Fat oxidation, lipolysis, and free fatty acid cycling in obesity-prone and obesity-resistant rats

S. Renee Commerford1,2,3, Michael J. Pagliassotti1,3, Christopher L. Melby2, Yuren Wei1, Ellis C. Gayles1, and James O. Hill1

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


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 beta -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.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 >= 48 g) or OR (body wt gain <= 40 g) groups. These weight gain criteria were selected a priori to ensure that OP and OR rats were distinct with respect to weight gain and that the weight gain criteria used to identify OP and OR remained consistent across cohorts. Rats with weight gains between 41 and 47 g were not studied further (58.7% of total rats fed the HFD). Rats were studied either after 1 wk of HFD (OR1, OP1, and LFD1) or after 5 wk of HFD feeding (OR5, OP5, and LFD5). Throughout the protocol, food intake was measured three times per week, and body weight was recorded once per week.

                              
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Table 1.   Diet composition

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 recover >= 4 days. During recovery, rats ate their respective diets ad libitum. For a rat to be studied, body weight was required to be >= 92.5% of presurgery weight. Rats not meeting this weight by 7 days postsurgery were not studied.

Experimental 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 · kg-1 · 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 (VO2) and CO2 production (VCO2) 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.

14CO2 production was determined by trapping expired gases in a solution of 2:1 benzathonium hydroxide-methanol over a 5-min period. One-milliliter aliquots were counted (Beckman LS6500) in triplicate to determine the rate of 14CO2 appearance in expired air.

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
FFA R<SUB>a</SUB><IT>=</IT>

<FR><NU>rate of [<IT>1-<SUP>14</SUP></IT>C] palmitate infusion (dpm<IT>/</IT>min)</NU><DE><SUP><IT>14</IT></SUP>C of palmitate (dpm<IT>/</IT>ml)<IT>/</IT>FFA concentration (<IT>&mgr;</IT>mol<IT>/</IT>ml)</DE></FR>
The denominator in this equation corrects for the fact that palmitate comprises some unknown fraction of the total plasma FFA pool, and thus adjusts, by direct proportion, Ra to FFA Ra. This method of determining FFA Ra has been used by others (1). Specific activity for both glycerol and FFA was calculated using a rolling average across t = 120, 130, and 140 min. Glycerol Ra was normalized to mean fat cell number (106 fat cells) and mean cell volume.

Whole body fat oxidation by indirect calorimetry. After correction to STPD (standard temperature and pressure, dry), VO2 and VCO2 were calculated using a rolling average over the three time points collected to give a single value for each. The mean value for both VO2 and VCO2 (ml/min) was extrapolated to 140 min to determine total VO2 and VCO2 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 VO2, 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
(1 cell<IT>/</IT>mean cell volume in pl)<IT>·</IT>(pl<IT>/0.95 </IT>ng lipid)<IT>·</IT>(<IT>10<SUP>9</SUP></IT>)<IT>=</IT>cell<IT>/</IT>g lipid
Fat cell number within a given tissue was calculated using the total mass of tissue times the number of cells per gram of tissue. Fat cell number was also determined on a subset of animals from each group (n = 5/group for each of epididymal, retroperitoneal, and mesenteric fat pads) after total lipid content was determined. This value was then applied to calculate fat cell number as follows
(fat pad weight in g)<IT>·</IT>(<IT>% </IT>lipid content)<IT>·</IT>(<IT>10<SUP>6</SUP></IT>)<IT>=&mgr;</IT>g lipid<IT>/</IT>fat pad

cell volume (pL<IT>/</IT>cell)<IT>·</IT>(<IT>0.95 </IT>ng lipid<IT>/</IT>pl)<IT>·</IT>(<IT>10<SUP>−3</SUP></IT>)<IT>=&mgr;</IT>g lipid<IT>/</IT>cell
Fat cell number was calculated by dividing microgram of lipid per cell by microgram of lipid per fat pad.

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
plasma palmitate oxidation<IT>=</IT>{expired<SUP><IT> 14</IT></SUP>CO<SUB><IT>2</IT></SUB> (dpm<IT>/</IT>min)<IT>/</IT>

[<IT>1-<SUP>14</SUP></IT>C]palmitate infusion rate (dpm<IT>/</IT>min)}<IT>×</IT>FFA R<SUB>a</SUB>
Palmitate oxidation was corrected to total FFA oxidation by accounting for the proportion of the plasma FFA pool that is comprised of palmitate (44% in chow-fed rats, Ref. 11). The acetate correction factor was applied to account for the underestimation of 14CO2 production from the oxidation of [1-14C]palmitate due to label fixation (25).

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
intracellular cycling<IT>=3</IT>(glycerol R<SUB>a</SUB>)<IT>−</IT>FFA R<SUB>a</SUB>
Extracellular cycling, defined as the hydrolysis of FFAs from glycerol that are then released from the adipocyte and are subsequently taken back up by the fat cell and reesterified, was calculated as
extracellular cycling<IT>=</IT>FFA R<SUB>a</SUB><IT>−</IT>plasma FFA oxidation

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.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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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Table 2.   Energy intake, body weight gain, body weight, fat pad weight, fat cell size, and fat cell number

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 (P <=  0.05). The increase in total fat pad weight at 1 vs. 5 wk of HFD was significantly greater in OP rats compared with either OR or LFD (P = 0.0243 and P = 0.0041, respectively). Neither fat cell volume nor fat cell number was significantly different among OP1, OR1, and LFD1. Among 5-wk rats, fat cell volume, but not fat cell number, was significantly greater in OP5 compared with OR5 and LFD5 (P = 0.0186 and P = 0.0044, respectively; Table 2).

Energy 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 · kg-1 · min-1) or among 5-wk rats (pooled 5 wk average: 0.093 ± 0.004 kcal · kg-1 · min-1).

Circulating 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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Table 3.   Circulating substrates, hormones, and urinary nitrogen

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-1 · min-1; Fig. 2B). When glycerol Ra was expressed relative to fat cell volume (Fig. 2C), no significant differences were detected among groups.


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Fig. 1.   Glycerol specific activity during the final 20 min of [2-3H]glycerol infusion. OP, obesity prone; OR, obesity resistant; LFD, low-fat diet controls. Values are means ± SE.



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Fig. 2.   Glycerol rate of appearance (Ra) expressed in µmol/min (A), relative to fat cell number (B), and relative to fat cell volume (C). Values reported are means ± SE of samples taken at 120, 130, and 140 min of infusion. Letters above bars are derived from Tukey's post hoc comparisons; values of bars sharing common letters are not significantly different (P <=  0.05). * Glycerol Ra expressed relative to 106 cells was logarithmically transformed. Data are means and SE without transformation.

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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Fig. 3.   14CO2 production during the final 20 min of [1-14C]acetate infusion. Values are means ± SE.


                              
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Table 4.   Acetate correction factor

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-1 · min-1, Fig. 5B), FFA Ra was not significantly different between 1-wk and 5-wk rats.


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Fig. 4.   Plasma free fatty acid (FFA) specific activity (A) and 14CO2 production (B) during the final 20 min of [1-14C]palmitate infusion. Values are means ± SE.



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Fig. 5.   A: absolute rates (µmol/min) of plasma FFA appearance (solid bars) and oxidation (open bars) in 1-wk and 5-wk rats. B: rates of FFA appearance (solid bars) and oxidation (open bars) relative to body wt in 1-wk and 5-wk rats. Values are means ± SE of samples taken at 120, 130, and 140 min of infusion. OP1, n = 12; OP5, n = 14; OR1, n = 11; OR5, n = 10; LFD1, n = 9; LFD5, n = 11. * Plasma FFA appearance significantly lower than plasma FFA oxidation (P <=  0.05). ** Plasma FFA oxidation significantly lower than respective 1-wk rate (P <=  0.05).

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 · kg-1 · 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 <=  0.05). Intracellular cycling (µmol/min, µmol · 106 cell-1 · min-1 and µmol · pl-1 · min-1) was not significantly different among 1 wk rats but was significantly higher in both OP5 and OR5 compared with LFD5 rats when expressed as µmol/min (Table 5). When normalized to either fat cell number or fat cell volume, however, only OP5 rats demonstrated significantly higher rates of intracellular cycling relative to LFD5 (P <=  0.05). Extracellular cycling rates were negative in both OP1 and LFD1 groups (significantly different from zero, P = 0.0007 and P = 0.0018, respectively) but were not for the remaining groups. Because the negative extracellular cycling rates were the result of either an underestimation of FFA Ra or an overestimation of plasma FFA oxidation, and thus not representative of a true rate, statistical comparisons are not reported.

                              
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Table 5.   Plasma FFA kinetics

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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Table 6.   Comparison of tracer- and indirect calorimetry-determined fat oxidation


    DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 and beta -hydroxyacyl-CoA dehydrogenase in skeletal muscle was significantly higher in OP compared with OR rats after 1 and 2 wk of HFD feeding, suggesting that OP rats might have a lower capacity to oxidize fat relative to carbohydrate compared with OR rats. Data presented here suggest that fat oxidation in the fasted state does not contribute to differences in weight gain or fat gain between OP and OR rats.

Two 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.


    ACKNOWLEDGEMENTS

We gratefully acknowledge the metabolic core of the Colorado Clinical Nutrition Research Unit (P30 DK-48520-01) for assistance with insulin measurements.


    FOOTNOTES

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.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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