Investigation of in vivo fatty acid metabolism in AFABP/aP2–/– mice

Rachel A. Baar,1 Carlus S. Dingfelder,1 Lisa A. Smith,2 David A. Bernlohr,2 Chaodong Wu,2 Alex J. Lange,2 and Elizabeth J. Parks1

1Department of Food Science and Nutrition, University of Minnesota, St. Paul; and 2Departments of Biochemistry, Molecular Biology, and Biophysics, University of Minnesota, Minneapolis, Minnesota

Submitted 15 June 2004 ; accepted in final form 3 September 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The metabolic impact of the murine adipocyte fatty acid-binding protein (AFABP/aP2) on lipid metabolism was investigated in the AFABP/aP2–/– mouse and compared with wild-type C57BL/6J littermates. Mice were weaned on a high-fat diet (59% of energy from fat) and acclimated to meal feeding. Stable isotopes were administered, and indirect calorimetry was performed to quantitate fatty acid flux, dietary fatty acid utilization, and substrate oxidation. Consistent with previous in situ and in vitro studies, fasting serum nonesterified fatty acid (NEFA) release was significantly reduced in AFABP/aP2–/– (17.1 ± 9.0 vs. 51.9 ± 22.9 mg·kg–1·min–1). AFABP/aP2–/– exhibited higher serum NEFA (1.4 ± 0.6 vs. 0.8 ± 0.4 mmol/l, AFABP/aP2–/– vs. C57BL/6J, respectively) and triacylglycerol (TAG; 0.23 ± 0.09 vs. 0.13 ± 0.10 mmol/l) and accumulated more TAG in liver tissue (2.9 ± 2.3 vs. 1.1 ± 0.8% wet wt) in the fasted state. For the liver-TAG pool, 16.4 ± 7.3% of TAG-fatty acids were derived from serum NEFA in AFABP/aP2–/–. In contrast, a significantly greater portion of C57BL/6J liver-TAG was derived from serum NEFA (42.3 ± 25.5%) during tracer infusion. For adipose-TAG stores, only 0.29 ± 0.04% was derived from serum NEFA in AFABP/aP2–/–, and, in C57BL/6J, 1.85 ± 0.97% of adipose-TAG was derived from NEFA. In addition, AFABP/aP2–/– preferentially oxidized glucose relative to fatty acids in the fed state. These data demonstrate that in vivo disruption of AFABP/aP2–/– leads to changes in the following two major metabolic processes: 1) decreased adipose NEFA efflux and 2) preferential utilization of glucose relative to fatty acids.

adipocyte fatty acid-binding protein; fatty acid turnover; triacylglycerol; nonesterified fatty acids; mouse model


THE ADIPOCYTE FATTY ACID-BINDING PROTEIN (AFABP/aP2) is a member of the mammalian intracellular fatty acid-binding protein multigene family (4). AFABP/aP2 is the product of the FABP4 gene mapped to mouse chromosome 3 and found in a cluster of other FABP genes, including the epithelial FABP (EFABP; see Refs. 2 and 4). AFABP/aP2 is highly expressed in white adipocytes, brown adipocytes, and macrophages and binds a variety of long-chain fatty acids in a 1:1 molar stoichiometry with affinities ranging from 100 to 300 nM (2). As a class, the fatty acid-binding proteins are hypothesized to be responsible for fatty acid trafficking in the cell, facilitating the intracellular diffusion of insoluble lipids between cellular compartments (1, 4, 11, 27). When maintained on a low-fat chow diet, AFABP/aP2 null mice develop normally, are fertile, and gain a similar amount of weight as their wild-type counterparts (14). Biochemically, primary adipocytes isolated from AFABP/aP2 null mice exhibit decreased efflux of fatty acids and an accumulation of cytosolic nonesterified fatty acid (NEFA; see Refs. 2 and 5). Published studies have reported that basal and stimulated adipocyte lipolysis was both unchanged in AFABP/aP2–/– mice (25) and reduced in the basal state, but not different with the stimulation (24) in vitro. However, stimulation of lipolysis in vivo was shown to be significantly reduced (29). The molecular basis for decreased lipolysis in AFABP/aP2–/– adipocytes is linked to a physical association between AFABP and the hormone-sensitive lipase, a complex that facilitates efflux of fatty acids from the fat cell (26). When AFABP/aP2 null mice were placed on a high-fat diet to induce obesity, insulin resistance was attenuated, as shown by improved glucose and insulin tolerance tests (14, 29). In contrast, overexpression of FABPs in adipocytes results in increased lipolysis and potentiated characteristics of insulin resistance (10). Because glucose disposal is largely mediated via the skeletal muscle, the AFABP/aP2 null mouse provides a model to investigate the influence of fatty acid flux through adipocyte triacylglycerol (TAG) pools on whole body substrate oxidation. Given the biological role of FABPs in lipolysis, we chose an experimental protocol likely to reveal maximal differences in metabolic parameters linked to energy metabolism and substrate utilization. This protocol included feeding a high-fat diet in a meal-feeding regimen that included a long-term (16-h) fast to promote robust lipolysis. Accordingly, the goals of the present study were to 1) quantitate in vivo fatty acid efflux from adipose tissue and fatty acid reesterification rates in tissues, 2) measure the contribution of dietary TAG and newly made fatty acids (via de novo lipogenesis) to tissue TAG stores, and 3) determine whether substrate oxidation differed between AFABP/aP2 null and wild-type mice. We hypothesized that NEFA efflux from the adipocyte would be reduced, leading to elevations in peripheral glucose oxidation in AFABP/aP2 null mice. Our findings supported this hypothesis; however, the data also suggest that altered regulation of adipose NEFA flux impacted whole body fatty acid utilization, leading to greater flux of dietary fatty acids toward liver TAG stores.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animal husbandry. Male AFABP/aP2 knockout mice and wild-type C57BL/6J littermates were weaned onto a high-fat diet (product no. F3282; Bio-Serv, Frenchtown, NJ) providing 15.8% of energy as protein (197.4 g/kg), 59.4% of energy as fat (358.0 g/kg), and 24.8% of energy as carbohydrate (358.2 g/kg). The primary source of fat in the diet was lard. Mice were maintained on a 12:12-h reverse light-dark cycle (dark cycle was from 6:00 AM to 6:00 PM), given free access to water, and acclimated to a meal-feeding schedule of 6 h/day from 11:30 AM to 5:30 PM. Stable isotopes were purchased from Isotec (Miamisburg, OH). All procedures were approved by the University of Minnesota Animal Care and Use Committee.

Stable isotope infusion during fasting (experiment I). Mice were allowed to eat at libitum for ~6 wk postweaning to develop diet-induced obesity. At this time, mice were acclimated to the meal-feeding regimen described above. At ~13 wk of age, an indwelling catheter was surgically placed under sterile procedure in the jugular vein and advanced to the right atrium, as previously described (23). Mice were allowed to recuperate for 3–4 days and were infused after a 16-h fast with stable isotope at a rate of 5 µl/min using a Harvard Apparatus syringe pump (model 11; Harvard Apparatus, Holliston, MA) to quantitate NEFA flux (30). The isotope [13C4]palmitate (70 µg·kg–1·min–1) complexed to albumin was infused for 20, 45, 60, 90, or 180 min, after which mice were killed with a 2 µl/g intramuscular injection of a 2:1 mixture of ketamine (100 mg/ml)-xylazine (20 mg/ml). Blood, liver, and adipose tissues were immediately harvested.

Stable isotope administration during feeding (experiment II). Mice, maintained on the high-fat diet and reverse light-dark cycle, were acclimated to the meal-feeding regimen for 8 days. For the purposes of identifying fatty acid sources in tissues, two stable isotopes, [13C1]acetate (4 g) and glyceryl tri(hexadecanoate-d31) (8.9 g), were added as described previously (6) to 200 g of the high-fat diet for 5 days. Food intake was assessed using a food cup designed for metabolic studies attached to the side of a standard cage. On day 5 after a 16-h fast, mice began feeding at the usual time and were allowed to consume food for 1 h. The rationale for allowing 1 h of food consumption is that the goal was to obtain data from animals actively eating, which occurs reliably (21). After this 1 h, the animals were killed as described above, and serum, liver, adipose, and muscle tissues were immediately harvested.

Serum and tissue collection/lipid extraction. Blood was obtained at the end of experiments I and II through an open-heart puncture, and serum was immediately separated. Liver and epididymal adipose (and muscle in experiment II only) tissue depots were excised and weighed; portions of the tissue samples were removed for lipid extraction and analysis, and extra samples were immediately frozen. Total tissue and serum lipids were extracted via the extraction method of Folch et al. (7) with trinonadecanoin and pentadecanoic acid as internal standards (Sigma, St. Louis, MO). Serum NEFA and TAG were separated by TLC and transesterified to methyl esters, as described previously (17), for analysis by gas chromatography-mass spectrometry (GC-MS).

GC-MS, fatty acid composition, and enzymatic assays. GC-MS was performed on an HP 6890 GC with Mass Selective Detector HP 5973 MS (Hewlet-Packard, Eagan, MN) using electron-impact ionization with an HP-1, 25-m column (Hewlet-Packard, Palo Alto, CA) and helium as the carrier gas. The molecular ions of methyl-palmitate from the NEFA pool in serum and TAG from serum, liver, and adipose were analyzed for mass-to-charge ratios of 270, 271, 272, 274, 300, and 301 (M0, M1, M2, M4, M30, and M31, respectively), as described previously (6). Comparable ion peak areas between a standard curve and biological samples were achieved by either adjusting the volume injected, diluting, or concentrating the sample when needed. Fatty acid composition of the TAG fraction in liver and adipose was determined by gas chromatography with flame-ionization detection (FID; see Ref. 19). Individual fatty acids were identified by retention time and compared with the internal standard for quantification. Serum TAG, NEFA, and glucose concentrations were measured in triplicate enzymatically (Wako Chemicals USA, Richmond, VA) using a Microtek EL340 microplate reader (Bio-Tek Instruments, Winooski, VT). For determination of hepatic and muscle content of fructose 2,6-bisphosphate (F-2,6-P2), the metabolite was extracted from frozen liver or muscle tissue in 10 vol of 50 mM NaOH and kept at 80°C for 5 min. The extract was cooled and neutralized at 0°C by addition of ice-cold 1 M acetic acid in presence of 20 mM HEPES. After centrifugation at 8,000 g for 10 min, supernatant was collected and assayed for F-2,6-P2 by the 6-phosphofructo-1-kinase activation method, as described previously (31).

Indirect calorimetry. Indirect calorimetry was assessed on mice, 12–16 wk of age, using a computer-controlled, open-circuit system (Applied Electrochemistry, Pittsburgh, PA). A primary gas standard containing 20.0% oxygen, 1.0% carbon dioxide, and nitrogen balance (National Specialty Gases, Research Triangle Park, NC) was used to calibrate carbon dioxide and oxygen sensors before each mouse entered the chamber for indirect calorimetry measurement. For all animals (n = 7 for each group), measurements were taken from 8:00 AM to 11:00 AM for fasting and 12:00 PM to 3:00 PM for fed-state data (i.e., in each mouse, fasting and fed-state data were collected on different days). All animals were weighed before and after the metabolic chamber measurements. Values of oxygen consumption (ml/min) and carbon dioxide production (ml/min) were taken every 2 min for 3 h; all time points between 1 and 2 h were averaged and used for calculations. This time of data collection allowed the animal to begin eating from 11:30 AM to 12:00 PM, and then 0.5 h to acclimate to eating in the chamber before data were used for analysis. The time of data collection for indirect calorimetry also coincided with the same time frame that data were collected for fed-state measurements of blood metabolite concentrations. Indirect calorimetry data were used to calculate respiratory quotient (RQ), energy expenditure, and glucose and fat oxidation rates as described (16).

Calculations and statistical analysis. Adipose total fatty acid release was calculated by the isotope dilution method (30) based on the percentage of palmitate (16:0) in the total NEFA pool, as determined by GC-MS. The rate of appearance of NEFA (Ra NEFA) was calculated as follows:


The rate was corrected for the percentage of NEFA that was palmitate, as determined by gas chromatography with FID detection. TAG derived from the NEFA pool was calculated by dividing steady-state enrichments of TAG-[13C4]palmitate from serum, liver, and adipose by the steady-state enrichments of [13C4]palmitate in the NEFA pool. Newly made fatty acids in liver, adipose, and muscle tissues were calculated using mass isotopomer distribution analysis (9). Results are expressed as means ± SD. Statistical differences between AFABP/aP2–/– mice and wild-type littermates were determined by unpaired Student's t-tests and ANOVA. For the comparison of tissue TAG-fatty acid composition with dietary TAG-fatty acids, a nonparametric one-sample sign test was used. Statistical analyses were performed using Microsoft Excel (2002 version; Microsoft, Redmond, WA) and Statview (version 5.0.1; SAS Institute, Berkeley, CA). A P value <0.05 was considered significant.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Physiological characteristics. Mice were studied between the ages of 11 and 19 wk. As shown in Table 1, body weights between the two groups of mice were not different at the end of experiment I (28.1 ± 1.7 vs. 26.9 ± 2.5 g in AFABP/aP2–/– and wild-type mice, respectively) or experiment II (29.6 ± 3.1 vs. 29.1 ± 1.9 g). Food intake was assessed daily in experiment II and demonstrated no difference between the groups (Table 1, P = 0.17). The AFABP/aP2 null mice had significantly lower concentrations of serum glucose compared with the wild-type mice in the fasting state (10.12 ± 6.84 vs. 17.84 ± 5.63 mmol/l, P = 0.005), a result that has been documented previously (14, 25). The values observed in the present study were higher than previously reported, most likely because of the effect of the anesthetic used (15). Surprisingly, serum NEFA concentrations were 70% higher in AFABP/aP2–/– mice (1.43 ± 0.55 vs. 0.84 ± 0.38 mmol/l, P = 0.002) and serum TAG concentrations were 76% higher (0.23 ± 0.09 vs. 0.13 ± 0.10 mmol/l, P = 0.006) in the fasting state. Serum NEFA and TAG found in this study most likely reflect metabolic differences associated with the meal-feeding regimen and the long-term (16-h) fasted state relative to ad libitum feeding and a short-term fast used in prior studies. In the fed state, NEFA concentrations were significantly higher in the AFABP/aP2–/– mice (2.20 ± 0.46 vs. 1.43 ± 0.57 mmol/l, P = 0.046); however, no significant difference was found between the groups for fed-state glucose or TAG concentrations. Liver weights between the groups were similar, although livers of AFABP/aP2–/– mice contained nearly threefold more TAG per wet weight of tissue compared with the wild type (2.9 ± 2.3 vs. 1.1 ± 0.8%, P = 0.008) in the fasted state. Isolated epididymal fat pads weighed 43% more compared with wild-type littermates (1.0 ± 0.3 vs. 0.7 ± 0.3 g, P = 0.001) in the fasted state. Consistent with in vitro lipolysis data, fasting serum NEFA release (Ra NEFA) from adipose tissue was 69% lower in the AFABP/aP2–/– mice (17.1 ± 9.0 vs. 51.9 ± 22.9 mg·kg–1·min–1, P = 0.001) relative to wild-type mice and 71% lower when analyzed as absolute fatty acid release per mouse (1.78 ± 0.98 vs. 6.11 ± 5.19 µmol·min–1·mouse–1, P = 0.009). Because serum NEFA can be transported in tissues and incorporated in TAG stores (8, 20), we calculated the percentage of various TAG pools derived from serum NEFA during the course of the isotope infusion. As shown in Table 2, for the liver-TAG pool, 16.4 ± 7.3% of TAG-fatty acids were derived from serum NEFA in AFABP/aP2–/– mice. In contrast, a significantly greater portion of wild-type liver-TAG was derived from serum NEFA (42.3 ± 25.5%) during tracer infusion. For adipose-TAG stores, only 0.29 ± 0.04% of this pool was derived from serum NEFA in AFABP/aP2–/– mice, and, in wild-type mice, 1.85 ± 0.97% of adipose-TAG was derived from NEFA (Table 2). These lower enrichments in adipose are because of the larger whole body adipose pool size compared with liver fat stores. Typically, the majority of serum TAG is found in very low-density lipoproteins released by the liver (3). In AFABP/aP2–/– mice, 20.8 ± 7.1% of serum TAG was derived from the hepatic production, whereas, in wild-type mice, the liver contributed 67.9 ± 28.5% the fatty acids ending up in serum TAG. Although the percentage of label incorporated in all tissues studied was significantly different between the two groups of mice, the observed lower fractional contribution of incorporation of NEFA label in tissue-TAG stores was because of the greater amounts of TAG in these tissues. For instance, when the percentage of liver derived from the serum NEFA pool was multiplied by the liver TAG content, no difference was found between the mouse strains in the absolute amount of reesterification that occurred during the infusion (0.019 ± 0.014 vs. 0.019 ± 0.010 mg·liver–1·min–1, P = 0.999 in AFABP/aP2–/– mice vs. wild-type mice, respectively).


View this table:
[in this window]
[in a new window]
 
Table 1. Characteristics of AFABP/aP2–/– and wild-type mice

 

View this table:
[in this window]
[in a new window]
 
Table 2. Metabolic characteristics of AFABP/aP2–/– and wild-type mice

 
In experiment II, no difference in the fractional synthesis rate of de novo lipogenesis (acetate incorporation into fatty acids) was found between the groups of mice for liver (10.8 ± 2.2 vs. 14.8 ± 5.1%, P = 0.072 in AFABP/aP2–/– mice vs. wild-type mice, respectively), adipose (3.6 ± 1.8 vs. 6.9 ± 4.4%, P = 0.086), or muscle tissue (6.3 ± 3.5 vs. 7.4 ± 3.7%, P = 0.328). The contribution of dietary TAG to tissue TAG pools was 20% higher in AFABP/aP2–/– liver tissue (P = 0.011) compared with the wild type. However, no differences were found in the contribution of dietary fatty acids to adipose or muscle tissue-TAG stores (Table 2).

Fatty acid composition and content in adipose and liver tissue in the fasted state. The fatty acid compositions of adipose- and liver-TAG stores were compared with the fatty acid composition of the diet. Percentages of saturated fatty acids (14:0, 16:0, and 18:0) were all lower in the tissues than in the diet, whereas the percentage of the polyunsaturated fatty acid, 18:2, was higher in tissue-TAG compared with dietary-TAG (Fig. 1). The percentage of liver-TAG 16:1, 18:1, and 20:1 was similar to that in the diet in AFABP/aP2–/– mice, whereas, in wild-type mice, liver-TAG monounsaturated fatty acids were different from that of the diet composition, with 16:1 and 18:1 being lower and 20:1 being higher (Fig. 1A). In AFABP/aP2–/– adipose-TAG, percentages of 16:1 and 18:1 were higher, whereas 20:1 was lower than in the diet (Fig. 1B). In wild-type adipose, only the percentage of 18:1 was higher than in the diet. In addition to the percentage composition, the absolute amounts of TAG-fatty acids were also quantified (data not shown). Given the 2.5-fold higher TAG concentration in AFABP/aP2–/– liver tissue, the concentrations of each of the individual fatty acids were found to be quantitatively higher in these mice.



View larger version (24K):
[in this window]
[in a new window]
 
Fig. 1. Liver triacylglycerol (TAG)-fatty acid composition (A) and adipose-TAG-fatty acid composition (B) in AFABP/aP2–/– and wild-type mice in the fasted state and comparison with dietary TAG-fatty acids. AFABP, adipocyte fatty acid-binding protein. Filled bars, data from AFABP/aP2–/– mice; open bars, wild-type mice; hatched bars, percentage of the fatty acids in the diet. Values with different superscript letters are significantly different from one another.

 
Indirect calorimetry. From the measurements of respiratory gases, whole body carbohydrate and fatty acid oxidation rates were quantified. For wild-type mice, the fasted RQ did not differ from the fed state (0.76 ± 0.02 vs. 0.79 ± 0.08, P = 0.205; Fig. 2A), a finding that was not surprising given the extremely high-fat nature of the diet. Interestingly, although AFABP/aP2–/– mice also had a low RQ in the fasted state (0.75 ± 0.01), their fed-state RQ rose significantly (0.80 ± 0.06, P = 0.031). Compared with fasting, fed-state energy expenditure was higher in both wild-type (0.49 ± 0.11 vs. 0.69 ± 0.10 J·g–1·min–1, fasted vs. fed, respectively) and AFABP/aP2–/– (0.44 ± 0.08 vs. 0.59 ± 0.07 J/g/min; Fig. 2B) mice. Between the groups, wild-type mice had a significantly higher energy expenditure compared with AFABP/aP2–/– mice in the fed state. For glucose oxidation rates, fasted rates were significantly lower than fed in both wild-type mice (5.82 ± 1.65 vs. 8.82 ± 3.71 mg·kg–1·min–1) and AFABP/aP2–/– mice (5.06 ± 2.72 vs. 10.89 ± 6.52 mg·kg–1·min–1, Fig. 2C). In wild-type mice, fat oxidation rose significantly from the fasted state to the fed state when expressed per kilogram mouse body weight (7.74 ± 2.62 vs. 10.80 ± 2.60 mg·kg–1·min–1; Fig. 2D) or when expressed as absolute fatty acid oxidation per mouse (0.94 ± 0.31 vs. 1.36 ± 0.32 µmol/min, P = 0.016). However, no differences were found between fasted- and fed-state fat oxidation rates in the AFABP/aP2–/– mice, when expressed relative to mouse body weight (7.28 ± 1.64 vs. 7.80 ± 2.32 mg·kg–1·min–1, P = 0.320) or in terms of total fat oxidized per mouse (0.84 ± 0.14 vs. 0.95 ± 0.35 µmol/min, P = 0.23). In summary, these data indicate that the well-documented improvement in insulin sensitivity found in AFABP/aP2–/– mice may result preferentially from oxidation of glucose in the fed state.



View larger version (28K):
[in this window]
[in a new window]
 
Fig. 2. Data from fasting and fed-state respiratory gas measurements and fructose 2,6-bisphosphate (F-2,6-P2) level, including respiratory quotients (RQ; A), energy expenditure (B), glucose oxidation (C), fat oxidation (D), and F-2,6-P2 production in muscle (E) and liver (F). Filled bars, data from AFABP/aP2–/–; open bars, data from wild-type mice. Data are means ± SD; sample sizes are 5–12 mice/group; NA, data not available.

 

    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The present study was undertaken to extend the in vitro and in situ analysis of AFABP/aP2 to the in vivo environment. AFABP/aP2 forms a physical complex with the adipocyte hormone sensitive lipase and facilitates efficient efflux of lipolytically-derived fatty acids from the fat cell (26). Consequently, the disruption of AFABP/aP2 results in reduced fatty acid efflux from adipose tissue cells in culture, as well as in response to pharmacological activation of lipolysis, and the present work clearly supports this finding using methods to quantitate metabolic turnover in vivo. AFABP/aP2 null mice exhibited a two- to threefold reduction in fatty acid release from adipose tissue in vivo after a 16-h fast, demonstrating for the first time that FABPs mediate efflux of fatty acids in normal physiology. A higher abundance of epididymal fat mass in the AFABP/aP2–/– mouse likely resulted from the decrease in lipolysis, since the other possible cause, greater food intake, was not observed in the present study (Table 1) or in previous research (25). Whether the larger epididymal depot in AFABP/aP2–/– is representative of a generalized expansion of adipose depots throughout the body is unknown, since other data have not found a difference in total adiposity, adipose cellularity, or depot size (Smith and Bernlohr, unpublished observations). Surprisingly, the reduction in rate of appearance of fatty acids in serum in the AFABP/aP2–/– mouse was also accompanied by elevations in serum NEFA concentrations in the fasted and fed states. Serum NEFA concentrations have been shown to be highly variable in mice, depending on the metabolic condition of the animals, and a large percentage of the NEFA in this pool can originate from spillover of dietary fatty acids during lipoprotein lipase-mediated lipolysis (28). Published studies have reported higher (24) and unaltered (10, 14) concentrations of serum NEFA in AFABP/aP2–/– mice relative to wild-type animals. Our studies revealed an increase in serum NEFA in AFABP/aP2–/– mice, a finding that may specifically reflect the use of the meal-feeding protocol that restricted mouse access to food 6 h/day in the dark cycle. Alternatively, it may be because of the source of the dietary fat (lard) or because of an interaction between this feeding regimen and the lack of weight gain typically observed when animals eat this diet ad libitum. This regimen was employed so that assessments of differences between fasting and fed states could be made with confidence (i.e., we could precisely control the duration of fasting and obtain fed-state data at a time when all animals were known to be eating). Previous studies have shown that, when this paradigm is used, mice eat normally (6), although excess weight gain, typically induced by consumption of a high-fat diet, is restrained (22). However, because both strains of mice were on the same feeding schedule, the differences observed between the AFABP/aP2–/– and wild-type mice remain valid and represent a viable paradigm for examining fatty acid flux in this rodent model. Previous studies in ad libitum-fed animals have shown that targeted disruption of AFABP leads to a subsequent increase in the expression of the other fatty acid-binding protein, keratinocyte (K)FABP, which only partially compensated for the loss of AFABP (5). In that study, although KFABP protein levels were increased, their total FABP level was still only ~5% of that of the wild type (5). The level of KFABP under the present meal-feeding paradigm is unknown. Another limitation of this study was that, because of technical constraints, the rate of release of fatty acids was assessed only in fasting animals. However, the elevation in free fatty acid concentration in both the fasting and fed states implicates fed-state events as the cause of a relative "backup" of fatty acids in the system. The data suggest that, although adipose fatty acid efflux was reduced, the reduction in peripheral fatty acid oxidation occurred earlier and to a greater extent, leading to an increase in serum NEFA concentrations. Several independent results from the present study support this concept. First, AFABP/aP2–/– mice had higher fasting concentrations of TAG in the blood and liver compared with wild-type mice. The two primary fates of NEFA in serum are either clearance to tissues (e.g., skeletal and cardiac muscle) followed by oxidation or clearance to liver followed by either storage as TAG or biosynthesis into lipoproteins (very low-density lipoprotein). The elevation of TAG in serum and liver suggests that these stores were expanded secondary to reduced oxidation of fatty acids by skeletal muscle. Second, the reduction in serum NEFA oxidation by muscle was mirrored by a reduction in the use of fatty acids derived from the diet. For example, when dietary palmitate (c16:0) was isotopically labeled, AFABP/aP2–/– mice accumulated a greater percentage of these fatty acids in liver-TAG pools over the 5-day period of labeling compared with wild-type animals. Internalization of dietary fatty acids by adipose and muscle in AFABP/aP2–/– mice may not have been reduced, since the percent contribution of labeled palmitate to TAG pools in these tissues was not different from in the wild type. Analysis of the different fatty acids in liver-TAG also demonstrated that liver fatty acid composition in AFABP/aP2–/– mice more closely resembled that of the diet, particularly for the monounsaturated fatty acids, which made up the majority of dietary fatty acids. Third, compared with wild-type mice, AFABP/aP2–/– mice exhibited a higher RQ in the fasting state, supporting a greater proportion of energy needs met by glucose oxidation during fasting. This was particularly surprising given the relatively long period of fasting in the present study (16 h), a duration after which usage of adipose tissue fatty acids would have been expected to be high. Last, when fed a high-fat diet, AFABP/aP2–/– mice demonstrated lower serum glucose concentrations, an increase in the amount of glucose burned in the fed state, and a failure to increase fat oxidation in the fed state. The extremely high-fat nature of the diet should have resulted in continued use of fatty acids to support energy expenditure in the fed state, as observed in the wild-type mice. However, even when dietary carbohydrate was in relatively low supply (~25% of energy in the diet), the AFABP/aP2–/– mouse exhibited greater glucose oxidation with feeding. The increased glucose oxidation by the muscle is supported by the elevation of glycolysis activator F-2,6-P2 seen in the AFABP/aP2–/– mouse vs. the wild type. The observed depression of F-2,6-P2 in the liver is indicative of increased hepatic glucose output and hepatic insulin resistance, which may be a compensatory response to lower blood glucose (18). The AFABP/aP2–/– mouse may represent an interesting case where muscle has increased insulin sensitivity and liver is insulin resistant, in which case the effect in the muscle dominates. All together, these observations point toward a block in fatty acid oxidation in the AFABP/aP2–/– mice.

The potential connection between trafficking of fatty acids within the adipocyte and regulation of substrate oxidation at the periphery points toward a potential role of adipokines in this process. Adipose tissue participates in the regulation of whole body energy utilization via the synthesis and secretion of a variety of hormones (adipokines). For example, one or more of the cell types present in adipose tissue synthesizes and secretes TNF-{alpha}, adiponectin, resistin, plasminogen activator inhibitor-1, IL-6, leptin, angiotensinogen, and growth and differentiation factor 3. Elevations in TNF-{alpha} have been shown to reduce insulin signaling at the insulin receptor, producing insulin resistance (12, 13). Hotamisligil et al. (14) have shown that AFABP/aP2–/– mice have a lower relative abundance of TNF-{alpha} mRNA than do wild-type littermates, although Shaughnessey et al. (25) found no difference in TNF-{alpha} from adipocytes derived from mice fed either a low-fat or high-fat diet. Because the levels of secreted adipokines in experimental animals after a long-term fast, as used in the studies described herein, have not been evaluated, such studies are currently underway.

In summary, the present results are the first to document reduced in vivo adipose fatty acid release in the AFABP/aP2–/– mouse. The metabolic mechanism by which lower adipose fatty acid efflux leads to the improved insulin sensitivity, previously documented in this mouse model, may be via reduced use of dietary fatty acids at the muscle beds. While supporting the concept that increased quantities of stored fat may function remotely to control substrate oxidation at skeletal muscle, these findings also highlight the importance of considering both fasting and fed state metabolism. Identifying the mediator(s) of these effects will be key to understanding the potential influence of adipocyte biology on fatty acid flux and whole body energy metabolism.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants 5 P30 DK-50456 (to E. J. Parks and C. Wu), DK-53189 (to D. A. Bernlohr), and DK-38354 (to A. J. Lange), and American Heart Association Scientist's Development Grant 0230022N (to E. J. Parks).


    FOOTNOTES
 

Address for reprint requests and other correspondence: E. Parks, Dept. of Food Science and Nutrition, Univ. of Minnesota, 1334 Eckles Ave., St. Paul, MN 55108 (E-mail: eparks{at}umn.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
 

  1. Bernlohr DA, Coe NR, and LiCata VJ. Fatty acid trafficking in the adipocyte. Semin Cell Dev Biol 10: 43–49, 1999.[CrossRef][ISI][Medline]
  2. Bernlohr DA, Simpson MA, Hertzel AV, and Banaszak LJ. Intracellular lipid binding proteins and their genes. Annu Rev Nutr 17: 277–303, 1997.[CrossRef][ISI][Medline]
  3. Chapman MJ. Comparative analysis of mammalian plasma lipoproteins. In: Methods in Enzymology. New York: Academic, 1996, p. 70–143.
  4. Coe NR and Bernlohr DA. Physiological properties and functions of intracellular fatty acid-binding proteins. Biochim Biophys Acta 1391: 287–306, 1998.[ISI][Medline]
  5. Coe NR, Simpson MA, and Bernlohr DA. Targeted disruption of the adipocyte lipid-binding protein (aP2 protein) gene impairs fat cell lipolysis and increases cellular fatty acid levels. J Lipid Res 40: 967–972, 1999.[Abstract/Free Full Text]
  6. Donnelly KL, Margosian MR, Sheth SS, Lusis AJ, and Parks EJ. Increased lipogenesis and fatty acid reesterification contribute to hepatic triacylglycerol stores in hyperlipidemic Txnip–/– mice. J Nutr 134: 1475–1480, 2004.[Abstract/Free Full Text]
  7. Folch J, Lees M, and Sloane Stanley GH. A simple method for the isolation and purification of total lipids from animal tissues. J Biol Chem 226: 497–509, 1957.[Free Full Text]
  8. Gibbons GF. A comparison of in-vitro models to study hepatic lipid and lipoprotein metabolism. Curr Opin Lipidol 5: 191–199, 1994.[Medline]
  9. Hellerstein MK and Neese RA. Mass isotopomer distribution analysis at eight years: theoretical, analytic, and experimental considerations. Am J Physiol Endocrinol Metab 276: E1146–E1170, 1999.[Abstract/Free Full Text]
  10. Hertzel AV, Bennaars-Eiden A, and Bernlohr DA. Increased lipolysis in transgenic animals overexpressing the epithelial fatty acid binding protein in adipose cells. J Lipid Res 43: 2105–2111, 2002.[Abstract/Free Full Text]
  11. Hertzel AV and Bernlohr DA. The mammalian fatty acid-binding protein multigene family: molecular and genetic insights into function. Trends Endocrinol Metab 11: 175–180, 2000.[CrossRef][ISI][Medline]
  12. Hotamisligil GS. Molecular mechanisms of insulin resistance and the role of the adipocyte. Int J Obes Relat Metab Disord 24: S23–S27, 2000.[CrossRef][Medline]
  13. Hotamsligil GS, Budavari A, Murray D, and Spiegelman BM. Reduced tyrosine kinase activity of the insulin receptor in obesity-diabetes. J Clin Invest 94: 1543–1549, 1994.[ISI][Medline]
  14. Hotamisligil GS, Johnson RS, Distel RJ, Ellis R, Papaioannou VE, and Spiegelman BM. Uncoupling of obesity from insulin resistance through a targeted mutation in aP2, the adipocyte fatty acid binding protein. Science 274: 1377–1379, 1996.[Abstract/Free Full Text]
  15. Illera JC, Gonzalez Gil A, Silvan G, and Illera M. The effects of different anaesthetic treatments on the adreno-cortical functions and glucose levels in NZW rabbits. J Physiol Biochem 56: 329–336, 2000.[ISI][Medline]
  16. Jequier E, Acheson K, and Schutz Y. Assessment of energy expenditure and fuel utilization in man. Annu Rev Nutr 7: 187–208, 1987.[CrossRef][ISI][Medline]
  17. Ohta A, Mayo MC, Kramer N, and Lands WEM. Rapid analysis of fatty acids in plasma lipids. Lipids 25: 742–747, 1990.[ISI][Medline]
  18. Okar DA, Manzano A, Navarro-Sabate A, Riera L, Bartrons R, and Lange AJ. PFK-2/FBPase-2: maker and breaker of the essential biofactor fructose-2,6-bisphosphate. Trends Biol Sci 26: 30–35, 2001.[CrossRef]
  19. Parks EJ, German JB, Davis PA, Frankel EN, Kappagoda CT, Rutledge JC, Hyson DA, and Schneeman BO. Reduced susceptibility of LDL from patients participating in an intensive atherosclerosis treatment program. Am J Clin Nutr 68: 778–785, 1998.[Abstract]
  20. Parks EJ, Krauss RM, Christiansen MP, Neese RA, and Hellerstein MK. Effects of a low-fat, high-carbohydrate diet on VLDL-triglyceride assembly, production and clearance. J Clin Invest 104: 1087–1096, 1999.[Abstract/Free Full Text]
  21. Parks EJ, Schneider TL, and Baar RA. Meal-feeding studies in mice: effects of different diets on blood lipids and energy expenditure. Comp Med. In press.
  22. Parks EJ, Schneider TL, and Baar RA. Metabolic studies in mice: effects of different diets and feeding regimens (Abstract). FASEB J 18: A867, 2004.
  23. Ren JM, Marshall BA, Mueckler MM, McCaleb M, Amatruda JM, and Shulman GI. Overexpression of Glut4 protein in muscle increases basal and insulin-stimulated whole body glucose disposal in conscious mice. J Clin Invest 95: 429–432, 1995.[ISI][Medline]
  24. Scheja L, Makowski L, Uysal KT, Wiesbrock SM, Shimshek DR, Meyers DS, Morgan M, Parker RA, and Hotamisligil GS. Altered insulin secretion associated with reduced lipolytic efficiency in aP2–/– mice. Diabetes 48: 1987–1994, 1999.[Abstract]
  25. Shaughnessey S, Smith E, Sarala K, Storch J, and Fried S. Adipocyte metabolism in adipocyte fatty acid binding protein knockout (aP2–/–) mice after short-term high-fat feeding: functional compensation by the keratinocyte fatty acid binding protein. Diabetes 49: 904–911, 2000.[Abstract]
  26. Shen WJ, Sridhar K, Bernlohr DA, and Kraemer FB. Interaction of rat hormone-sensitive lipase with adipocyte lipid-binding protein. Proc Natl Acad Sci USA 96: 5528–5532, 1999.[Abstract/Free Full Text]
  27. Simpson MA, LiCata VJ, Coe NR, and Bernlohr DA. Biochemical and biophysical analysis of the intracellular lipid binding proteins of adipocytes. Mol Cell Biochem 192: 33–40, 1999.[CrossRef][ISI][Medline]
  28. Teusink B, Voshol PJ, Kahlmans VEH, Rensen PCN, Pijl H, Romijn JA, and Havekes LM. Contribution of fatty acids released from lipolysis of plasma triglycerides to total plasma fatty acid flux and tissue-specific fatty acid uptake. Diabetes 52: 614–620, 2003.[Abstract/Free Full Text]
  29. Uysal KT, Scheja L, Wiesbrock SM, Bonner-Weir S, and Hotamisligil. Improved glucose and lipid metabolism in genetically obese mice lacking aP2. Endocrinology 141: 3388–3396, 2000.[Abstract/Free Full Text]
  30. Wolfe RR. Radioactive and Stable Isotope Tracers in Biomedicine. New York: Wiley and Sons, 1992.
  31. Wu C, Okar DA, Stoeckman AK, Peng LJ, Herrera AH, Herrera JE, Towle HC, and Lange AJ. A potential role for fructose-2,6-bisphosphate in the stimulation of hepatic glucokinase gene expression. Endocrinology 145: 650–658, 2004.[Abstract/Free Full Text]