at this time (OP1, OR1) or after 5 wk (OP5, OR5).
Fat pad weight (sum of epididymal, retroperitoneal, and mesenteric fat pads) at HFD1 was 26% greater and at HFD5 was 43% greater
(P
0.05) in OP vs. OR. Free fatty acid rates of appearance
(FFA Ra) and oxidation were not significantly different
between OP and OR at 1 or 5 wk. Glycerol Ra, when expressed
in absolute terms (µmol/min), increased from 1 to 5 wk of HFD feeding
in both OP and OR, but significantly so only in OP. Likewise, increased
rates of intracellular FFA cycling [estimated as (3 × glycerol
Ra)
FFA Ra] were observed in both OP
and OR rats from 1 to 5 wk of HFD feeding, but significantly so in OP
rats only. When expressed relative to fat cell volume (µmol · pl
1 · min
1),
neither lipolysis nor intracellular cycling was significantly different
between OP and OR, regardless of time on HFD. These data suggest that
1) if low rates of fat oxidation contribute to obesity
development in OP rats, the contribution does not occur at times when
fat oxidation is at or near maximum rates (i.e., 24-h fasted
conditions), and 2) between 1 and 5 wk of HFD feeding, basal
lipolysis and reesterification may work to expand fat cell volume and
increase fat pad weight in both OP and OR rats, although more so in OP rats.
fatty acid metabolism; reesterification; triglyceride cycling; high-fat diet
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INTRODUCTION |
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OBESITY IS CHARACTERIZED by increased fat stores, where energy intake exceeds energy expenditure. In rodents, a high-fat diet (HFD) promotes obesity through hyperphagia (9, 17, 18). However, even under conditions of isocaloric feeding, there exists a direct relationship between percent body fat and fat content of the diet (2). These data suggest that a HFD promotes nutrient partitioning in a way permissive to expansion of fat pad stores. Importantly, HFD-induced obesity occurs in some but not all rats. For example, after 1 wk of ad libitum HFD feeding, rats prone to obesity development (OP, top tertile of weight gain) eat more, gain more weight, and have a higher percent body fat than do rats resistant to obesity development (OR, lowest tertile of weight gain). By 5 wk of HFD, weekly weight gain in OP rats is only slightly greater than that observed in OR rats. Thus this model permits comparisons between rats that are developing obesity (i.e., OP rats after 1 wk of HFD) and rats in which the obese state is established (OP rats after 5 wks of HFD).
Data from several studies previously reported from our
laboratory (10, 20) suggest that OP rats either respond
differently to early exposure to the HFD relative to OR rats, or that,
upon beginning the HFD feeding period, OP rats are metabolically
distinct from OR rats. For instance, both lipoprotein lipase mRNA and
activity were significantly greater in adipose tissue but were lower in skeletal muscle of OP rats after 1 and 2 wk of HFD feeding, but not
after 5 wk of HFD (20). Likewise, skeletal muscle enzyme profiles of OP rats (e.g., the ratio between the maximal activities of
phosphofructokinase to those of -hydroxyacyl-CoA dehydrogenase) indicated an increased capacity for carbohydrate over fat oxidation at
both 1 and 2 wk of HFD feeding, but not at 5 wk (10).
These findings imply that, with early exposure to the HFD, OP rats
would oxidize less fat under conditions of maximal fat oxidation.
Although comparisons of 24-h respiratory quotient (RQ) have been made
between OP and OR at 5 wk of HFD feeding (4), nothing is
known regarding the ability of OP rats to oxidize fat early in their
exposure to the HFD. We were therefore interested to learn whether, as implied by their enzymatic profiles, OP rats were characterized by a
decreased capacity to oxidize fat under conditions of near-maximum fat
oxidation (i.e., after a 24-h fast) at 1 wk but not at 5 wk of HFD.
Both fat pad weight and percent body fat are significantly greater in OP rats by 1 wk of HFD feeding. Fat stores expand when nutrient uptake by adipocytes exceeds their release. If lipolysis is lower relative to nutrient uptake, or if rates of reesterification exceed nutrient release, fat pad mass will increase. The mechanism by which an animal on a HFD partitions nutrients toward fat storage is presently unknown, but it may relate to impairments in insulin action. Lipolysis is very sensitive to the antilipolytic effects of insulin (14), so that even under basal conditions, insulin concentrations will have a significant effect on lipolysis. Some data in OP rats suggest that they are relatively more insulin resistant than are OR rats with longer-term HFD feeding, but not at 1 wk of HFD (4, 10, 20, 22). In addition to lipolysis, the partitioning of fatty acids between reesterification and oxidation will influence fat accumulation and therefore obesity development. As such, a second aim of the present investigation was to characterize rates of lipolysis and reesterification in rats prone and resistant to HFD-induced obesity under the same conditions in which fat oxidation was measured. In conjunction with these estimates, catecholamines and corticosterone concentrations were measured, because these two hormones can regulate fat metabolism.
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METHODS |
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Experimental Animals
Male Crl(WI)BR rats (7 wk of age) were housed at the University of Colorado Health Sciences Center Animal Resource Center for either 3 or 7 wk. Rats weighed 135-180 g on arrival. Nine cohorts of between 16 and 50 animals per cohort were studied (total number of rats studied was 92). Rats were individually caged under controlled conditions (12:12-h light-dark cycle).Diet Protocol
On arrival, animals were provided ad libitum access to a semi-purified low-fat diet (LFD; 12% fat, 68% carbohydrate, 20% protein; Research Diets, New Brunswick, NJ; Table 1) for 2 wk (baseline period). After the baseline period, rats either remained on the LFD or were switched to an HFD (45% fat, 35% carbohydrate, 20% protein; Research Diets; Table 1). Rats on the HFD were separated on the basis of body weight gain after 1 wk of ad libitum access into either OP (body weight gain
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Animal Preparation
After either 1 or 5 wk of HFD feeding and several days before an animal's day of study, catheters were implanted into the jugular vein (tracer infusions) and carotid artery (blood sampling), with animals under general anesthesia as previously described (23). Rats were allowed to recoverExperimental Protocol
At 0900 on the morning before the day of study, body weight and the previous day's food intake were measured, and all food was removed. Studies commenced between 0900 and 1200 of the following morning (fasted conditions). Because fuel stores present within a tissue can potentially impact upon the substrate mixture used to meet energy needs, we chose to fast rats for 24 h, because this length of fast reduces liver glycogen to minimal concentrations and equates skeletal muscle glycogen in OP and OR rats (21). On the morning of the study day, extensions were placed onto exteriorized catheters to allow easy access without disturbing animals. Before the tracer infusion was initiated, an arterial blood sample was drawn, rats were placed into a metabolic chamber (airflow = 1.0 l/min), and a preinfusion respiratory gas sample was collected. An arterial infusion of normal saline (no heparin) was then initiated to maintain arterial line patency.[1-14C]palmitate (0.05 µCi/min) and
[2-3H]glycerol (0.10 µCi/min; NEN Life Science
Products, Boston, MA), resuspended in 22% bovine serum albumin (Sigma,
St. Louis, MO), were used to estimate plasma free fatty acid (FFA)
turnover, plasma FFA oxidation, and whole body lipolysis. Data in a
subset of rats (those fed either chow or HFD, n = 6),
which compared the method described here [liquid scintillation
counting (LSC) method] to estimate FFA Ra with that in
which plasma palmitate was directly determined (HPLC method), indicated
that the two methods were comparable (6.0 ± 1.7 mg · kg1 · min
1 for the LSC
method vs. 6.7 ± 1.8 mg · kg
1 · min
1 for the
HPLC method; r2 = 0.995). Tracers were
infused via the jugular vein catheter for 140 min. At 120, 130, and 140 min of tracer infusion, arterial blood samples were collected
concurrently with respiratory gases and expired
14CO2 samples. At t = 140, pentobarbital sodium (50 mg/kg) was administered, and epididymal,
retroperitoneal, and mesenteric fat pads were taken and immediately
weighed. Portions of each pad were placed in Krebs Ringer
phosphate (KRP) buffer for determination of fat cell size and
number, which were determined within 6 h of tissue removal. Urine
was removed from the bladder by use of a tuberculin syringe and
combined with urine collected throughout the infusion period, and total
urine volume was recorded.
Determination of Acetate Correction Factor
In separate studies, [1-14C]acetate was used to estimate an acetate correction factor for each group (necessary for calculation of plasma FFA oxidation; Ref. 25). [1-14C]acetate reconstituted with 0.9% normal saline was infused into the jugular vein at a rate of 0.05 µCi/min for 140 min. Rats were maintained in metabolic chambers (airflow = 1.0 l/min) throughout the period of infusion. Expired CO2 was collected immediately before initiation of the constant infusion and at 120, 130, and 140 min after the start of infusion. No blood samples were taken in these studies.Analytical Procedures
Blood and urine.
Blood samples were drawn into heparinized syringes. For glycerol, 50 µl of whole blood were deproteinized (26). The
supernatant was used to determine glycerol concentrations
(30) and [3H]glycerol, after having been
taken to dryness to drive off any 3H2O. The
remaining blood was immediately centrifuged, and the plasma was used to
determine circulating substrates, hormones, and 14C-labeled
FFA. Plasma glucose concentration was determined using a Beckman
glucose analyzer (Fullerton, CA). FFA concentration was determined
spectrophotometrically (Wako NEFA-C kit, Richmond, VA). Insulin (Linco
Research, St. Charles, MO) and corticosterone (Coat-a-Count,
Diagnostics Products, Los Angeles, CA) were determined by
radioimmunoassay. Catecholamines were stored at 70°C with reduced
glutathione (5 mM) until analysis. Analysis was performed by HPLC as
previously described (12). Urinary nitrogen corrected for
ammonia was determined spectrophotometrically (Sigma kit no. 640-A).
Determination of 14C-FFAs was made from extracts of plasma
according to Dole (7), as modified by Trout
(28). Briefly, an extraction mixture (1.0 ml of 40:10:1
isopropyl alcohol-heptane-1.0 N H2SO4) was
added to 200 µl of plasma and vortexed. Heptane (600 µl) and
distilled, deionized H2O (400 µl) were added, and each
sample was vortexed for
2 min. The resulting top layer was removed to
a glass culture tube, and an equal volume of 0.05%
H2SO4 was added. Each sample was vortexed for 5 min and centrifuged at 1,350 rpm for 5 min, and the top layer was again
removed and transferred to a glass scintillation vial. Titration
mixture (100 µl; 0.01% thymol blue in 95% ethanol) was added, and
the solution was titrated with 0.018 N NaOH (made fresh daily) to
titration end point. The resulting extract was evaporated to dryness
with a low-pressure stream of N2 gas and was reconstituted
with 0.9% saline. Scintillation cocktail was added, the samples were
allowed to sit overnight, and radioactivity was counted on the
following day (LSC6500, Beckman, Fullerton, CA). Plasma standards
spiked with a known quantity of [1-14C]palmitate were
extracted in duplicate along with samples, and counts were compared
with those attained from an aliquot of nonextracted spiked plasma to
determine percent recovery. Percent recovery of radioactivity from
plasma standards averaged 87.2%. Duplicity between labeled plasma
standards (relative percent difference) never exceeded 7.2%, and the
day-to-day coefficient of variation for percent recovery was 6.0%.
Respiratory gases.
Respiratory gases were collected into modified Douglas bags for the
measurement of O2 consumption
(O2) and CO2 production (
CO2) between 110 and 113, 120 and 123, and 130 and 133 min. Samples were drawn at a rate of 1.0 l/min through
O2 and CO2 analyzers (Ametek, Pittsburg, PA),
and the deviation from atmospheric air was recorded.
Fat cell volume and number. Fat cell size and number were determined by methods previously described (6). Fat cell diameter was determined under a microscope after cells had been digested for 10 min in collagenase (2 mg of collagenase/ml KRP buffer) at 37°C (9). Methylene blue was used to stain the cell membrane.
Calculations
Glycerol Ra and FFA turnover.
Glycerol Ra and FFA Ra were calculated using
the steady-state equation of Steele (27). FFA
Ra was calculated using [14C]palmitate
concentration (dpm/ml) relative to total FFA concentration (µmol/ml)
as follows
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Whole body fat oxidation by indirect calorimetry.
After correction to STPD (standard temperature and pressure, dry),
O2 and
CO2 were calculated using a rolling
average over the three time points collected to give a single value for each. The mean value for both
O2 and
CO2 (ml/min) was extrapolated to 140 min
to determine total
O2 and
CO2 over the course of the experiment.
Energy expenditure and substrate oxidation after correction for protein
oxidation (6.25 × g of urinary nitrogen; see Ref. 16) were
calculated from
O2, and the nonprotein
RQ (npRQ) was calculated using equations previously described (8, 16).
Fat cell volume and number.
Fat cell volume was calculated using the relationship between diameter
and volume: 4.19 (diameter/2)3. The number of fat cells per
gram of tissue was calculated using cell volume and the density of
lipid
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Acetate correction factor. An acetate correction factor was calculated in a subset of rats (n = 4/group) as the Ra of expired 14CO2 divided by the rate of infusion of [1-14C]acetate. The value used for expired 14CO2 (dpm/min) was a rolling average of values collected at 120, 130, and 140 min of the [1-14C]acetate infusion.
Plasma FFA oxidation.
The rate of plasma palmitate oxidation was calculated as follows
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Intracellular and extracellular triglyceride cycling.
Triglyceride cycling, in which FFAs are hydrolyzed from glycerol and
then reesterified to triglyceride, can occur in vivo (3,
31). Triglyceride cycling can be divided into intracellular and
extracellular cycling. Intracellular cycling, defined as hydrolysis of
FFAs from glycerol that are then reesterified within the adipocyte without exiting the cell, was calculated as
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Data analyses.
One-way ANOVA was used to detect differences across groups. When
necessary, multiple comparison tests were made using the method of
Tukey (29). Two-way ANOVA was used to determine the significance of any group-by-time interactions, with linear contrasts used to determine significant differences among groups. One-way ANOVA
was used to compare tracer-determined (with and without the acetate
correction factor) and indirect calorimetry-determined rates of fat
oxidation, with method as the dependent variable. For all comparisons,
significance was set at P 0.05. For the LFD1 group, one
animal was an outlier for all processes, as estimated from
14C kinetics, and thus any analyses of variables determined
from 14C were made with only 9 animals in the LFD1 group.
For all other analyses, this animal was included. Data for insulin and
intracellular cycling were log transformed to achieve homogeneity of variance.
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RESULTS |
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Body Weight, Energy Intake, and Body Weight Gain
Rats entered the study with body weights that were not significantly different (Table 2). During the baseline period, energy intake, body weight gain, and body weight were not significantly different among groups. The first week of HFD feeding resulted in anticipated outcomes based on the study design: energy intake, body weight gain, and body weight were significantly greater in OP than in OR (Table 2). Of note, over the 5 wk of HFD, cumulative energy intake by OP5 rats exceeded that in OR5 rats by only 11%, but body weight gain in OP5 rats was 24% greater than that observed in OR5 rats.
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Fat Pad Weights, Fat Cell Volume, and Fat Cell Number
In animals studied after 1 wk of HFD, total fat pad weight (summed weights of epididymal, retroperitoneal, and mesenteric fat pads) was not significantly different across groups (Table 2). After 5 wk of HFD feeding, total fat pad weight was significantly greater in OP5 compared with either OR5 or LFD5 (PEnergy Expenditure and RQ
Energy expenditure and npRQ were not significantly different across groups (pooled average for energy expenditure: 0.037 ± 0.002 kcal/min; pooled average for npRQ: 0.723 ± 0.011, respectively). Energy expenditure, normalized to body weight, was not significantly different among 1-wk rats (pooled 1-wk average: 0.12 ± 0.006 kcal · kgCirculating Substrates, Urinary Nitrogen, and Plasma Hormones
Glycerol concentrations were significantly higher in OP5 compared with LFD5 rats (Table 3). Glucose concentrations were significantly lower in OP5 rats relative to LFD5 rats (Table 3). Norepinephrine concentrations were significantly higher in OR1 rats compared with LFD1 rats (Table 3). No other significant differences existed in substrates, hormones, or urinary nitrogen among groups (Table 3).
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Glycerol Ra
By 120 min of infusion, glycerol specific activity had reached steady state in both 1-wk and 5-wk rats (Fig. 1). Glycerol Ra (µmol/min) was not significantly different among 1-wk rats (Fig. 2A) but was significantly greater in both OP5 and OR5 relative to LFD5. Glycerol Ra was increased twofold in OP5 vs. OP1 (7.3 ± 0.4 vs. 3.6 ± 0.4 µmol/min), whereas it was only 38% greater in OR5 compared with OR1 (5.8 ± 0.8 vs. 4.2 ± 0.4 µmol/min). Similar results were obtained when glycerol Ra was expressed relative to fat cell number (µmol · 106 cells
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Acetate Correction Factor
Initial body weight, energy intake, body weight gain, body weight, and fat pad weights were not different for rats used to determine an acetate correction factor from values for rats used in studies in which fat oxidation was determined (data not shown). The rate of 14CO2 production (dpm/min) for both 1-wk and 5-wk animals was in steady state by 120 min of the [1-14C]acetate infusion (Fig. 3). No significant differences in the acetate correction factor were found among groups (Table 4).
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Plasma FFA Kinetics and Plasma FFA Oxidation
Steady state was reached for plasma FFA specific activity by 120 min of infusion in both 1-wk and 5-wk rats (Fig. 4A). When expressed in absolute terms (µmol/min, Fig. 5A) or relative to body weight (µmol · kg
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The rate of 14CO2 production (dpm/min) for both
1-wk and 5-wk animals was in steady state by 120 min (Fig.
4B). The only significant differences in rates of plasma FFA
oxidation were observed when expressed relative to body weight
(µmol · kg1 · min
1).
Rates were significantly lower in OP5 and LFD5 vs. OP1 and LFD1,
respectively (Fig. 5B).
FFA Reesterification and Triglyceride Cycling
Reesterification (ratio of FFA Ra to glycerol Ra) was significantly greater in OP5 vs. LFD5 only (P
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Comparison of Tracer-Determined and Indirect Calorimetry-Determined Rates of Fat Oxidation
With no application of the acetate correction factor, and with the assumption that palmitate comprises 44% of the total FFA pool (11), tracer-estimated rates of fat oxidation were significantly lower than rates determined by indirect calorimetry, regardless of group (Table 6). With the application of the acetate correction factor (Table 4), rates of fat oxidation were no longer significantly different from rates determined by indirect calorimetry in some (OR1 and LFD1) but not all groups. When data were pooled, tracer-determined rates of fat oxidation left uncorrected were significantly lower than both corrected rates and rates determined by indirect calorimetry. Likewise, with the data pooled, corrected rates determined by tracer methods were significantly lower than rates estimated by indirect calorimetry.
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DISCUSSION |
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General Findings
Previous data from our laboratory suggested that, compared with OR rats, the 24-h RQ was higher in OP rats fed a HFD for 4-8 wk (4). More recent data demonstrated the "daytime RQ," measured at a time when rats eat very little and thus representative of a fasted RQ, was also significantly higher in OP rats (19). However, no comparisons of in vivo plasma FFA oxidation have been made between rats earlier in their exposure to a HFD, a time when obesity development in this model is at its most accelerated rate. Thus the primary purpose of this investigation was to determine whether fasted rates of fat oxidation, lipolysis, or reesterification contributed to the development of obesity in OP rats. Data reported here suggest that, if low rates of fat oxidation contribute to obesity development, the contribution does not occur at times when fat oxidation is at or near maximum rates (i.e., 24-h fasted conditions). Lipolysis and reesterification had not been previously studied using this model of HFD-induced obesity. Data presented here suggest that, between 1 and 5 wk of HFD feeding, basal lipolysis and reesterification may work in concert to expand fat cell volume and increase fat pad weight in both OP and OR rats, although more so in OP rats (Fig. 2 and Table 5).Fat Oxidation
In the 24-h-fasted state, OP1 rats oxidized fat at rates no different from OR1 rats (Fig. 5). It can be argued that the 24-h fast used in the present study maximized fat oxidation in both OP and OR rats, masking any differences between them. However, we have previously demonstrated that the ratio between the maximum activities of phosphofructokinase andTwo independent methods were used to estimate fat oxidation. Comparison of the results from these two methods yielded several important findings (Table 6). First, as previously reported (25), when corrected for 14C label fixation, tracer methods of plasma FFA oxidation better matched indirect calorimetry determinations of whole body fat oxidation [rates of fat oxidation (n = 67): 3.2 ± 0.09, 2.4 ± 0.09, and 1.7 ± 0.06 mg/min; indirect calorimetry, tracer methods with and without correction, respectively; Table 6]. Second, the acetate correction factor used was not different, regardless of diet or group. Thus 14C label fixation appears to be unaffected by significant alteration of the fat and carbohydrate contents of the diet. In addition, because the acetate correction factor determined here (0.72 on average) was somewhat higher than that reported in 12-h-fasted adults (0.56; Ref. 25), it is advisable to determine a correction factor within the model being used.
It has been previously reported in humans that tracer estimates of fat oxidation remain slightly lower than values measured by indirect calorimetry, even when corrected for label fixation as performed here (25). The difference between the two methods could be due to oxidation of intratissue triglyceride stores, which would be included in indirect calorimetry measures but not captured by tracer methods. Conversely, in this study, we assumed that palmitate comprised 44% of the total FFA pool (11). It may be that, in our animals, palmitate comprised some lesser percentage of the total FFA pool. To equate the two methods, palmitate would have had to have occupied 33% of the total FFA pool, with no oxidation of intratissue triglyceride stores assumed.
It should be noted that the so-called "VA-mode" of infusion/sampling was used to estimate rates of appearance and oxidation. This method will result in lower rates compared with the "AV-mode" of infusion/sampling. It is unlikely, however, that the choice of infusion/sampling sites will significantly impact comparisons between groups (15). There were two groups (OP1 and LFD1) in which rates of plasma FFA oxidation exceeded plasma FFA Ra, because, in these two groups, either plasma FFA oxidation was overestimated or plasma FFA Ra was underestimated. It is worth noting that, when compared in total (n = 67), rates of FFA appearance were nearly equal to rates of plasma FFA oxidation (9.5 ± 0.5 vs. 9.9 ± 0.4 µmol/min).
Basal Lipolysis and Reesterification
The greater fat pad weight in OP5 rats was due largely to increased fat cell volume (Table 2) and did not appear to relate to differences in basal concentration of catecholamines, corticosterone, or insulin (Table 3). This would predict that reesterification was in excess of lipolysis in OP rats relative to OR rats. Regardless of how expressed, lipolysis (estimated using glycerol Ra) was not significantly different between OP and OR rats (Fig. 2, A-C). Thus reduced basal lipolysis, per se, does not contribute to the expansion of fat cell volume and fat pad weight in OP rats. However, whereas glycerol Ra increased in both OP and OR rats from 1 to 5 wk (significantly so in OP), FFA Ra did not (Fig. 5A). Taken together, these data suggest that, relative to the 1-wk rats, a greater proportion of the FFAs derived from lipolysis subsequently underwent reesterification within the adipocyte in the 5-wk rats (measured either as the ratio between FFA Ra and glycerol Ra or as intracellular cycling). Indeed, the significantly greater increase in lipolysis from 1 to 5 wk in OP rats relative to OR rats was accompanied by a significantly greater increase in intracellular triglyceride cycling in OP rats (Fig. 2A and Table 5).Recent data suggest that glycerol Ra may not accurately represent adipose tissue lipolysis (5, 13). If so, we have overestimated adipose tissue lipolysis in both OP and OR rats. However, our conclusion that a greater mismatch between fasted adipose tissue lipolysis and reesterification contributed to or was a consequence of obesity development in OP rats is still reasonable, because it was based on a greater increase in glycerol Ra from 1 to 5 wk in OP rats relative to OR rats. Only if the excess glycerol Ra observed in OP5 rats originated from tissues other than adipose tissue or from circulating very low-density lipoprotein triglyceride would this conclusion be in error.
The mechanism(s) that might permit rates of reesterification to increase to a greater extent in OP rats relative to OR rats from 1 to 5 wk of HFD cannot be delineated from the data presented here. The greater increase in absolute rates of lipolysis between 1 and 5 wk in OP rats would be a likely source for intracellular FFAs. The greater increase in intracellular reesterification implies that within adipocytes of OP rats, there is either greater availability of 1) glycerol-3-phosphate resulting from an increased glucose uptake and subsequent metabolism to glycerol-3-phosphate, or 2) monoacyl- and/or diacylglycerides for free fatty acid reesterification. Future work is required to establish the physiological role of increased reesterification in this model of obesity development.
The differences described in this study between OP and OR rats were measured under 24-h-fasted conditions and do not reflect a typical metabolic setting in either group of rats. Knowing whether or not lipolysis and reesterification differ between OP and OR rats under insulin-stimulated conditions requires additional work, although fasting insulin concentrations measured here suggest that whole body insulin action was not significantly different between OP and OR rats (Table 3). Adipose tissue metabolism and insulin action have not been well characterized in this model. If a mismatch between lipolysis and reesterification under insulin-stimulated conditions is contributing to obesity development, we would predict that adipose tissue of OP1 rats would be more sensitive to insulin action than would the adipose tissue of OR1 rats.
The rates of extracellular reesterification reported in Table 5 for OP1 and LFD1 rats were significantly less than zero (P = 0.0007 for OP1 and P = 0.0018 for LFD1). Because extracellular reesterification was calculated as the difference between FFA Ra and plasma FFA oxidation, the negative rates were the result of either an underestimation of FFA Ra or an overestimation of plasma FFA oxidation in these two groups. As such, we presented the means ± SE for extracellular cycling, but because the values could not be reflective of any physiological process, we chose not to make any statistical comparisons. If anything, these data suggest that whole body extracellular reesterification is essentially zero under fasted conditions.
In conclusion, there are several important findings from this study. First, fasted rates of fat oxidation were not impaired in OP rats. Thus the higher 24-h RQ (4) and higher daytime RQ (19) reported previously for OP rats must be driven by processes other than low fasted rates of fat oxidation. Second, basal rates of lipolysis were not significantly different between OP and OR rats, and with 5 wk of HFD, lipolytic rates actually increased in both OP and OR rats, although significantly so only in OP rats. At the same time, intracellular reesterification increased as well. Taken together, these findings implicate an earlier or greater increase in intracellular reesterification relative to lipolysis with HFD feeding to be one process contributing to the larger fat cell volume, and thus fat pad mass, in OP rats.
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ACKNOWLEDGEMENTS |
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We gratefully acknowledge the metabolic core of the Colorado Clinical Nutrition Research Unit (P30 DK-48520-01) for assistance with insulin measurements.
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FOOTNOTES |
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This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-47416 (M. J. Pagliassotti) and DK-38088 (J. O. Hill) and the Colorado Agricultural Experiment Station Project 616 (C. L. Melby).
Address for reprint requests and other correspondence: S. R. Commerford, Arizona State University, Dept of ESPE, PE Building East, Rm. 107B, Tempe, AZ 85287-0404 (E-mail: Renee.Commerford{at}asu.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.
Received 23 August 1999; accepted in final form 17 May 2000.
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