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The Journal of Clinical Endocrinology & Metabolism Vol. 86, No. 2 797-803
Copyright © 2001 by The Endocrine Society

Original Studies

Ex Vivo Measurement of Lipoprotein Lipase-Dependent Very Low Density Lipoprotein (VLDL)-Triglyceride Hydrolysis in Human VLDL: An Alternative to the Postheparin Assay of Lipoprotein Lipase Activity?1

Valerie Pruneta, Delphine Autran, Gabriel Ponsin, Christophe Marcais, Laurence Duvillard, Bruno Verges, Francois Berthezene and Philippe Moulin

Laboratoire de Métabolisme des Lipides (V.P., D.A., G.P., P.M.) CNRS ESA 5014 and Service d’Endocrinologie et des Maladies de la Nutrition (P.M., F.B.), Hôpital de l’Antiquaille, Lyon; Laboratoire de Biochimie (C.M.), Centre Hospitalier Lyon-Sud, Lyon; Service d’Endocrinologie-Diabétologie-Maladies Métaboliques, INSERM U498 (L.D., B.V.), Centre Hospitalier Universitaire Dijon, Dijon, France

Address correspondence and requests for reprints to: Valérie Pruneta, Laboratoire de Métabolisme des Lipides, CNRS ESA 5014, Hôpital de l’Antiquaille, 1 rue de l’Antiquaille, 69005 Lyon, France.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The plasma lipolysis of triglyceride (TG)-rich lipoproteins is mainly due to the activity of lipoprotein lipase (LPL). Albeit important for our analysis of certain physiopathological situations, the determination of the magnitude of LPL-dependent lipolysis is not easy to perform. This essentially results from the binding of LPL to the luminal surface of vascular endothelium. The measurements of the whole putative LPL activity have been achieved after injection of heparin, a procedure that releases LPL from endothelium. However, the physiopathological relevance of this postheparin lipolysis assay (PHLA) remains questionable because it has never been demonstrated that the bulk of endothelium-bound LPL was active.

It has been recently shown that a small part of LPL is associated to circulating lipoproteins in nonheparinized plasma, raising the possibility that the lipolysis mediated by this circulating LPL might reflect the overall LPL-dependent TG hydrolysis in plasma. To address this question, we developed a new lipolysis assay in which the very low density lipoprotein (VLDL)-bound LPL-dependent VLDL-TG hydrolysis (LVTH) was directly determined through the measurement of nonesterified fatty acid (NEFA) release during in vitro incubations. LVTH measurements were performed in control subjects, in type 2 diabetics, and in either heterozygous or homozygous LPL-deficient patients. In the latter group, LVTH values were extremely low. Those of heterozygous patients and of diabetics were similarly decreased by about 40% with respect to control group. Plasma TG concentrations exhibited an inverse relationship with LVTH level. In a subgroup of subjects, LVTH and PHLA were positively correlated and the inverse correlation of LVTH with plasma or VLDL-TG concentration was stronger than that obtained with PHLA. To further study the validity of this new assay, we measured LVTH in nine subjects who were studied for their catabolism of VLDL labeled with stable isotope. No relation was observed between the direct hepatic removal of VLDL and LVTH, whereas the latter was strikingly correlated with the rate of conversion of VLDL to intermediary density lipoprotein.

Collective consideration of these findings strongly suggests that LVTH is a physiologically relevant index which could advantageously replace the measurements of PHLA in numerous physiopathological situations.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
PLASMA TRIGLYCERIDES (TG) are transported by circulating chylomicrons (CM) and very low density lipoprotein (VLDL) particles, and they are hydrolyzed through the action of lipoprotein lipase (LPL; EC 3.1.1.34) (1, 2). This reaction results in the production of nonesterified fatty acids (NEFA) available for tissue uptake and subsequent utilization or storage. Thus, in addition to its interest for our overall understanding of TG metabolism, the measurement of LPL-mediated lipolysis may be of critical importance in certain physiopathological situations. LPL is synthesized in parenchymal cells and transported across the endothelium where it binds to heparan sulfate proteoglycans (HSPG) at the luminal surface of vascular endothelium (3). The in vivo magnitude of LPL-dependent lipolysis is not easy to evaluate. Due to a strong posttranscriptional regulation, the measurement of LPL gene expression is poorly informative (4). It is thus necessary to perform a direct assay of the enzyme catalytic activity. Because the bulk of LPL is bound to the vascular endothelium, its activity has been commonly assayed in blood samples collected after iv injection of heparin, a process that releases LPL from HSPG (5, 6, 7). However, this assay has several drawbacks. The injection of heparin to control subjects raises ethical problems and prevents the possibility of serial measurements. From a technical viewpoint, heparin also releases hepatic lipase (HL) from hepatic sinusoid capillaries, which results in a significant contribution of HL to the total postheparin lipolytic activity (PHLA) (8, 9). To specifically measure plasma LPL activity, it is thus necessary either to substract HL activity from total lipolytic activity (10, 11), or to selectively inhibit HL using a specific antiserum (12, 13), or to separate HL from LPL by affinity chromatography on a heparin-Sepharose column (9). Whichever the procedure used, it necessarily introduces a supplementary variability that decreases the accuracy of the assay.

In spite of its limitations, PHLA assay has been used to determine the total LPL activity theoretically available for plasma lipolysis, although it has never been demonstrated that all of the HSPG-bound LPL is necessarily active. In fact, it has recently been shown that LPL can dissociate from the endothelial surface and move into the blood (14, 15). It is now clear that in nonheparinized plasma, a small part of LPL is associated with circulating lipoproteins and especially to apoB-containing lipoproteins (16, 17, 18, 19, 20). The question as to whether circulating LPL was catalytically active has been controversial (18, 20). Although several reports have now shown that circulating LPL was active (17, 20), the level of preheparin LPL activity was not correlated to PHLA (21, 22, 23, 24). This raises the important question of the physiopathological relevance of the circulating LPL activity. Preheparin LPL measurements were found previously decreased in plasma of patients with type IV or type IIb dyslipidemia (25). Although correlation between VLDL kinetics and circulating LPL was not provided in these reports and no study was performed in LPL genetic deficiency, the data suggest that plasma LPL activity might reflect the LPL-mediated TG lipolysis in pathological situations.

In the present study, we describe a new lipolysis assay in which the VLDL-bound LPL-dependent VLDL-TG hydrolysis (LVTH) is directly determined through the measurement of nonesterified fatty acid (NEFA) release during in vitro incubations. In addition, we provide an evaluation of the physiopathological relevance of this new assay on the basis of two studies. In the first, we have analyzed the levels of LVTH in different groups of subjects, including controls, type 2 diabetics, and LPL-deficient patients. In the second study, we have related LVTH levels to the dynamics of VLDL catabolism, as determined by mass spectrometry using L-[1-13C] leucine-labeled apolipoprotein B as the marker.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
All experiments were carried out using a buffer (TSE) containing 150 mmol/L NaCl, 10 mmol/L Tris-HCl, 2 mmol/L EDTA, and 0.02% NaN3, pH 7.4.

Subjects and samples

Blood samples were obtained after an overnight fast, from 20 informed and consenting healthy subjects (12 women and 8 men), from 13 type 2 diabetic patients, and from 10 patients with genetic LPL deficiency (5 homozygous, 5 heterozygous). The diabetics and the obligate heterozygous exhibited a mild hypertriglyceridemia and hypoalphalipoproteinemia (Table 1Go). The study was conducted in agreement with the recommendations of our local ethical committees. The mutations in the LPL gene were characterized by sequencing after screening by SSCP. Two patients had a G->A mutation at the acceptor splice site of intron 1, one had the Gly 188 Glu mutation, one a Y 288 ter mutation, and one a 192 ter mutation. Blood samples for the determination of plasma lipid and lipoprotein concentrations were collected into tubes containing Na2-EDTA; separate aliquots (10 mL) for LVTH measurements were drawn in prechilled tubes kept on ice. Plasma were then separated by low-speed centrifugation at 4 C within 1 h to avoid in vitro lipolysis (26). When subsequent analysis were not performed on the same day, plasma samples were immediately frozen at -80 C.


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  		Table 1. Characteristics of subjects

 
Plasma lipoproteins

The major plasma lipoproteins [VLDL, low density lipoprotein (LDL), and high density lipoprotein (HDL)] were separated by a combination of preparative ultracentrifugation and precipitation of apoB-containing lipoproteins (27, 19), with the exception of homozygous LPL-deficient plasma. In the latter, chylomicrons were first removed after centrifugation of the samples at 35,000 x g for 1 h at 12 C, and then VLDL and LDL were separated by sequential ultracentrifugation as previously described (28). Plasma lipids were assayed using commercial kits [Roche Molecular Biochemicals (Mannheim, Germany) for cholesterol and TG, and Oxoid (Dardilly, France) for NEFA].

Determination of VLDL-TG hydrolysis

After filtration on a 0.22 µm-filter, plasma samples (0.6 mL) were fractionated by fast protein liquid chromatography (FPLC) at 4 C using a Superose 6 HR 10/30 column (Pharmacia LKB, Uppsala, Sweden) to separate lipoproteins in TSE buffer containing 10 IU/mL heparin for stabilization of LPL during the procedure. This procedure described by Zambon et al. (20) does not release LPL from VLDL. Chromatography was carried out with a flow rate of 0.3 mL/min under a pressure of 150 psi. Fractions of 0.3 mL were collected. The peak corresponding to VLDL (fractions 11–18) was concentrated using Centricon 30 and assayed for TG content. VLDL (0.3 µmol TG) were immediately incubated at 37 C in 0.5 mL of buffer, and lipolysis was monitored according to time. When desired, exogenous bovine LPL was added to LPL-deficient VLDL before incubation, as previously described (28). Blanks were obtained by incubations of samples in the presence of 2 mmol/L Paraoxon (E600) (Sigma, St. Louis, MO), which totally blocked LPL-mediated lipolysis (26), and they were substracted to the amounts of NEFA released during the incubations. As shown in Fig. 1Go, NEFA production increased linearily during the first hour of incubation before reaching a plateau. Routine measurements of LVTH were thus performed after 1 h of incubation. The results were then corrected for plasma VLDL-TG concentration and expressed as the amounts of NEFA released per mL of plasma and per hour. The interassay variability of these measurements was of 4.3%. To check residual HL activity, a goat antihuman HL polyclonal antibody was used at rising concentration as previously described (29). To determine the effect of freezing, plasma obtained from five healthy subjects were aliquoted and LVTH was measured before and after 2 weeks and 2 months of storage at -80 C. No significant change of LVTH level was found after 2 weeks of freezing (105.4 ± 16.7 vs. 109.9 ± 23.8 nmol NEFA/mL·h), whereas a mild 8.7% decrease was observed after 2 months (P = 0.0455).



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  		Figure 1. Time-course of VLDL-bound LPL-dependent VLDL-TG hydrolysis (LVTH). The amounts of NEFA resulting from TG hydrolysis were measured during incubations at 37 C of VLDL isolated from control plasma. The data were substracted for blanks that were determined in samples containing 2 mmol/L paraoxon to block LPL. Values are shown as the mean ± SEM of five independent measurements.

 
Determination of PHLA

Blood samples were obtained 10 min after an iv injection of 30 IU heparin/kg body weight. LPL activity was measured by using radiolabeled triolein emulsion after HL inhibition by SDS as previously described (30).

ApoB kinetic studies

The kinetic study was performed in the fed state by iv administration of L-[1-13C] leucine (99 atom%, Eurisotop, St. Aubin, France), with a primed bolus of 0.7 mg/kg immediately followed by a 16 h constant infusion of 0.7 mg/kg·h as previously described (31). VLDL, IDL, and LDL were isolated from plasma and delipidated using diethylether-ethanol 3:1. Apo B-100 was isolated in each lipoprotein fraction by preparative SDS-PAGE. After hydrolysis, amino acids of apo B-100 were recovered from an AG-50W-x8 200–400 mesh cation exchange column (Bio-Rad Laboratories, Inc., Richmond, CA). Amino acids were converted to N-acetyl O-propyl esters and were analyzed with a Finnigan Mat {delta} C isotope ratio mass spectrometer (Finnegan Mat, Bremen, Germany). Data were analyzed with the Stimulation Analysis And Modeling (SAAM) II program (SAAM Institute, Inc., Seattle, WA) using a multicompartmental model as previously described (32). Direct fractional catabolic rates (FCRs) of VLDL apoB and FCR from VLDL to IDL or LDL were calculated as follows:


where k (i, j) is the fractional transfer coefficient from compartment j to i. Mj represents the mass expressed as concentration per liter of plasma of compartment j. Data were normalized to the plasma volume of each subject.

Statistical analysis

Statistical analysis was performed using the StatView 4.5 software. Data from patients and controls were compared using Student’s t test for unpaired values or ANOVA. The relationships between variables were analyzed after calculation of linear correlation coefficients.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
To evidence the involvement of LPL bound to VLDL in the lipolysis of VLDL-TG, we performed a series of experiments that is described in Fig. 2Go. The VLDL-TG hydrolysis observed during the incubation of VLDL isolated from a LPL-deficient patient was extremely low compared with that from a group of three control subjects. However, this low hydrolysis was normalized when LPL-deficient VLDL was supplemented with exogenous bovine LPL. Moreover, addition of antihuman HL antibody did not decrease TG hydrolysis in VLDL isolated by FPLC from control subjects (113.9 ± 30.7 vs. 111.2 ± 29.3 nmol NEFA/mL·h at the highest concentration of antibody, mean ± SD of four independent experiments). It was therefore concluded that VLDL-TG hydrolysis occurred without any detectable contribution of HL, but that it was exclusively a VLDL-associated LPL-dependent process.



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  		Figure 2. Evidence that LVTH level is related to the presence of VLDL-bound LPL. LVTH values were determined during incubations at 37 C of VLDL isolated either from control plasma (•) or from one LPL-deficient plasma before ({blacksquare}) and after ({square}) supplementation with exogenous LPL. Values for controls are shown as the mean ± SD of three individuals, whereas those for the LPL-deficient patient represent the mean ± SD of three independent determinations.

 
Plasma LVTH activity was determined in a group of 20 control subjects and in three types of patients known to have mild or severe hypertriglyceridemia. These included 13 type 2 diabetics, 5 patients homozygous for LPL gene mutations, and 5 patients with heterozygous LPL deficiency (Fig. 3Go). The plasma LVTH activity of type 2 diabetics was 37% lower than that of control subjects (P < 0.0001). Similarly, that of patients with heterozygous mutations exhibited a 42% decrease (P = 0.0004). In contrast, almost no LVTH was detectable in the samples obtained from homozygous LPL-deficient subjects. No overlap was observed between the latter group and those of either control subjects or patients with heterozygous LPL mutations. In none of the groups mentioned above was plasma LVTH activity affected by either sex or age.



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  		Figure 3. Plasma LVTH levels in 20 control subjects, in 13 type 2 diabetics as well as in 5 heterozygous and homozygous LPL-deficient patients. Data are shown as the means ± SD together with individual values. Statistical comparison between groups was assessed by one-way ANOVA.

 
Plasma TG concentrations were inversely related with LVTH according to an hyperbolic-type curve that was linearized after double reciprocal transformation of the data, thereby permitting the valid calculation of a linear correlation coefficient (r = -0.56, P = 0.0002) (Fig. 4Go). This negative correlation was found in control subjects (r = -0.51, P = 0.022), with a larger variability in subjects with type 2 diabetes (r = -0.39, P = 0.195). Similar results were obtained when plasma TG values were replaced by those of VLDL-TG (r = -0.63, P < 0.0001). As usually observed, we found a negative correlation between the plasma concentrations of TG and those of HDL-C (r = -0.67, P < 0.0001). However no significant correlation appeared between HDL-C and LVTH (Fig. 5Go). In control subjects and in diabetic patients LDL-C was positively correlated to LVTH (r = 0.46, P = 0.040 and r = 0.60, P = 0.028, respectively), whereas in the other groups, the number of data were to low to permit any accurate statistical analysis. Finally, plasma NEFA concentrations were positively correlated to LVTH (r = 0.63, P = 0.0021 and r = 0.80, P = 0.0005 for control subjects and diabetic patients, respectively). LVTH and PHLA were found positively correlated (r = 0.67, P = 0.047) in a subgroup of nine subjects that had simultaneous determination of both LPL assays (Fig. 6Go). In this subgroup, the inverse correlation of LVTH with plasma TG was stronger than that obtained with PHLA (r = -0.69, P = 0.057 vs. r = -0.36, P = 0.355). Similar results were obtained with VLDL-TG concentration (r = -0.73, P = 0.023 vs. r = -0.30, P = 0.455).



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  		Figure 4. Relationship between plasma TG concentrations and LVTH levels. The individual values obtained for control subjects (•), for type 2 diabetics ({circ}) and for heterozygous LPL-deficient patients (+) are shown in the inset. The data were linearized by double reciprocal transformation to permit the accurate calculation of the linear correlation coefficient (r = -0.56, P < 0.001). The values from homozygous LPL-deficient patients were largely out of scale and were therefore excluded from this figure only for the sake of presentation.

 


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  		Figure 5. Relationship between plasma HDL-C or LDL-C concentrations and LVTH levels in control subjects (•), in type 2 diabetics ({circ}) and in heterozygous LPL-deficient patients (+). LVTH did not correlate with HDL-C (A), whereas positive correlations were observed with LDL-C in control subjects (r = 0.46, P = 0.0403) and in type 2 diabetics (r = 0.60, P = 0.0278) (B). In heterozygous LPL-deficient patients, the number of data were too low to permit an accurate statistical analysis.

 


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  		Figure 6. Correlation between LVTH and (PHLA) postheparin LPL activity (r = 0.67, P = 0.0466).

 
To further characterize the physiological relevance of LVTH measurements, the latter were determined in a subgroup of nine subjects who were studied for their time course of VLDL catabolism after apo B labeling with L-[1-13C] leucine. LVTH was found highly correlated to the rate of conversion of VLDL into IDL (r = 0.82, P < 0.0042, Fig. 7AGo). The correlation between the rate of conversion was found stronger with LVTH than that obtained with PHLA in the five patients that had both measurements (r = 0.82, P = 0.0042 vs. r = 0.68, P = 0.24). LVTH as well as PHLA was not correlated with direct hepatic removal of VLDL (r = 0.20, P = 0.621 and r = -0.34, P = 0.619 respectively, Fig. 7BGo).



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  		Figure 7. Relationships between LVTH levels and the kinetics of VLDL catabolism in nine control subjects. Both the FCR of direct hepatic removal of VLDL and of conversion into IDL were determined by mass spectrometry, using L-[1-13C] leucine-labeled apolipoprotein B as the marker. LVTH was highly correlated to the rate of conversion of VLDL into IDL (r = 0.82, P = 0.0042, A), but not with the direct hepatic removal of VLDL (r = 0.20, P = 0.6213, B).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
This work describes a new assay designed to account for plasma TG hydrolysis through the measurements of VLDL-bound LPL-dependent VLDL lipolysis. Because this assay is based upon the TG hydrolysis of endogenous VLDL, it accounts for the actual LPL-dependent lipolytic capacity of circulating VLDL rather than for a simple LPL activity assay. This is in contrast with previous assays where pre- or postheparinic plasma samples were tested for LPL activity using a lipid emulsion containing radioactive triolein as the substrate (6, 7, 8, 9, 10, 11, 12, 13). Although the latter assay likely results in a good evaluation of the active LPL enzyme concentration, its relevance with respect to the analysis of physio-pathological situations remains questionable, for two important reasons. Firstly, the in vivo level of plasma LPL-dependent lipolysis not only depends upon the active enzyme concentration but also upon the concentration and composition of substrate lipoproteins, which are criteria that are not taken into account in the radioisotopic assay (33). Secondly, the postheparinic radioisotopic assay is supposed to account for the sum of circulating plus endothelium-bound LPL activity. Thus, it represents the total of the putatively active LPL occurring at a given time, which in no way means that the bulk of this LPL was really physiologically active at this time. From this viewpoint, it is important to note that LPL activity was reported to increase by 100 to 300 times after heparin injection but that no correlation was observed between either LPL activity (isotopic assay) or concentration (sensitive ELISA) determined before and after heparin injection (22, 23, 24). These considerations probably explain for a large part certain drawbacks of the postheparin assay that have been pointed out by others. For example, significant residual LPL activities were found in homozygous subjects with null mutation, and frequently postheparin LPL activity in heterozygous remained indistinguishable from that of controls (34, 35, 36). Further evidence of the lack of capacity of postheparin LPL activity measurements to detect mild alteration in lipolytic activity in pathological condition was the inconsistent decrease in postheparin LPL activity in hypertriglyceridemia (24, 37, 38). Similarly, Taskinen et al. were only able to evidence a change in the LPL/HL ratio in the insulin-resistant state (39).

The new assay that we describe in this report might bring several significant improvements with respect to ethical considerations, technical aspects, and physiopathological relevance. Clearly, our LVTH assay avoids the injection of heparin, whereas 10, 60, or 100 UI/kg have been commonly used in PHLA measurements (7, 10, 22, 24, 40). This simplifies blood drawing and particularly allows serial measurements of plasma samples. In addition, LVTH assay does not imply the use of radioactive materials. However, albeit simple, the accurate determination of LVTH requires basic precautions. Blood must be drawn on ice and plasma immediately separated at 4 C to avoid spontaneous lipolysis, and because it has been suggested that LPL was transferred from TG-rich lipoproteins to LDL after a few minutes at room temperature (20, 26). Storage at -80 C is mandatory because a rapid decrease of lipolytic activity has been described in samples stored at -20 C. As to the preparation of VLDL, it is absolutely necessary to avoid the use of ultracentrifugation that leads to a major loss of VLDL-bound LPL (16, 28). In our hands, the separation of VLDL performed by FPLC was found rapid and convenient. The lack of LVTH activity observed in patients with homozygous LPL deficiencies clearly shows that our assay specifically measures LPL activity without detectable contribution of HL activity. This is of particular interest inasmuch as the neutralization of HL activity has always been a major concern in PHLA assay. Variable residual HL activities have been described in various reports, irrespective of the method used to discriminate between LPL and HL activities (salt concentration, or addition of protamine or SDS or anti-HL antibodies) (7, 8, 9, 10, 11, 12, 13, 30, 40). Because of these difficulties, it was frequently observed that the summation of LPL plus HL activities did not correspond to the total lipolytic activity directly measured in the samples.

The lack of HL contribution in our LVTH assay increases its interest from a physiopathological viewpoint. Indeed, it permits the clear discrimination of homozygous LPL-deficient patients because no overlap of data occurred with those of heterozygous patients, although the latter had LVTH activities approximately half of those of control subjects. To be considered as a good index reflecting the in vivo LPL-mediated lipolysis, LVTH measurements must be consistent with already well established processes. For example, plasma TG concentrations are believed to be largely driven by lipolysis, leading to the classical inverse relationship between triglyceridaemia and the levels of lipolysis. Consistent with this concept, we found that LVTH activities of type 2 diabetics were comparable with those of heterozygous LPL-deficient patients, whereas their plasma TG concentrations were also similar. In addition, we observed that the values of total or VLDL triglyceridemia and those of LVTH distributed around the same correlation line, for all subjects, whether patients or controls, with a larger variability in the former group, probably due to the contribution of hepatic overproduction of VLDL. These data clearly meet the criteria that have to be expected from a physiologically relevant lipolysis index. Moreover, although LVTH and PHLA were positively correlated, the correlations between LVTH and TG parameters (plasma concentration, VLDL conversion, VLDL-TG concentration) were systematically stronger than these obtained with PHLA in the subgroup of subjects that had a simultaneous determination of both LPL measurements. As usually observed, plasma triglyceridemia was positively correlated to HDL-C concentrations (41). However, no correlation was found between the latter and LVTH either in control or in type 2 diabetics, in contrast with previous works reporting that postheparin LPL activity was positively correlated with HDL-C (22, 24, 42). Although we have no clear explanation for that difference, our data are in agreement with other reports where preheparin LPL activity was measured (22, 23). In addition, transgenic mice overexpressing LPL failed to exhibit any increase in HDL-C (43).

To further analyze the overall validity of LVTH measurements, we specifically performed a series of experiments in which the kinetics of VLDL catabolism were measured in nine control subjects. The results clearly indicated that LVTH highly correlated with the conversion of VLDL to IDL, but not with the direct hepatic uptake of VLDL. These data demonstrate that LVTH assay quantitatively accounts for the lipolysis occurring in physiological conditions. Consistent with this finding, LVTH was also correlated to the plasma NEFA concentrations and to those of LDL-C. The positive correlation between LVTH and plasma NEFA concentration in type 2 diabetic subjects suggests indirectly that plasma NEFA concentration might be influenced both by the level of insulin resistance (increased lipolysis from intraabdominal fat) and LPL activity.

Finally, LVTH appears as a physiologically relevant index of the LPL-mediated lipolysis of plasma TG, which could advantageously replace the measurement of postheparin LPL activity at least in patients with type 2 diabetes and genetic LPL deficiency and may be in others numerous situations.


   Acknowledgments
 
We thank Jacobs Chantal and the nursing staff for their expert technical assistance.


   Footnotes
 
1 This work was supported by Claude Bernard-Lyon I University and by the Hospices Civils de Lyon. Back

Received March 14, 2000.

Revised July 31, 2000.

Revised October 24, 2000.

Accepted October 25, 2000.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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