Transport of lipoprotein lipase across endothelial cells (triglyceride/fatty acids/adipose tissue/proteoglycans/glycosaminoglycans)

Transcription

1 Proc. Natl. Acad. Sci. USA Vol. 88, pp , March 1991 Medical Sciences Transport of lipoprotein lipase across endothelial cells (triglyceride/fatty acids/adipose tissue/proteoglycans/glycosaminoglycans) UDAY SAXENA, MICHAEL G. KLEIN, AND IRA J. GOLDBERG* Department of Medicine and Specialized Center of Research in Arteriosclerosis, Columbia University, College of Physicians and Surgeons, New York, NY 132 Communicated by Richard J. Havel, December 26, 199 (received for review March 8, 199) ABSTRACT Lipoprotein lipase (LPL), synthesized in muscle and fat, hydrolyzes plasma triglycerides primarily while bound to luminal endothelial cell surfaces. To obtain information about the movement of LPL from the basal to the luminal endothelial cell surface, we studied the transport of purified bovine milk LPL across bovine aortic endothelial cell monolayers. 125I-labeled LPL ('25I-LPL) added to the basal surface of the monolayers was detected on the apical side of the cells in two compartments: (i) in the medium of the upper chamber, and (ii) bound to the apical cell surface. The amount of 125I-LPL on the cell surface, but not in the medium, reached saturation with time and LPL dose. Catalytically active LPL was transported to the apical surface but very little LPL activity appeared in the medium. Heparinase treatment of the basal cell surface and addition of dextran sulfate (.15 IAM) to the lower chamber decreased the amount of 125I-LPL appearing on the apical surface. Similarly, the presence of increasing molar ratios of oleic acid/bovine serum albumin at the basal surface decreased the transport of active LPL across the monolayer. Thus, a saturable transport system, which requires heparan sulfate proteoglycans and is inhibited by high concentrations of free fatty acids on the basal side of the cells, appears to exist for passage of enzymatically active LPL across endothelial cells. We postulate that regulation of LPL transport to the endothelial luminal surface modulates the physiologically active pool of LPL in vivo. Lipoprotein lipase (LPL) bound to endothelial cell-surface heparan sulfate proteoglycans hydrolyzes triglycerides in plasma lipoproteins, especially chylomicrons and very low density lipoproteins (1, 2). LPL is not synthesized by the vascular endothelial cells but is produced by underlying adipocytes and myocytes (3-5). The physiologic actions of LPL are mediated primarily by the pool of LPL that is located at the luminal surface of the endothelium. Therefore, after its synthesis and secretion, LPL must be transported from the basal to the luminal side of the endothelial cell. How LPL is routed from its initial site of synthesis to the endothelial surface is not understood. Molecules can be transported across endothelial cells by nonspecific and specific mechanisms (6, 7). The nonspecific processes include uptake by fluid phase and adsorptive pathways, which lead to transcytosis. Plasmalemmal vesicles, formed from the plasma membrane, shuttle between endothelial cell luminal and basal surfaces, discharging their contents to the plasma or interstitial fluid compartments. Albumin transport across endothelial cells is an example of such a process (8, 9). Smaller molecules like free fatty acids (FFAs) are thought to transfer across endothelial cells via lateral diffusion in cell membranes (1). Some movement can also occur through gaps in the intercellular tight junctions, which form a barrier between the plasma and interstitial The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C solely to indicate this fact. compartments (6). Specific transport may require initial interaction of a molecule with high-affinity receptors present on the cell surface (11, 12). Insulin, for example, is bound to its receptor, internalized by endothelial cells, and then transported from the apical to the basal endothelial cell surface in endocytic vesicles. LPL must be transported across endothelial cells in the opposite direction, from the basal to the apical surface of the cells. LPL transport was studied by using endothelial cell monolayers grown on a permeable polycarbonate filter system, which separates culture dishes into two compartments. Native LPL and 251I-labeled LPL (125I-LPL) were added to the lower chamber and the appearance of radioactivity and LPL hydrolytic activity in the upper-chamber medium and on the apical cell surface were measured. In addition, the role of proteoglycans in the transport of LPL was explored. MATERIALS AND METHODS Bovine Milk LPL: Purification and Radioiodination. LPL was purified from fresh unpasteurized milk, radioiodinated, and stored at -7TC as described (13). At the time of storage, LPL preparations had a specific activity of approximately 2-3 mmol offfa per hour per mg of protein. Labeled LPL had a specific radioactivity activity of -5 dpm per ng of protein and >9%o of the counts were precipitated by 1%o trichloroacetic acid. Heat-inactivated LPL, prepared by heating LPL for 1 hr at 52C, was used in some experiments. Endothelial Cell Monolayers. Primary cultures of bovine aortic endothelial cells were established as described for porcine aortic endothelial cells (13). Cells were plated onto 25-mm polycarbonate filters (pore diameter, 3. gm; Nuclepore), according to the method of Shasby et al. (14). Each gelatin- and fibronectin-coated filter was seeded with 8-1 x 15 cells in 1.5 ml of Dulbecco's modified Eagle's medium (DMEM) containing 1% bovine calf serum, antibiotics (1 units of penicillin per ml and 1 gg of streptomycin per ml; Hazelton Research Products, Lenexa, KS), and glutamine (1%). The media in the upper chambers (1.5 ml) and lower chambers (2.6 ml), separated by the filter, were replaced every other day. Experiments were conducted 5-6 days after seeding the endothelial cells. For 12 hr prior to each experiment, the cells were maintained in DMEM without calf serum to reduce the number of surface proteoglycan binding sites occupied by serum proteins. The barrier function of the endothelial cell monolayer was examined by several methods. After a 1-hr incubation at 37C, there was a 3.5-fold greater amount of 125I-LPL in the medium of the upper chamber when filters that did not contain any cells were used compared to the filters containing the monolayers; a 1-fold greater amount of 1251-LPL was Abbreviations: LPL, lipoprotein lipase; BSA, bovine serum albumin; FFA, free fatty acid. *To whom reprint requests should be addressed at: Department of Medicine, Columbia University, College of Physicians and Surgeons, 63 West 168 Street, New-York, NY

2 Medical Sciences: Saxena et al. found in the upper chamber if the filters were not coated with collagen and fibronectin. In pilot experiments, the seeding density of the cells was increased 5-fold to =4 million cells per well. This did not change the transport of '25I-LPL into the medium, compared to the filters on which the usual number of cells was seeded, demonstrating that increasing the number of cells seeded did not increase the barrier function of the monolayer. Transport rates of [14C]albumin from the basal to the apical side of the monolayers were <1.5% per hr, a rate similar to that reported by Stoll and Spector (15). At the conclusion of each LPL transport experiment, the monolayers were stained with 2% toluidene blue to verify the uniformity of the monolayer. Transport Studies. On the day of the experiment, culture media from both chambers were aspirated and the cells were washed three times with DMEM containing 3% bovine serum albumin (DMEM-BSA). All further steps were carried out with DMEM-BSA. After washing, LPL (radioiodinated or unlabeled) was added to the lower chamber, and then the chambers were incubated at 37C for up to 1 hr. At the end of the incubation period, the chambers were transferred to the cold room (4C) and washed three times with cold DMEM-BSA. LPL associated with the apical cell surface was then released by the addition of DMEM-BSA containing heparin (1 units/ml; Organon) at 4C for 1 min to the upper and lower chambers. 1"I-LPL radioactivity present in the three compartments-(i) medium from the upper chamber, (ii) apical cell surface, and (iii) basal cell surface-was measured. In other experiments, LPL enzymatic activity in these three compartments was assayed by using a high specific activity substrate emulsion (16). To confirm that active LPL was present on the apical surface of endothelial cells and to avoid inhibition of the assay by heparin, LPL was also directly assayed while bound to the cells (13). Cellular LPL Measurements LPL associated with the cell (representing internalized LPL) was measured after removing surface-associated LPL with heparin, lifting the cell monolayers with.25% trypsin (37C for 5 min), centrifuging at 25 x g for 1 min, and washing the cell pellets with DMEM-BSA. To analyze the cell-associated 125I-LPL, the cell pellets were lysed in buffer containing 1 mm Tris-HCl (ph 7.5), 1% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate, and 1% octyl glucoside. The cell extract was sonicated for 2 sec with a Branson sonifier. The extracts were then centrifuged at 25 x g for 1 min and the supernatants were precipitated with 1%o trichloroacetic acid. The precipitates were then analyzed by SDS/PAGE using a 7.5% gel. RESULTS Concentration and Time Dependence of LPL Transport Across Monolayers. To determine whether LPL was transported across endothelial cells from the basal to the apical surface, increasing concentrations of radioiodinated LPL (.5-12 ug) were added to the lower chamber and the cells were incubated for 1 hr at 37 C. With increasing amounts of LPL (Fig. LA), the amount of 1251-LPL bound to the apical surface increased with the addition of up to 6,ug of 125I-LPL, and then remained unchanged as more LPL was added (up to 12,ug). In contrast, there was a linear increase in the amount of 115I-LPL in the medium from the upper chamber. Most (9% + 2%) of the 125I-LPL found in the medium and that released by heparin from the apical surface was precipitable by 1% trichloroacetic acid. These results demonstrate that 125I-LPL is transported across endothelial cells without much degradation. The time course of LPL transport was studied by using 2 pug of 1251-LPL added to the lower chamber. The amount of 125I-LPL found on the apical surface (released by heparin) FIG. 1. Proc. Nadl. Acad. Sci. USA 88 (1991) 2255 A 8- B C D ' 6 -j fi' 4 i 2 C -I a. -C v- n.l LPL added (IJg) :j2 L % -j 1 II.i II IIII I'i Time (minutes) Time (minutes) 2 4 rime (minutes) n. -i O 4 *1 a C -J.8I -a 6 U 4 CD 2 Q 4 Time and dose dependence of LPL transport across endothelial cell monolayers. (A) Effect of addition of increasing amounts of LPL. 125I-LPL in DMEM-BSA was added to the lower chamber (basal surface) of endothelial cell monolayers and the cells were incubated at 37TC for 1 hr. Amounts of 15I-LPL in the upper-chamber medium () and released from the apical surface by heparin (A) are shown. Values presented are the means of three separate observations ± SEM. (B) Time course of transport of 1"I-LPL across endothelial cell monolayers. 125I-LPL (2 pag) in DMEM-BSA was added to the lower chamber of endothelial cell monolayers. The cells were incubated at 37"C and 125I-LPL present in the upper-chamber medium (e) and on the apical cell surface (A) was measured at the indicated times. Each data point represents the average oftriplicate dishes. (C) Time course oftransport of LPL activity. Unlabeled LPL (1jLg) in DMEM-BSA was added to the lower chamber and the experiment was then performed as described for 125I-LPL (see B), except that triglyceride hydrolytic activity of LPL in the upper-chamber medium (e) and on the apical surface (A) was assayed by using a high specific activity substrate emulsion (16). The results are the averages of duplicate determinations and are presented as FFA generated per ml ofmedium per hr. (D) Direct measurement of LPL activity on the apical surface of endothelial cells. In a separate experiment, bovine LPL(lOpug) in DMEM-BSA was added to the lower chamber and at the times indicated the media in the lower and upper chambers were removed, the cells were washed, and 1.5 ml ofdmem- BSA containing 1 Al of substrate emulsion was added to the upper chamber and LPL activity was determined as described. Data shown are averages of assays from duplicate dishes for each time point. increased for 2 min, remained relatively constant until 6 min, and then decreased (Fig. 1B). A linear increase of

3 2256 Medical Sciences: Saxena et al. radioactivity occurred in the upper-chamber medium for up to 12 min. A similar time-dependent transport of LPL enzymatic activity released from the apical surface by heparin was found; LPL activity increased for 2 min and then decreased slightly over the next 4 min (Fig. 1C). In contrast, there was very little activity present in the medium at all times between 5 and 6 min. These data measuring enzyme activity suggest that only enzyme bound to the apical surface is enzymatically active. Radioiodinated LPL present in the medium in the prior experiments may represent transport of enzymatically inactive LPL molecules. Alternatively, LPL in the medium may have lost its catalytic activity during the experiments. Measurements of LPL activity released from the apical cell surface by heparin might have underestimated the actual amount of transported LPL due to instability of the enzyme or interference with triglyceride hydrolysis in the assay by heparin. LPL activity assayed by addition of emulsion directly to the upper-chamber medium is shown in Fig. 1D. A time course of LPL transport similar to that in Fig. 1C was obtained; LPL activity on the apical surface peaked at 2 min. A comparison of the activity of LPL transported to the apical surface with that of the original preparation was performed. Using the data obtained in Fig. 1B and the LPL activity in Fig. 1D, at 2 min the specific activity of transported LPL was 2.1 mmol of FFA mg-1'hr-1. This specific activity was greater than the specific activity of 1.6 mmol of FFA-mg-1hr-1 of the same preparation of purified LPL incubated in buffer for 2 min at 37C. In contrast, at 2 min the specific activity of LPL in the upper-chamber medium was markedly reduced (.5 mmol of FFA mg-1-hr-1). Thus, LPL on the apical cell surface appeared to be protected against inactivation when compared to LPL in the medium or control incubated LPL. To further examine the hypothesis that enzymatically inactive LPL may be preferentially transported directly to the medium, LPL was deliberately heat inactivated. Although this enzyme was enzymatically inactive, SDS/PAGE analysis showed that the heat-inactivated LPL was the same size as unheated LPL. Use of the heated enzyme produced a 2.8-fold increase in the amount of LPL present in the medium compared to unheated LPL. By contrast, compared to control LPL, heat inactivation of LPL reduced the amount bound to the apical surface by 4%. These data suggest that inactive LPL may be preferentially transported to the medium Requirement of Basal Heparan Sulfate Proteoglycans for LPL Transport. To test whether specific transport of LPL across the cells requires initial binding of LPL to heparan sulfate proteoglycans on the basal cell surface, the basal side of the monolayers was treated with heparinase. In another experiment, LPL binding to proteoglycans was reduced by the addition of dextran sulfate to the lower chamber. The heparinase treatment decreased the amount of 125I-LPL bound to the basal cell surface by >8o relative to control cells (not treated by heparinase) (Fig. 2). The amount of 125I-LPL transported to the apical cell surface decreased by 62%. Heparinase pretreatment did not alter the binding of LPL to the apical cell surface, suggesting that heparan sulfate binding sites on that surface were not affected by the pretreatment. With dextran sulfate addition to the lower chamber, there was an 88% decrease in the amount of 125I-LPL bound to the basal cell surface compared to control incubation (no dextran sulfate) and a corresponding decrease in the amount of 125I-LPL found on the a ical surface (95% decrease). In contrast, the amount of 5I-LPL transported to the medium in the upper chamber did not change with dextran sulfate addition and decreased only 11% with heparinase treatment. These results suggest that the binding of LPL to heparan sulfate proteoglycans on the basal cell surface is _ -. on 2-.1 If Proc. Natl. Acad. Sci. USA 88 (1991) G-1 -Henari'nase Dextrar, -:"IfratL FIG. 2. Effect of heparinase pretreatment and dextran sulfate on the transport of LPL. The basal surface of endothelial cell monolayers was treated with.1 unit of heparinase (ICN) for 2 hr at 37C prior to adding 125I-LPL (8,ug). In a separate experiment, 125I-LPL (8,ug) was added to the bottom chamber together with dextran sulfate (.15 1uM). The cells were then incubated at 3TC for 1 hr. 125I-LPL in the upper-chamber medium (n) and 125I-LPL released from the apical (a) and basal (a) cell surfaces after addition of heparin were determined. Results are expressed as percent of a control incubation. required for the transport of LPL to the apical cell surface, whereas the transport into the upper-chamber medium is minimally affected by the removal of heparan sulfate binding sites. Internalization of LPL During Transport. To examine whether LPL bound to the basal cells surface was internalized under conditions in which LPL was transported to the apical cell surface, 125I-LPL was bound to the basal surface for 2 hr at 4C and the monolayers were then incubated for 2 min at either 4C or 37C and cellular 125I-LPL was measured LPL on the apical cell surface was 41% lower at 4 C than at 37rC and there was a >8%o decrease in the amount of cell-associated radioactivity at 4C. When 1251-LPL and dextran sulfate were added together to the lower chamber, a 64% decrease in the amount of LPL radioactivity associated with the cells relative to control cells was observed. However, when dextran sulfate was added to the upper chamber of monolayers in which 1251-LPL was present in the lower chamber, there was no decrease (and in some experiments there was a small increase) in the cell-associated radioactivity. Analysis of the cell-associated radioactivity (Fig. 3) showed that the mobility of 1251-LPL associated with the cells was identical to that of control 1251-LPL. Because internalization occurs under the same conditions that demonstrate transport of 125I-LPL to the apical surface, it suggests that internalization is required for proteoglycan-mediated LPL transport. A second question is whether LPL that moves to the upper-chamber medium requires intracellular pathways or transfers via paracellular mechanisms. If LPL in the upper chamber is decreased with treatments that affect cellular FIG. 3. SDS/polyacrylamide gel analysis of control and internalized 1251-LPL. Internalized LPL from endothelial cells (lane 2) and 125I-LPL incubated under similar conditions in the absence of cells (lane 1) were applied to a 7.5% polyacrylamide gel. The gel was run at 2 ma for 6 min, fixed, and exposed to Kodak X-Omat film for 2 weeks. Approximately 2 cpm was applied to each lane.

4 functions, this would support a role for an intracellular pathway of LPL transport to the medium. Incubation at 4C reduced the amount of LPL found in the upper chamber after 1 hr by -5O%. When cells were treated with colchicine or sodium cyanide, however, an increase rather than a decrease in LPL was found in the upper chamber. This occurred despite a decrease in the amount of internalized 1251-LPL (53% decrease with colchicine and 45% decrease with sodium cyanide), also observed at 4C, and suggested that these metabolic inhibitors may have disrupted the integrity of the endothelial monolayers (17). Thus, whether LPL movement to the upper-chamber medium is via cellular or paracellular mechanisms cannot be concluded from these experiments. Inhibition of LPL Transport by Increased FFA Concentrations. We have previously shown that FFAs can dissociate LPL bound to endothelial cell surfaces (13, 18) and hypothesized that addition of FFA should decrease the transport of LPL to the apical endothelial surface. To examine the effect of FFAs on LPL transport, increasing molar ratios of oleic acid/bsa were added to the lower chamber together with unlabeled LPL. The addition of oleic acid markedly decreased the amount of LPL enzymatic activity that was transported to the apical cell surface (Fig. 4A). No LPL activity was measurable in the medium (data not shown), consistent with the results shown in Fig. 1C. To specifically assess the effects of fatty acids on the transport of LPL protein that was bound to the basal surface of the cells, 1251-LPL was allowed to bind to the basal surface for 2 hr at 4C. The medium containing the unbound"-5i-lpl was removed from the lower chamber and replaced with medium containing increased molar ratios of oleic acid/bsa. The cells were incubated for 1 hr at 37C. With increasing C.) a. ) c -J if) NY FIG. 4. Medical Sciences: Saxena et al o1 8' 6 4f 2 'A.5:11:1 2:1 4:1 2:1 Oleic acid/bsa, mol Effect of oleic acid/bsa molar ratios on the transport of LPL. (A) LPL activity. LPL (8 ug) was added together with increasing molar ratios of oleic acid/bsa to the lower chamber and the cells were incubated at 37C for 1 hr. After the incubation, LPL activity on the apical surface was determined by using a radioactive triglyceride emulsion as described. (B) '"I-LPL. 125I-LPL (8,ug) was allowed to bind to the basal surface of the monolayer for 2 hr at 4TC. The medium in both chambers was changed, increasing molar ratios of oleic acid/albumin were added to the lower chamber, and the endothelial cell monolayer was incubated for 1 hr at 37C. Radioactivity in the upper-chamber medium () and on the apical cell surface (*) was determined. Proc. Natl. Acad. Sci. USA 88 (1991) 2257 concentrations of fatty acids in the lower-chamber medium, less 125I-LPL was found on the apical surface (Fig. 4B). Therefore, higher concentrations ofoleic acid decreased total LPL protein transport, as well as transport of catalytically active LPL to the apical surface. In contrast to cell-surface LPL, the amount of '25I-LPL transported to the medium increased with addition of FFA. This increase of 1"I-LPL in the medium may have been due to increased nonspecific transport of LPL by the cells or to an increase in transport of LPL that was inactivated by high concentrations of FFA. DISCUSSION Our results demonstrate that LPL protein and LPL catalytic activity are transported across endothelial cell monolayers. The transported LPL was found in two compartments: (i) LPL bound to the apical cell surface, which was specifically assessed by its release with heparin, and (ii) LPL present in medium in the upper chamber. Results presented in Fig. 1 C and D showed that the LPL bound to the apical cell surface effectively hydrolyzed a substrate triglyceride emulsion. Although 125I-LPL was found in the medium, very little active LPL was found in this compartment. Data from other experiments showed that the LPL increases in these compartments were not parallel (Fig. 1 A and B). The medium LPL increased linearly with addition of increasing concentrations of LPL to the lower chamber, whereas the increase in the amount of LPL bound to the apical cell surface appeared to reach saturation. LPL transport to the apical surface and to the upperchamber medium appears to involve two distinct pathways. Several experiments suggest that apical surface LPL and LPL in the medium are not precursors of each other. By blocking the binding of LPL to heparan sulfate proteoglycans, we were able to markedly decrease only the LPL on the apical surface. We therefore believe that the apical surface LPL is not the precursor of most of the LPL in the medium. An alternative possibility, that LPL is first transported to the upper-chamber medium and then associates with heparan sulfate proteoglycans on the apical surface, is also unlikely because the LPL increase on the apical cell surface occurs prior to, not after, LPL appearing in the medium. In addition, if LPL was first bound to the basal surface (Fig. 4B) or was present in the lower chamber (Fig. la), similar amounts of LPL appeared on the apical surface despite marked differences in the amount of LPL in the upper-chamber medium. Thus, our results are most consistent with two transport pathways: one that delivers LPL to the apical surface of endothelial cells, and one through which inactive LPL travels to the upper-chamber medium. Is the pathway of LPL transport to the apical surface demonstrated in our studies operative in vivo? One issue to be considered when responding to this question is whether the pathway allows LPL to retain its enzymatic activity. As shown in Fig. 1D, LPL transported to the apical surface of endothelial cells does not lose its enzymatic activity and, compared to LPL incubated in medium alone, actually has a greater specific activity. Moreover, the calculated specific activity of apical surface LPL is likely to be an underestimate of the true specific activity of the enzyme. This is because LPL bound to solid supports is markedly less active than LPL in solution (19). A second issue is whether the efficiency of LPL transport is comparable to other receptor-mediated transcytosis pathways. King and Johnson (11) reported that =8% of insulin added to the apical surface of endothelial cells was transported to the lower chamber within 1 hr. In our studies, LPL was added to the lower-chamber medium, separated from the basal surface of the cells by the polycarbonate filter. Thus, the concentration of LPL directly in contact with the cells

5 2258 Medical Sciences: Saxena et al. Proc. Natl. Acad. Sci. USA 88 (1991) was initially somewhat less than that in the chamber. If the amount of LPL on the apical surface is compared to that added to the lower chamber (Fig. 1), its transport appears to be very low (<2%). However, when LPL was first bound to the basal cell surface at 4'C (Fig. 4B) followed by a 1-hr incubation at 37C, the amount of LPL present on the apical surface was 15-22% of that originally on the basal surface. Therefore, once bound to the basal surface, LPL is efficiently transported across endothelial cell monolayers. Our studies suggest a crucial role for heparan sulfate proteoglycans in the movement of enzymatically active LPL across endothelial cells. A requirement for the participation of basal cell-surface heparan sulfate proteoglycans in the transport of LPL to the apical cell surface was tested by digestion of glycosaminoglycans with heparinase and by addition of dextran sulfate, which prevents association of the enzyme with heparan sulfate. Both interventions decreased the amount of LPL transported to the apical surface, suggesting that LPL is transported to the apical surface by receptor-mediated processes in which heparan sulfate proteoglycans serve as receptors for LPL. Such a role for proteoglycans in cellular transport has not previously been described. Cell-surface proteoglycans are integral membrane-bound proteins and are ideally suited to serve this function (2). Furthermore, we have reported (21) that the interaction of LPL with its endothelial cell-surface binding site is resistant to dissociation with acid ph, perhaps explaining why LPL is not targeted to lysosomes. Using immunohistochemistry, Blanchette-Mackie et al. (22) have demonstrated LPL protein within endothelial cells that may be in the process of transcytosis. The transport of inactive LPL into the medium may be via a nonspecific fluid phase or adsorptive mechanisms, processes that are operative in vascular endothelial cells (7). Such mechanisms are sensitive to molecular shape, size, and charge (6, 23). In this regard, nonspecific transport pathways may preferentially transport inactive, monomeric LPL molecules as opposed to dimeric active LPL (24, 25). Therefore, the LPL transport pathway leading to the medium may not be involved in movement of catalytically active LPL, but it might allow for disposal of inactive enzyme from the intercellular space. The recent description of a mutant, inactive, non-heparin-binding LPL molecule found in the plasma of a hyperchylomicronemic patient (26) suggests that a nonproteoglycan-mediated pathway for transport of LPL functions in vivo. Addition of oleic acid to the basal surface of endothelial cells decreased the transport of active LPL across endothelial cells. This reduction may be due to diminished binding of LPL to the basal surface, since high concentrations of oleic acid decrease LPL binding to endothelial cells (13, 18). This decrease in LPL transport to the luminal cell surface may play a role in regulation of LPL activity specifically in adipose tissue. Adipose tissue LPL activity is relatively high in fed animals and is reduced during fasting (2, 3). Although feeding increases LPL activity, Semb and Olivecrona (27) demonstrated that LPL protein synthetic rate is the same in adipose tissue from fed and fasting animals. Ong and Kern (28) reported that human LPL mrna and protein levels are not different during fasting and feeding. Thus, alterations in LPL activity may be regulated by mechanisms other than those affecting LPL mrna transcription and LPL protein synthesis. Paradoxically, Doolittle et al. (29) showed that LPL mrna and protein in rat adipose tissue increased during fasting, while LPL activity decreased. These investigators postulated that more LPL protein may be enzymatically inactive in adipose from fasting compared to fed animals. Adipocyte hormone-sensitive lipase is activated during starvation, causing the release of FFA into the interstitial space. If our in vitro results are relevant to the in vivo situation, the generation of FFA during starvation could decrease LPL transport to the endothelial luminal surface. This would lead to increased LPL protein in interstitial fluid, where it can be rapidly inactivated, and to decreased LPL activity in adipose tissue. Thus, a change in LPL activity without a change in LPL mass could occur. Such a mechanism for regulation of LPL via changes in LPL transport may be a posttranslational process that, in part, modulates LPL activity during feeding and fasting. 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