Influence of diet on the modeling of adipose tissue triglycerides during growth

Daniel Z. Brunengraber,3 Brendan J. McCabe,3 Takhar Kasumov,3 James C. Alexander,1 Visvanathan Chandramouli,2 and Stephen F. Previs3

Departments of 1Mathematics, 2Medicine, and 3Nutrition, Case Western Reserve University School of Medicine, Cleveland, Ohio 44106

Submitted 26 March 2003 ; accepted in final form 6 June 2003


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We have studied the accretion of lipids in growing mice. We measured the rates of synthesis and degradation of triglycerides in epididymal fat pads of mice maintained for 44 days on a low-fat, high-carbohydrate diet (I) or a high-fat, lowcarbohydrate diet (II). 2H2O was added to the drinking water for 14 days. Rates of incorporation/washout of 2H to/from C1 of triglyceride-glycerol showed that triglyceride synthesis was greater than triglyceride degradation (net triglyceride balance was ~2.5 times greater in II than in I). The data also show that the contribution of de novo lipogenesis to triglyceride-bound palmitate was ~3 times greater in I than in II. This was consistent with a greater relative intake of carbohydrate in I vs. II. The rates of incorporation and washout of newly synthesized (2H-labeled) palmitate into and from triglycerides were also measured. Those data suggested a remodeling of triglyceride-bound fatty acids. On measuring the profile of triglyceride-bound fatty acids, we observed a decrease in the relative abundance of triglyceride-bound palmitate and stearate and an increase in triglyceride-bound oleate and linoleate. This was observed in I and II. In summary, diet substantially affects the deposition and modeling of triglycerides in adipose tissue during growth. 2H2O can be used to examine the mechanisms responsible for the accumulation of triglycerides, e.g., factors that affect 1) triglyceride synthesis and degradation and 2) the source of fatty acids that are used in esterification.

deuterium; stable isotopes; lipid metabolism; pediatric obesity; insulin resistance


THERE IS AN ALARMING INCREASE in the number of reported cases of pediatric obesity (26, 32). Like obese adults, obese children present with metabolic abnormalities (e.g., impaired glucose tolerance, dyslipidemia) that are associated with an increased risk of developing a chronic disease (e.g., diabetes, cardiovascular disease) (8, 9, 21, 24, 32). A special concern regarding obese children is that many will remain obese as adults, which further increases the likelihood of developing a chronic disease. Presumably, the development of effective treatments for pediatric obesity will benefit from knowledge of how specific biochemical reactions drive lipid accumulation during growth.

We hypothesized that, during growth, the nature of one's caloric intake will affect the rate of, and the mechanism of, fat accumulation. For example, as the ratio of dietary fat to carbohydrate increases, the rate of triglyceride accretion will increase. Also, the relative amount of carbohydrate in the diet will affect the relative contribution of de novo lipogenesis to the pool of triglyceride-bound fatty acids. We developed the use of 2H2O to measure the rates of the reactions involved in triglyceride accumulation (e.g., triglyceride synthesis and triglyceride degradation and the contribution of de novo lipogenesis to the pool of triglyceride-bound fatty acids). This approach also allows us to determine the rate of remodeling of fatty acids that are bound to triglycerides.

Application of the isotope labeling procedures (i.e., the chemical methods) required that we formulate a mathematical model to calculate the rates of triglyceride synthesis and degradation during non-steady-state metabolic conditions (e.g., during periods of net accumulation or loss). The need for this model is best explained using a simple example. Briefly, rates of triglyceride synthesis and degradation are determined according to the following rationale. Subjects are maintained on 2H2O for several days. During that time, triglycerides in adipose tissue become labeled. The rate of synthesis is determined from the rate of incorporation of 2H into tissue triglycerides. 2H is then eliminated from body water. The rate of triglyceride degradation is determined by measuring the decay of labeled triglycerides. A problem arises, however, when the rate of degradation of triglycerides is calculated via the decrease in labeling. This occurs because triglycerides are continuously being synthesized. Because the newly synthesized triglycerides are unlabeled, the apparent isotope decay rate will overestimate the true decay rate to extent of dilution from the newly synthesized unlabeled molecules.

In this report, we show that 1) 2H2O can be used to determine how the ratio of dietary carbohydrate to fat (or other factors) affects the rates of the specific reactions involved in lipid accretion and 2) special care must be used when interpreting data that are collected during conditions of growth (i.e., non-steady-state metabolism), since substantial remodeling of adipose tissue triglycerides can occur. We found that, although there is net accumulation of fat during growth, both triglyceride synthesis and triglyceride degradation are highly active. Last, de novo lipogenesis appears to be very active during growth.


    MATERIALS AND METHODS
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 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 Development of the 2H2O...
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Chemical Supplies

Unless specified, all chemicals and reagents were purchased from Sigma-Aldrich. 2H2O (99.9 atom percent excess) and [2H5]glycerol (98.5 atom percent excess) were purchased from Isotec (Miamisburg, OH). Ion exchange resins and HPLC columns were purchased from Bio-Rad (Hercules, CA). Gas chromatography-mass spectrometry supplies were purchased from Agilent Technologies (Wilmington, DE). Enzymes were purchased from Roche (Indianapolis, IN).

Biological Experiments

Theoretical. 2H from 2H2O is bound to carbon-1 (C1) of triose phosphates (i.e., dihydroxyacetone phosphate and glyceraldehyde 3-phosphate) during their formation and isomerization (25, 29) (Fig. 1). Reduction of dihydroxyacetone phosphate yields glycerol 3-phosphate; 2H remains bound to C1. Because glycerol 3-phosphate is used in the synthesis of triglycerides, we hypothesized that triglyceride synthesis could be quantified via the 2H labeling of triglyceride-glycerol. After the washout of 2H from body water, the rate of triglyceride degradation could be calculated by measuring the breakdown of 2H-labeled triglyceride-glycerol. It is well known that 2H from 2H2O is incorporated into fatty acids during fatty acid synthesis (1, 10, 18, 19). Thus the contribution of de novo lipogenesis to lipid accumulation could be determined by measuring the labeling of triglyceride-bound fatty acids. Last, the rate of remodeling of triglycerides could be determined by measuring the rates of incorporation, and removal, of newly synthesized (i.e., 2H-labeled) fatty acids into, and from, triglycerides.



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Fig. 1. Incorporation/washout of 2H (D) to/from triglycerides. In the presence of 2H2O(D2O), 2H is incorporated into the glycerol moiety of triglycerides. During de novo lipogenesis, 2H is incorporated into newly synthesized fatty acids. Rates and mechanisms of triglyceride synthesis can be determined by measuring the incorporation of 2H into the various parts of the triglyceride molecule. After the washout of 2H from body water, rates of triglyceride breakdown can be determined by measuring the decay of 2H-labeled triglycerides.

 

Determination of triglyceride synthesis and degradation. Male C57BL/6J mice (~5 wk old) were purchased from Jackson Laboratories (Bar Harbor, ME). On arrival, the mice were randomized to two groups. Each group was given free access to either diet I or diet II, purchased from Research Diets (New Brunswick, NJ; diet I, no. D12450 [GenBank] B, kcal distribution = 10% fat, 70% carbohydrate, and 20% protein; diet II, no. D12451 [GenBank] , kcal distribution = 45% fat, 35% carbohydrate, and 20% protein). After 10 days, the mice were given a single intraperitoneal injection of 2H2O-saline (16.25 µl/g body wt, 0.9 g NaCl in 1,000 ml 99% 2H2O; assuming that body water accounts for 65% body wt, this should establish 2.5% 2H enrichment of body water). Mice were returned to their cages and allowed free access to their respective diet and drinking water enriched to 5% 2H.

Mice were killed after 5, 10, or 14 days on 2H2O (n = 3 per day per group). On day 14, the remaining mice in each group were switched from 2H-labeled drinking water to tap water. Mice were killed on days 20 and 34 (n = 3 per day per group). At the time they were killed, blood and tissue samples were quick-frozen in liquid nitrogen and stored at –80°C.

Analytical Procedures

2H labeling of body water. The 2H labeling of body water was determined by exchange with acetone as described by Yang et al. (38), with minor modifications. First, it was not necessary to decrease the ionization energy below 70 eV to obtain a correct basal (M + 1)/M signal ratio in acetone. Presumably, the ionization effect(s) reported by Yang et al. do not occur on our instrument. Second, assays were performed using 40 µl of sample or standard, 2 µl of 10 N NaOH, and 4 µl of a 5% (vol/vol) solution of acetone in acetonitrile.

The 2H labeling of acetone was determined using an Agilent 5973N-MSD equipped with an Agilent 6890 GC system. A DB17-MS capillary column (30 m x 0.25 mm x 0.25 µm) was used in all analyses. The temperature program was: 60°C initial, increase by 20°C/min to 100°C, increase by 50°C/min to 220°C, and hold for 1 min. The split ratio was 40:1 with a helium flow of 1 ml/min. Acetone elutes at ~1.5 min. The mass spectrometer was operated in the electron impact mode. Selective ion monitoring of mass-to-charge ratios (m/z) 58–60 was performed using a dwell time of 10 ms/ion.

2H labeling of triglyceride-bound glycerol. Total glycerides were extracted from frozen tissues and hydrolyzed in ethanol-KOH at 70°C for 3 h (16). H2O (2 ml) was added, and the solution was acidified to ~pH 1 by adding 6 N HCl. The pH was monitored by using a pH meter (Accumet 900; Fisher Scientific). Fatty acids were first extracted using diethyl ether (3x with an equal volume). The pH of the aqueous solution was then adjusted to ~7.0 (using 10 N NaOH), and free glycerol was recovered by passing the solution over an ion exchange column, made by layering an AG 50W-X8 resin (formate form) over an AG 1-X8 resin (hydrogen form). Glycerol was recovered by washing with H2O.

Although glycerol appears as a symmetrical molecule, it has biological asymmetry (5, 30, 34). That is, glycerokinase selectively phosphorylates C3. In this study, we determined the labeling of 2H that is bound to C1triglyceride-glycerol and C3triglyceride-glycerol by use of the hexamethylenetetramine (HMT) method, as previously described (17). Briefly, reaction of triglyceride-glycerol with periodate yields C1triglyceride-glycerol + C3triglyceride-glycerol as formaldehyde. Reaction of triglycerideglycerol with glycerokinase plus ATP yields glycerol 3-phosphate, which, on reaction with periodate, yields only C1triglyceride-glycerol as formaldehyde. In each case, formaldehyde is reacted with NH4OH to generate HMT, which is assayed by gas chromatography-mass spectrometry. Using this approach, we directly measured the labeling of 2H that is bound to C1 + C3 and C1 alone. The labeling of 2H that is bound to C3 was calculated as [(C1 + C3)/0.5] – C1. This calculation assumes that the two hydrogens bound to C1 and C3 are equally labeled. Note that the samples used in this analysis were obtained from mice that were maintained for <=14 days on 2H2O (i.e., mice that were killed on days 5, 10, and 14).

Rose and colleagues (25, 29) demonstrated that the reactions catalyzed by triose phosphate isomerase and aldolase are stereospecific. That is, although C1 of dihydroxyacetone phosphate is bound to two hydrogens, triose-phosphate isomerase specifically incorporates label to one hydrogen, whereas aldolase incorporates label to the opposite hydrogen. We followed the methods of Rieder and Rose (25) and Rose et al. (29) to determine the equilibration of label on each hydrogen bound to C1triglyceride-glycerol. Briefly, samples of triglyceride-glycerol from a particular mouse were split into two fractions (A and B). Fractions A and B were reacted with glycerokinase, ATP, glycerol-3-phosphate dehydrogenase, and NAD. However, fraction B was also reacted with triose phosphate isomerase. After 3 h at 37°C, excess NADH was added, and the reaction was run for another 20 min. Glycerol 3-phosphate was isolated using ion exchange chromatography and further purified by HPLC. HMT was generated from C1 and analyzed by gas chromatography-mass spectrometry. By means of this strategy, samples from fraction A include the labeling of both hydrogens bound to C1 of triglycerideglycerol, whereas the sample in fraction B contains only a single hydrogen bound to triglyceride-glycerol (i.e., that which was derived during the hydrolysis of fructose 1,6-bisphosphate by aldolase). Note that the samples used in this analysis were obtained from mice that were maintained on 2H2O for 14 days.

The 2H labeling of HMT was determined using an Agilent 5973N-MSD equipped with an Agilent 6890 GC system. A DB17-MS capillary column (30 m x 0.25 mm x 0.25 µm) was used in all analyses. The temperature program: 100°C initial, hold for 2 min, increase by 20°C/min to 220°C, and hold for 4 min. The split ratio was 15:1 with a helium flow of 1 ml/min. HMT elutes at ~6.1 min. The mass spectrometer was operated in the electron impact mode. Selective ion monitoring of m/z 140 and m/z 141 was performed using a dwell time of 10 ms/ion.

Concentration of triglyceride-bound glycerol. A known quantity of tissue was hydrolyzed and processed after a known amount of [2H5]glycerol was added. After isolation by ion exchange, glycerol was converted to its trimethylsilyl derivative. This was done by adding 75 µl of bis(trimethylsilyl)trifluoroacetamide plus 10% trimethylchlorosilane (Regis, Morton Grove, IL) and heating at 75°C for 30 min. The sample was then analyzed using gas chromatography-electron impact ionization mass spectrometry. The concentration of glycerol was determined from the ratio of m/z (205 + 206)/(205 + 206 + 208).

2H labeling and concentration profile(s) of triglyceridebound fatty acids. A known quantity of tissue was hydrolyzed and extracted after a known amount of heptadecanoic acid (17:0) was added. Fatty acids were derivatized using diazomethane. Briefly, diazomethane was prepared and used as previously described (4). Fatty acid methyl esters were formed by dissolving the extracted fatty acids in 50 µl of methanol and adding ~300 µl of ether-diazomethane. The sample was allowed to react at room temperature for 45 min. Excess solvent was removed by evaporating to dryness. The fatty acid methyl esters were then dissolved in 100 µl of chloroform and analyzed by gas chromatography-electron impact ionization mass spectrometry.

The 2H enrichment of palmitate was determined by monitoring m/z 270–272. Unless noted, the total labeling of palmitate [i.e., M1 + (2 x M2), where M1 and M2 refer to the percentage of excess palmitate molecules with one or two 2H atoms, respectively] is shown.

The concentration of palmitate (16:0), stearate (18:0), oleate (18:1), and linoleate (18:2) was determined by comparing the abundance of m/z 270–272, 298–300, 296, and 294, respectively, to that of heptadecanoate (17:0) m/z 284. To account for possible differences in the ionization efficiency of each fatty acid, the profile was compared against standards prepared by mixing known quantities of each fatty acid.

Mathematical modeling. The modeling of triglyceride synthesis and degradation is done in two steps. First the concentration of triglyceride-bound glycerol is modeled, and then the concentration of triglyceride-bound fatty acids is modeled. The first step allows us to determine the net accumulation of triglycerides, whereas the second step allows us to determine the contribution of de novo lipogenesis vs. diet to the pool of triglyceride-bound fatty acids and whether there is a remodeling of triglyceride-bound fatty acids.

The incorporation of 2H from water into triglycerides in adipose tissue is modeled using a single-compartment model, assuming that the labeling of plasma water reflects that of water in adipose tissue. The time-dependent labeling of 2Hin water is c(t). 2H is eliminated from plasma at a rate c'. The parameters of basic interest, the rates of triglyceride synthesis and degradation (S and D, respectively, expressed in mg/day), are estimated from data using nonlinear least squares fitting.

Because the mass of triglycerides (expressed in mg) increases linearly with time (m, expressed in mg/day), the total amount of labeled triglycerides at time t satisfies the differential equation

where h equals the 2H labeling of triglycerides in adipose tissue (expressed in mg 2H). The parameter c(t) is essentially constant while mice are maintained on 2H-labeled drinking water, yet it decays exponentially (at rate c') when mice are switched to tap water. The parameter c' is estimated from the data.

Differential equations can then be solved in closed form for "before" and "after" removal of 2H-labeled drinking water. Those equations are

where c0 = c(0) and v0 = v(0) represent the 2H labeling (expressed in percent enrichment) of body water and triglyceride-glycerol, respectively, at time t = 0 (the time when 2H2O is withdrawn), and {Gamma}(a,z) is the incomplete gamma function

The model was solved by using a linear least squares fit of the data regarding the mass of triglycerides in adipose tissue.

The only remaining parameters that require estimation are the rate of synthesis (S) and the rate of degradation (D, which equals S – m) and v0. A Levenberg-Marquardt nonlinear fit (equal weights on all data points) is done for these parameters.

Once the parameters of triglyceride synthesis and breakdown are determined, the synthesis of fatty acids is then modeled. Here, an additional compartment is needed, since fatty acids can be created de novo or incorporated from the diet. Accordingly, a new parameter p, the proportion of de novo production, is introduced

The remaining formulas are similar to those described above, and a similar nonlinear least squares fit is made to the data.

Calculations. Unless noted, data are expressed as means ± SE. Statistical differences were calculated using a t-test.


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Figure 2 shows the body weight and epididymal fat pad weight of growing mice maintained on different diets. Body weight was similar in the two groups until the last time point (Fig. 2A). However, the weights of the epididymal fat pads (2 pads were pooled from each mouse) were different at all time points (Fig. 2B).



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Fig. 2. Change in body weight and epididymal fat pads in growing mice. Mice were maintained on a low-fat, high-carbohydrate diet (diet I, {bullet}) or a high-fat, low-carbohydrate diet (diet II, {circ}). At the time of death, the body weight (A) and the wet weight of total epididymal fat pads (B, 2 fat pads were pooled from each mouse) were determined. There was no difference in body weight until the last day. The weight of the fat pads was different at all points in time (P < 0.05).

 

Figure 3 shows the change in the total amount of triglyceride-bound glycerol and triglyceride-bound fatty acids from the epididymal fat pads. Regression analyses showed a linear increase in the quantity of triglyceride-glycerol (diet I: y = 4.3x + 179, r2 = 0.996; diet II: y = 11.4x + 271, r2 = 0.992) and triglyceridefatty acids (diet I: y = 15.8x + 537, r2 = 0.971; diet II: y = 36.6x + 766, r2 = 0.992).



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Fig. 3. Change in total triglyceride-bound glycerol (A) and triglyceride-bound fatty acids (B) in epididymal fat pads from growing mice. Total amount of triglyceride was determined by hydrolyzing a known amount of tissue and measuring the quantity of glycerol and fatty acids (i.e., palmitate, stearate, oleate and linoleate). There was a linear increase in the total amount of glycerol and fatty acids. The total quantity of fatty acids was ~3 times that of glycerol; this was observed in both groups. Diet significantly affected the amount of glycerol and fatty acids at all points.

 

2H2O was used to quantify the rates of synthesis and breakdown of lipids. Figure 4 shows that the labeling of body water remained constant while mice were maintained on 2H-labeled drinking water. The 2H enrichment of plasma water was ~50% that of the drinking water. The apparent exchange/dilution of label is in agreement with other reports (1) and presumably occurs via respiration and digestion of food. Between days 14 and 20, 2H was washed out of body water by changing the drinking water from 2H2O to H2O. On day 20, the 2H labeling of plasma water decreased to ~0.45% 2H. The calculated t1/2 of body water is ~2.5 days. This is consistent with experiments in which the elimination of body water was measured over the course of 1 wk (4).



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Fig. 4. Labeling of body water in growing mice. MPE, mole percent excess. A steady-state labeling of body water was maintained by using a primed infusion of 2H2O. The prime was achieved by administering an intraperitoneal injection; the constant infusion was achieved by allowing free access to 2H-labeled drinking water. 2H was washed out from body water by switching mice to tap water.

 

Figure 5 shows the 2H labeling of triglyceride-glycerol and triglyceride-palmitate. Data are shown as the total labeling of 2H bound to C1 of triglyceride-glycerol and the total labeling of triglyceride-bound palmitate. While the mice were maintained on 2H2O there was continuous labeling of triglycerides; after switching to H2O, the labeling in triglycerides decreased.



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Fig. 5. Labeling of triglyceride-bound glycerol (A) and triglyceride-bound palmitate (B) isolated from epididymal fat pads. While mice were maintained on 2H2O, there was a constant increase in the enrichment of triglyceride-bound glycerol and triglyceride-bound palmitate. When 2H was washed out of body water, the 2H labeling in triglycerides decreased. Data are shown as total labeling of 2H bound to C1 of triglyceride-bound glycerol and the total labeling of triglyceride-bound palmitate [i.e., M1 + (2 x M2)].

 

Figure 6 shows a fit of the data in Figs. 3, 4, 5 after application of the equations presented in MATERIALS AND METHODS. There was a good fit between the observed data and the computed parameters.



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Fig. 6. Best fit of the 2H-labeling of triglyceride-bound glycerol (A) and triglyceride-bound fatty acids (B). Rates of triglyceride synthesis and breakdown were determined by fitting the glycerol and fatty acid (i.e., palmitate) data in Figs. 3, 4, 5 and using the equations described in MATERIALS AND METHODS. Shown is the best fit of the data that were obtained for each group.

 

Table 1 contains the rates of triglyceride turnover (i.e., rates of triglyceride synthesis and degradation, contribution of de novo lipogenesis, and rates of incorporation and removal of newly synthesized palmitate into and from triglycerides). Regardless of diet, the rates of triglyceride synthesis were greater than the rates of triglyceride degradation. The accumulation of triglycerides is consistent with the fact that growing mice were studied. The net accumulation of triglycerides was ~2.5 times greater on diet II compared with diet I. The contribution of de novo lipogenesis was about three times greater on diet I than on diet II. Regardless of diet, there was net accumulation of palmitate. The accumulation of palmitate was ~7.5 times greater on diet I than on diet II.


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Table 1. Rates of lipid synthesis and breakdown

 

The relative distribution of triglyceride-bound fatty acids was measured. Although there was an accumulation of total triglyceride-bound fatty acids (Fig. 3), Fig. 7 shows that there was an extensive remodeling of the triglyceride-bound fatty acids in both groups. Namely, the relative amount of saturated fatty acids decreased (e.g., diet I: 16:0, 0.39 ± 0.05 day 5 to 0.29 ± 0.02 day 34, P < 0.01 and 18:0, 0.10 ± 0.01 day 5 to 0.06 ± 0.01 day 34, P < 0.01; diet II: 16:0, 0.29 ± 0.02 day 5 to 0.21 ± 0.01 day 34, P < 0.01 and 18:0, 0.09 ± 0.01 day 5 to 0.06 ± 0.01 day 34, P < 0.01), and the relative amount of unsaturated fatty acids increased (e.g., diet I: 18:1, 0.27 ± 0.03 day 5 to 0.37 ± 0.07 day 34, P < 0.05 and diet II: 18:1, 0.32 ± 0.02 day 5 to 0.41 ± 0.02 day 34, P < 0.01).



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Fig. 7. Relative distribution of triglyceride-bound fatty acids. In both groups, there was a decrease in the relative amounts of palmitate and stearate (saturated fatty acids) and an increase in the relative amounts of oleate and linoleate (unsaturated fatty acids). A: diet I (low-fat, high-carbohydrate), filled bars, day 5; gray bars, day 34. B: diet II (high-fat, low-carbohydrate), open bars, day 5; gray bars, day 34.

 


    DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 Development of the 2H2O...
 DISCLOSURES
 REFERENCES
 
Lipid accretion is an important component of normal growth. For example, between the ages of ~5 and 20 yr, healthy individuals accumulate comparable amounts of protein and fat. Unfortunately, epidemiological data show a dramatic increase in the number of obese children (26). We suspect that determining how specific biochemical reactions affect lipid accretion during growth is an important first step in the development of treatments for preventing and/or reversing pediatric obesity (and thereby reducing the risk of developing future disease).

We developed the use of 2H2O to study the influence of diet on the biochemical basis of triglyceride accumulation during growth. First, we discuss the development of this method. Second, we discuss the implications of our data on future studies.


    Development of the 2H2O Method
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 Development of the 2H2O...
 DISCLOSURES
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A prerequisite for using 2H2O was to establish and maintain a steady-state labeling of body water (the precursor pool). This is easily achieved by administering a priming bolus of 2H2O and then allowing free access to 2H-labeled drinking water (Figs. 1 and 4). As shown, 2H is readily washed out of body water by changing the drinking water from 2H2O to tap water. Therefore, it is possible to calculate multiple parameters related to triglyceride turnover by measuring the incorporation/washout of 2H to/from lipids.

Rates of triglyceride synthesis are calculated after determining the precursor-to-product labeling ratio (7). Although determination of the precursor labeling is straightforward (i.e., measure the labeling of body water), we questioned the correct "product labeling". For example, we (17) previously reported that the ratio of 2H bound to C3 vs. C1 of VLDLtriglyceride-glycerol is ~0.93; however, the ratio of 2H bound to C3 vs. C1 of blood glycerol is ~0.72 (Ref. 17 and Tables 2 and 3 therein, respectively). Those data suggest that the labeling of the hydrogens bound to different carbons of triglyceride-glycerol may not be equivalently labeled. In the present study, we observed that the labeling of 2H that is bound to C3 of triglyceride-glycerol obtained from adipose tissue of mice is ~75% that of C1. This was true whether we examined samples that were obtained from mice maintained on diet I or diet II (not shown). Presumably, incomplete equilibration of 2H on the different carbons of glycerol reflects the fact that glycerol 3-phosphate, used in the synthesis of triglycerides, can be derived from glucose and/or phosphoenolpyruvate carboxykinase (PEPCK; i.e., glyceroneogenesis). For example, phosphoglucoisomerase incorporates 2H on C1glucose 6-phosphate to yield [1-2H]fructose 6-phosphate. Conversion to [1-2H]fructose 1,6-bisphosphate and hydrolysis yields two triose phosphates; during hydrolysis and equilibration, 2H is incorporated on C1dihydroxyacetone phosphate and C1glyceraldehyde 3-phosphate. Reduction to glycerol 3-phosphate yields two molecules of glycerol 3-phosphate with 2H on C1; only one of those molecules also has 2H on C3 (i.e., that which was bound to C1 of fructose 1,6-bisphosphate). The conversion of substrates to glycerol 3-phosphate via PEPCK to glycerol 3-phosphate could also preferentially label C1triglyceride-glycerol vs. C3triglyceride-glycerol (6).

Rieder and Rose (25) and Rose et al. (29) demonstrated that the two hydrogens bound to C1 of dihydroxyacetone phosphate derive from different reactions. Therefore, after establishing that the labeling of C1triglyceride-glycerol reflects total synthesis (above), we examined whether the hydrogens bound to C1triglyceride-glycerol are equally labeled. We analyzed samples obtained from mice that were maintained for 14 days on 2H2O. The total labeling of C1triglyceride-glycerol was ~2.21% (diet I) and ~2.50% (diet II; Fig. 5). Samples treated with triose phosphate isomerase had an enrichment of ~1.03% (diet I) and ~1.17% (diet II). Thus, regardless of diet, the two hydrogens of C1triglyceride-glycerol reached ~88% equilibrium.

On the basis of the observations described above (i.e., the different labeling on C1triglyceride-glycerol vs. C3triglyceride-glycerol and the equilibration of 2H between the two hydrogens of C1triglyceride-glycerol), we concluded that the correct product labeling is the labeling of 2H bound to C1triglyceride-glycerol divided by 2.

Application of the 2H2O Method

There is uncertainty regarding how the ratio of dietary fat to carbohydrate affects the development of obesity (14, 36). We hypothesized that the relative distribution of one's caloric intake would substantially affect the pathway(s) of lipid accretion during growth. 2H2O was used to test our hypothesis. Although the predominant difference between the diets was the ratio of dietary fat to carbohydrate, the caloric intake was slightly greater in mice that were fed a high-fat, low-carbohydrate diet (diet II) compared with those fed a low-fat, high-carbohydrate diet (diet I) (~10.5 vs. ~9.6 kcal · mouse1 · day1, respectively, P < 0.075).

We observed that, during growth, the rate of accumulation of triglycerides in epididymal fat pads (i.e., the balance between the rates of synthesis and breakdown) is related to the intake of dietary fat (i.e., 4.4 µmol/day on diet I vs. 10.9 µmol/day on diet II; Table 1). Although our overall observation is consistent with the literature [i.e., percent dietary fat affects rate of lipid accretion (35)], these data are the first direct measurements of the rates of triglyceride synthesis and degradation in vivo. More importantly, the data show that, despite the very active triglyceride synthesis, triglyceride degradation also occurs at a considerable rate (i.e., degradation occurs at ~50% the rate of synthesis). Our findings agree with the original work of Schoenheimer and Rittenberg (31), who concluded that ~50% of dietary fat moves through adipose tissue each day. This also supports the hypothesis put forth by Frayn (11), that adipose tissue may act as a buffer for daily lipid flux.

Using 2H2O allowed us to examine how the relative amount of dietary carbohydrate affects the relative contribution of lipogenesis (1, 10, 18, 19, 37). This was done by dividing the total enrichment of palmitate [i.e., M1 + (2 x M2), where M1 and M2 refer to the percentage of excess palmitate molecules with one or two 2H atoms, respectively] by the enrichment of body water times 22 (Fig. 6) [i.e., the average number of hydrogens incorporated during synthesis (1, 10)]. We found that the contribution of de novo lipogenesis to the pool of triglyceride-bound palmitate was about three times greater in mice maintained on diet I than on diet II (Table 1). Similar data were obtained by measuring the labeling of stearate (not shown). The observation is consistent with the fact that mice on diet I consume more carbohydrate than mice on diet II. One intriguing observation is that the type of carbohydrate also varied between the diets. For example, the ratio of complex to simple carbohydrates was greater in diet I than in diet II. Although the intake of simple carbohydrates is usually more closely associated with increased lipogenesis (14, 36), our data show that de novo lipogenesis is very active during growth despite the type of carbohydrate(s) being consumed (Table 1).

Finally, we were able to determine the rate of remodeling of triglyceride-bound fatty acids. This was done by measuring the rates of incorporation and removal of newly synthesized (i.e., 2H-labeled) palmitate into and from triglycerides. Initial inspection of the labeling data suggested that there must be a major remodeling. For example, if triglycerides are accumulating at 4.4 µmol/day (e.g., Table 1, diet I), then 13.2 µmol of fatty acids must be incorporated per day. Because palmitate typically accounts for a large percentage (e.g., ~35%) of the total fatty acids found in adipose tissue, one should expect that palmitate would accumulate at ~4.6 µmol/day (i.e., 13.2 x 0.35). We found that the net palmitate flux was substantially less than what was expected on the basis of the net triglyceride flux (e.g., net palmitate flux was only 0.8 µmol/day; Table 1, diet I). This was observed in both groups (Table 1). On measuring the distribution of the triglyceride-bound fatty acids, we found that the relative amount of palmitate decreased over time (Fig. 7). These observations are consistent with the original data presented by Hirsch (15). Namely, triglycerides in adipose tissue are in dynamic exchange with the diet; the remodeling that we observed reflects the fact that the fatty acid content of diet that the mice were raised on (at Jackson Laboratories) is not the same as what we fed them (purchased from Research Diets, New Brunswick, NJ). Also, these data suggest that tracing the turnover of triglycerides via glycerol labeling has an advantage over using a labeled fatty acid (28), since various fatty acids have different fates (3, 33). Therefore, measuring the incorporation of one fatty acid into triglyceride may not accurately reflect the fate of other fatty acids.

In summary, our data show that diet influences 1) the rate of triglyceride accumulation and 2) the pathway(s) that affects lipid accretion. Also, although growth is characterized by net accumulation of lipid, lipid breakdown is very active, and substantial remodeling of triglycerides can occur. Because future experiments are aimed at studying the regulation of triglyceride turnover in adipose tissue, we reviewed the literature to identify parameters that could affect these measurements. In particular, several studies have examined compartmentation/heterogeneity of adipose tissue. We discuss the significance of those reports below.

First, Rodbell (27) demonstrated that adipose tissue is contaminated with vascular tissue. If unaccounted for, this may affect measurements of protein and DNA content of fat tissue. However, because vascular tissue contains virtually no triglyceride (27), quantitation of glyceride kinetics in adipose depots should be possible after a tissue biopsy without purification from vascular tissue.

Second, the lipid composition of a tissue can affect measurements of turnover. However, because ~90% of the glycerides found in adipose tissue are triglycerides, it does not seem that one needs to separate various glyceride species (e.g., triglyceride, diglyceride, phospholipids). Also, pulse-chase studies have shown that the small pool of diglycerides is in rapid equilibrium with the large pool of triglycerides (39).

Third, Angel (2) studied the intracellular structural heterogeneity. This author demonstrated that >90% of all lipid in adipocytes is localized in the cytoplasmic pool. A minor part of the total pool (<10%) is associated with other compartments (e.g., mitochondria, microsomes, etc.).

Fourth, chemical heterogeneity has been described. This can occur within the triglyceride pool because of the positional specificity of the lipases (12, 22). For example, lipoprotein lipase, which hydrolyzes extracellular triglycerides before entry into adipocytes, has different specificity from hormone-sensitive lipase, which hydrolyzes intracellular triglycerides before release from adipocytes.

Finally, vascularization can affect heterogeneity of adipose tissue (13). For example, investigators have observed variations in the number of, and area of, blood vessels within a single adipose tissue. Presumably, the greater the degree of perfusion, the greater the metabolic activity.

In summary, 2H2O can be used to measure the total rates of triglyceride synthesis and degradation in adipose tissue in vivo. 2H2O is uniquely suited for investigations in free-living conditions because catheterization is not required. Although the dose of 2H2O used in our experiments is safe for rodents (23), to administer less 2H and not compromise analytical sensitivity we recently developed an alternative approach to processing gas chromatography-mass spectrometry data (20).

In conclusion, the ratio of dietary fat to carbohydrate affects the rates of, and the mechanisms of, triglyceride accumulation in adipose tissue. Our data show that triglycerides in adipose tissue are in a dynamic state during growth. Although future studies by our laboratory are aimed at studying how different interventions may affect the biochemical basis of lipid accumulation during growth, 2H2O is also ideal for studying how to affect lipid kinetics in other conditions (e.g., obese or lipoatrophic subjects).


    DISCLOSURES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 Development of the 2H2O...
 DISCLOSURES
 REFERENCES
 
This project was supported by funds from the Mt. Sinai Health Care Foundation of Cleveland, the Diabetes Association of Greater Cleveland (Grant no. 449–01), and Merck Pharmaceuticals. V. Chandramouli was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-14507.


    ACKNOWLEDGMENTS
 
We thank Dr. Timothy Kern for generously donating mice that were used in several pilot studies and Dr. Vernon Anderson for helpful discussions regarding the stereospecificity of 2H incorporation into triose phosphates via enzymatic reactions.


    FOOTNOTES
 

Address for reprint requests and other correspondence: S. F. Previs, Dept. of Nutrition, Dental Bldg., Rm. 201, Case Western Reserve Univ. School of Medicine, 10900 Euclid Ave., Cleveland, OH 44106–4906 (sxp29{at}po.cwru.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.


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