Human Hepatic and Lipoprotein Lipase: The Loop Covering the Catalytic Site Mediates Lipase Substrate Specificity
1995; Elsevier BV; Volume: 270; Issue: 43 Linguagem: Inglês
10.1074/jbc.270.43.25396
ISSN1083-351X
AutoresKlaus A. Dugi, Helén L. Dichek, Silvia Santamarina-Fojo,
Tópico(s)Diabetes Management and Research
ResumoHepatic lipase (HL) and lipoprotein lipase (LPL) are key enzymes that mediate the hydrolysis of triglycerides (TG) and phospholipids (PL) present in circulating plasma lipoproteins. Relative to triacylglycerol hydrolysis, HL displays higher phospholipase activity than LPL. The structural basis for this difference in substrate specificity has not been definitively established. We recently demonstrated that the 22-amino acid loops ("lids") covering the catalytic sites of LPL and HL are critical for the interaction with lipid substrate (Dugi, K. A., Dichek, H. L., Talley, G. D., Brewer, H. B., Jr., and Santamarina-Fojo, S.(1992) J. Biol. Chem. 267, 25086-25091). To determine whether the lipase lid plays a role in conferring the different substrate specificities of HL and LPL, we have generated four chimeric lipases. Characterization of these chimeric enzymes using TG (triolein and tributyrin) or PL (dioleoylphosphatidylcholine (DOPC) vesicles, DOPC proteoliposomes, and DOPC-mixed liposomes) substrates demonstrated marked differences between their relative PL/TG hydrolyzing activities. Chimeric LPL containing the lid of HL had reduced triolein hydrolyzing activity (49% of the wild type), but increased phospholipase activity in DOPC vesicle, DOPC proteoliposome, and DOPC-mixed liposome assay systems (443, 628, and 327% of wild-type LPL, respectively). In contrast, chimeric HL containing the LPL lid was more active against triolein (123% of the wild type) and less active against DOPC (23, 0, and 30%, respectively) than normal HL. Similar results were obtained when the lipase lids were exchanged in chimeric enzymes containing the NH2-terminal end of LPL and the COOH-terminal domain of HL. Exchange of the LPL and HL lids resulted in a reversal of the phospholipase/neutral lipase ratio, establishing the important role of this region in mediating substrate specificity.In summary, the lid covering the catalytic domains in LPL and HL plays a crucial role in determining lipase substrate specificity. The lid of LPL confers preferential triglyceride hydrolysis, whereas the lid of HL augments phospholipase activity. This study provides new insight into the structural basis for the observed in vivo differences in LPL and HL function. Hepatic lipase (HL) and lipoprotein lipase (LPL) are key enzymes that mediate the hydrolysis of triglycerides (TG) and phospholipids (PL) present in circulating plasma lipoproteins. Relative to triacylglycerol hydrolysis, HL displays higher phospholipase activity than LPL. The structural basis for this difference in substrate specificity has not been definitively established. We recently demonstrated that the 22-amino acid loops ("lids") covering the catalytic sites of LPL and HL are critical for the interaction with lipid substrate (Dugi, K. A., Dichek, H. L., Talley, G. D., Brewer, H. B., Jr., and Santamarina-Fojo, S.(1992) J. Biol. Chem. 267, 25086-25091). To determine whether the lipase lid plays a role in conferring the different substrate specificities of HL and LPL, we have generated four chimeric lipases. Characterization of these chimeric enzymes using TG (triolein and tributyrin) or PL (dioleoylphosphatidylcholine (DOPC) vesicles, DOPC proteoliposomes, and DOPC-mixed liposomes) substrates demonstrated marked differences between their relative PL/TG hydrolyzing activities. Chimeric LPL containing the lid of HL had reduced triolein hydrolyzing activity (49% of the wild type), but increased phospholipase activity in DOPC vesicle, DOPC proteoliposome, and DOPC-mixed liposome assay systems (443, 628, and 327% of wild-type LPL, respectively). In contrast, chimeric HL containing the LPL lid was more active against triolein (123% of the wild type) and less active against DOPC (23, 0, and 30%, respectively) than normal HL. Similar results were obtained when the lipase lids were exchanged in chimeric enzymes containing the NH2-terminal end of LPL and the COOH-terminal domain of HL. Exchange of the LPL and HL lids resulted in a reversal of the phospholipase/neutral lipase ratio, establishing the important role of this region in mediating substrate specificity. In summary, the lid covering the catalytic domains in LPL and HL plays a crucial role in determining lipase substrate specificity. The lid of LPL confers preferential triglyceride hydrolysis, whereas the lid of HL augments phospholipase activity. This study provides new insight into the structural basis for the observed in vivo differences in LPL and HL function. INTRODUCTIONHepatic lipase (HL) 1The abbreviations used are: HLhepatic lipaseLPLlipoprotein lipaseHDLhigh density lipoprotein(s)DOPCdiolehoylphosphatidylcholine. and lipoprotein lipase (LPL) are critical enzymes that play a central role in lipoprotein metabolism. Their main function is to hydrolyze triglycerides and phospholipids present in circulating plasma lipoproteins, including chylomicrons, very low and intermediate density lipoproteins, and HDL(1Jackson R.L. Boyer P.D. The Enzymes. XVI. Academic Press, New York1983: 141-186Google Scholar). In the process, they generate free fatty acids, which can be utilized for storage or generation of energy. By influencing HDL metabolism(2Goldberg I.J. Blaner W.S. Vanni T.M. Moukides M. Ramakrishnan R. J. Clin. Invest. 1990; 86: 463-473Crossref PubMed Scopus (98) Google Scholar, 3Kekki M. Atherosclerosis. 1980; 37: 143-150Abstract Full Text PDF PubMed Scopus (52) Google Scholar, 4Patsch J.R. Prasad S. Gotto Jr., A.M. Patsch W. J. Clin. Invest. 1987; 80: 341-347Crossref PubMed Scopus (329) Google Scholar), these two lipases may also play a role in reverse cholesterol transport.Together with pancreatic lipase, HL and LPL are members of the human lipase family and share a high degree of primary sequence homology(5Hide W.A. Chan L. Li W.H. J. Lipid Res. 1992; 33: 167-178Abstract Full Text PDF PubMed Google Scholar). Their catalytic domain consists of a Ser-His-Asp triad(6Davis R.C. Stahnke G. Wong H. Doolittle M.H. Ameis D. Will H. Schotz M.C. J. Biol. Chem. 1990; 265: 6291-6295Abstract Full Text PDF PubMed Google Scholar, 7Faustinella F. Smith L.C. Semenkovich C.F. Chan L. J. Biol. Chem. 1991; 266: 9481-9485Abstract Full Text PDF PubMed Google Scholar, 8Emmerich J. Beg O.U. Peterson J. Previato L. Brunzell J.D. Brewer Jr., H.B. Santamarina-Fojo S. J. Biol. Chem. 1992; 267: 4161-4165Abstract Full Text PDF PubMed Google Scholar). Conservation of disulfide bonds suggests a similar folding pattern(9Kirchgessner T.G. Svenson K.L. Lusis A.J. Schotz M.C. J. Biol. Chem. 1987; 262: 8463-8466Abstract Full Text PDF PubMed Google Scholar), and by homology to the reported three-dimensional structure of pancreatic lipase(10Winkler F.K. D'Arcy A. Hunziker W. Nature. 1990; 343: 771-774Crossref PubMed Scopus (1028) Google Scholar), LPL and HL may be organized into two distinct amino- and carboxyl-terminal domains. Both lipases are anchored to the capillary endothelium via glycosaminoglycans and can be released by intravenous administration of heparin(1Jackson R.L. Boyer P.D. The Enzymes. XVI. Academic Press, New York1983: 141-186Google Scholar, 11Brunzell J.D. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill Book Co., New York1989: 1165-1180Google Scholar).There are, however, striking differences between LPL and HL. LPL is primarily synthesized by adipocytes, muscle cells, and macrophages, whereas HL mRNA is primarily detected in hepatocytes(12Semenkovich C.F. Chen S.H. Wims M. Luo C.C. Li W.H. Chan L. J. Lipid Res. 1989; 30: 423-431Abstract Full Text PDF PubMed Google Scholar). LPL is inhibited by 1 M NaCl (11Brunzell J.D. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill Book Co., New York1989: 1165-1180Google Scholar) and requires its cofactor, apoC-II, for full activation(13LaRosa J.C. Levy R.I. Herbert P. Lux S.E. Fredrickson D.S. Biochem. Biophys. Res. Commun. 1970; 41: 57-62Crossref PubMed Scopus (450) Google Scholar, 14Havel R.J. Fielding C.J. Olivecrona T. Shore V.G. Fielding P.E. Egelrud T. Biochemistry. 1973; 12: 1828-1833Crossref PubMed Scopus (246) Google Scholar). HL, on the other hand, is fully active even in the presence of high salt concentrations and in the absence of a cofactor, although its activity may be modulated by apoA-II (15Jahn C.E. Osborne Jr., J.C. Schaefer E.J. Brewer Jr., H.B. Eur. J. Biochem. 1983; 131: 25-29Crossref PubMed Scopus (65) Google Scholar, 16Mowri H.-O. Patsch W. Smith L.C. Gotto Jr., A.M. Patsch J.R. J. Lipid Res. 1992; 33: 1269-1279Abstract Full Text PDF PubMed Google Scholar) and apoE(17Thuren T. Wilcox R.W. Sisson P. Waite M. J. Biol. Chem. 1991; 266: 4853-4861Abstract Full Text PDF PubMed Google Scholar, 18Thuren T. Sisson P. Waite M. Biochim. Biophys. Acta. 1991; 1083: 217-220Crossref PubMed Scopus (25) Google Scholar, 19Thuren T. Weisgraber K.H. Sisson P. Waite M. Biochemistry. 1992; 31: 2332-2338Crossref PubMed Scopus (64) Google Scholar).The two enzymes also differ in their substrate specificity, demonstrating preferences in their fatty acid positional specificity (20Morley N. Kuksis A. J. Biol. Chem. 1972; 247: 6389-6393Abstract Full Text PDF PubMed Google Scholar, 21Wilcox R.W. Thuren T. Sisson P. Kucera G.L. Waite M. Lipids. 1991; 26: 283-288Crossref PubMed Scopus (9) Google Scholar), fatty acid chain length(22Deckelbaum R.J. Hamilton J.A. Moser A. Bengtsson-Olivecrona G. Butbul E. Carpentier Y.A. Gutman A. Olivecrona T. Biochemistry. 1990; 29: 1136-1142Crossref PubMed Scopus (145) Google Scholar, 23Masuno H. Okuda H. Biochim. Biophys. Acta. 1986; 879: 339-344Crossref PubMed Scopus (6) Google Scholar), and degree of fatty acid saturation(24Wang C.S. Kuksis A. Manganaro F. Lipids. 1982; 17: 278-284Crossref PubMed Scopus (48) Google Scholar, 25Miller C.H. Parce J.W. Sisson P. Waite M. Biochim. Biophys. Acta. 1981; 665: 385-392Crossref PubMed Scopus (22) Google Scholar). In addition, LPL and HL differ in their preferred lipoprotein substrates. Thus, patients with LPL deficiency accumulate primarily chylomicrons as well as very low density lipoproteins in plasma(11Brunzell J.D. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill Book Co., New York1989: 1165-1180Google Scholar), whereas HL deficiency results in elevated plasma concentrations of intermediate density lipoproteins and HDL(26Connelly P.W. Maguire G.F. Lee M. Little J.A. Arteriosclerosis. 1990; 10: 40-48Crossref PubMed Scopus (96) Google Scholar, 27Demant T. Carlson L.A. Holmquist L. Karpe F. Nilsson-Ehle P. Packard C.J. Shepherd J. J. Lipid Res. 1988; 29: 1603-1611Abstract Full Text PDF PubMed Google Scholar). Characterization of these patients demonstrates that in vivo, the preferred substrate of LPL are the large triglyceride-rich lipoproteins, whereas HL is more active in the hydrolysis of smaller lipoproteins such as HDL(28Kuusi T. Kinnunen P.K. Nikkila E.A. FEBS Lett. 1979; 104: 384-388Crossref PubMed Scopus (158) Google Scholar, 29Jansen H. van Tol A. Hulsmann W.C. Biochem. Biophys. Res. Commun. 1980; 92: 53-59Crossref PubMed Scopus (189) Google Scholar), especially HDL2(30Shirai K. Barnhart R.L. Jackson R.L. Biochem. Biophys. Res. Commun. 1981; 100: 591-599Crossref PubMed Scopus (152) Google Scholar), and intermediate density lipoproteins (26Connelly P.W. Maguire G.F. Lee M. Little J.A. Arteriosclerosis. 1990; 10: 40-48Crossref PubMed Scopus (96) Google Scholar, 31Nozaki S. Kubo M. Sudo H. Matsuzawa Y. Tarui S. Metab. Clin. Exp. 1986; 35: 53-58Abstract Full Text PDF PubMed Scopus (33) Google Scholar, 32Auwerx J.H. Marzetta C.A. Hokanson J.E. Brunzell J.D. Arteriosclerosis. 1989; 9: 319-325Crossref PubMed Google Scholar). Another physiologically important difference in the function of HL and LPL is the relative phospholipase versus triacylglycerol hydrolase activity of the two enzymes. Several studies have demonstrated that, relative to triglyceride hydrolysis, HL is a more active phospholipase than LPL(1Jackson R.L. Boyer P.D. The Enzymes. XVI. Academic Press, New York1983: 141-186Google Scholar, 33Ehnholm C. Shaw W. Greten H. Brown W.V. J. Biol. Chem. 1975; 250: 6756-6761Abstract Full Text PDF PubMed Google Scholar, 34van Tol A. van Gent T. Jansen H. Biochem. Biophys. Res. Commun. 1980; 94: 101-108Crossref PubMed Scopus (64) Google Scholar, 35Deckelbaum R.J. Ramakrishnan R. Eisenberg S. Olivecrona T. Bengtsson-Olivecrona G. Biochemistry. 1992; 31: 8544-8551Crossref PubMed Scopus (65) Google Scholar). In fact, inhibition of HL by infusion of antisera in cynomolgus monkeys led to a marked accumulation of not only triglycerides, but also of phospholipids in different lipoprotein fractions, especially HDL(36Goldberg I.J. Le N.A. Paterniti Jr., J.R. Ginsberg H.N. Lindgren F.T. Brown W.V. J. Clin. Invest. 1982; 70: 1184-1192Crossref PubMed Scopus (174) Google Scholar). Similar changes in plasma phospholipid levels have been observed in some patients with HL deficiency(32Auwerx J.H. Marzetta C.A. Hokanson J.E. Brunzell J.D. Arteriosclerosis. 1989; 9: 319-325Crossref PubMed Google Scholar), but not with LPL deficiency(37Faustinella F. Chang A. Van Biervliet J.P. Rosseneu M. Vinaimont N. Smith L.C. Chen S.-H. Chan L. J. Biol. Chem. 1991; 266: 14418-14424Abstract Full Text PDF PubMed Google Scholar, 38Demant T. Gaw A. Watts G.F. Durrington P. Buckley B. Imrie C.W. Wilson C. Packard C.J. Shepherd J. J. Lipid Res. 1993; 34: 147-156Abstract Full Text PDF PubMed Google Scholar). This difference in relative phospholipase to triacylglycerol hydrolase activity of the two enzymes may be important for modulating the function of LPL and HL in vivo.Recent studies involving the analysis of chimeric LPL-HL mutants (39Wong H. Davis R.C. Nikazy J. Seebart K.E. Schotz M.C. Proc. Natl. Acad. Sci. U. S. A. 1991; 88: 11290-11294Crossref PubMed Scopus (57) Google Scholar, 40Dichek H.L. Parrott C. Ronan R. Brunzell J.D. Brewer Jr., H.B. Santamarina-Fojo S. J. Lipid Res. 1993; 34: 1393-1401Abstract Full Text PDF PubMed Google Scholar, 41Davis R.C. Wong H. Nikazy J. Wang K. Han Q. Schotz M.C. J. Biol. Chem. 1992; 267: 21499-21504Abstract Full Text PDF PubMed Google Scholar) have suggested that the COOH-terminal domains of LPL and HL (Fig. 1) may play a role in determining the preferred substrate of the respective lipase(42Dugi K.A. Dichek H.L. Talley G.D. Brewer Jr., H.B. Santamarina-Fojo S. J. Biol. Chem. 1992; 267: 25086-25091Abstract Full Text PDF PubMed Google Scholar). In addition, removal of the COOH-terminal 58 amino acids of LPL by chymotryptic cleavage resulted in the inability of LPL to bind to chylomicrons(43Lookene A. Bengtsson-Olivecrona G. Eur. J. Biochem. 1993; 213: 185-194Crossref PubMed Scopus (44) Google Scholar), suggesting that this region of the enzyme may mediate binding to lipoproteins. However, to date, the structural basis for the differences in substrate specificity between the two lipases remains poorly defined. We have recently demonstrated that the lid covering the catalytic domain of the human lipases (Fig. 1) is critically involved in the interaction of the lipases with their lipid substrate(42Dugi K.A. Dichek H.L. Talley G.D. Brewer Jr., H.B. Santamarina-Fojo S. J. Biol. Chem. 1992; 267: 25086-25091Abstract Full Text PDF PubMed Google Scholar). In this report, we investigate the role of the lipase lid in determining the substrate specificity of HL and LPL for phospholipids and triglycerides by generating four different chimeric lipases in which the LPL and HL lids have been exchanged. Our studies demonstrate that the LPL lid enhances triglyceride hydrolysis, whereas the HL lid augments phospholipase function, thus establishing the important role of the lid in mediating lipase substrate specificity.MATERIALS AND METHODScDNA Expression VectorThe parent plasmid (pCMV) used for site-directed mutagenesis and transfection is a pUC18-derived vector containing the cytomegalovirus immediate early promoter and the polyadenylation site of SV40 as described previously(44Beg O.U. Meng M.S. Skarlatos S.I. Previato L. Brunzell J.D. Brewer Jr., H.B. Fojo S.S. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 3474-3478Crossref PubMed Scopus (55) Google Scholar). A 1473-base pair fragment of normal human LPL cDNA (pCMV-NLPL) (45Wion K.L. Kirchgessner T.G. Lusis A.J. Schotz M.C. Lawn R.M. Science. 1987; 235: 1638-1641Crossref PubMed Scopus (349) Google Scholar) or a 1522-base pair fragment of normal human HL cDNA (pCMV-NHL) (46Datta S. Luo C.C. Li W.H. VanTuinen P. Ledbetter D.H. Brown M.A. Chen S.H. Liu S. Chan L. J. Biol. Chem. 1988; 263: 1107-1110Abstract Full Text PDF PubMed Google Scholar, 47Stahnke G. Sprengel R. Augustin J. Will H. Differentiation. 1987; 35: 45-52Crossref PubMed Scopus (64) Google Scholar, 48Martin G.A. Busch S.J. Meredith G.D. Cardin A.D. Blankenship D.T. Mao S.J.T. Rechtin A.E. Woods C.W. Racke M.M. Schafer M.P. Fitzgerald M.C. Burke D.M. Flanagan M.A. Jackson R.L. J. Biol. Chem. 1988; 263: 10907-10914Abstract Full Text PDF PubMed Google Scholar) was cloned into the XbaI and HpaI restriction sites of pCMV. The DNA sequence of each fragment, which spanned the signal peptide through the termination codon, was confirmed by DNA sequence analysis using the dideoxynucleotide chain termination method (49Sanger F. Nicklen S. Coulson A.R. Proc. Natl. Acad. Sci. U. S. A. 1977; 74: 5463-5467Crossref PubMed Scopus (52355) Google Scholar) and T7 DNA polymerase (Sequenase, U. S. Biochemical Corp.).Synthesis of Mutant cDNAThe mutant HL and LPL cDNAs were synthesized by the overlap extension polymerase chain reaction (50Ho S.N. Hunt H.D. Horton R.M. Pullen J.K. Pease L.R. Gene (Amst.). 1989; 77: 51-59Crossref PubMed Scopus (6797) Google Scholar) using either pCMV-NLPL or pCMV-NHL as template. Polymerase chain reaction was performed in an automated DNA thermal cycler (Perkin-Elmer) as described (51Saiki R.K. Gelfand D.H. Stoffel S. Scharf S.J. Higuchi R. Horn G.T. Mullis K.B. Erlich H.A. Science. 1988; 239: 487-491Crossref PubMed Scopus (13398) Google Scholar) utilizing DNA polymerase from Pyrococcus furiosus (Stratagene Inc., La Jolla, CA) and 30 cycles with 1-min denaturation at 95°C, 1-min annealing at 50°C, and 2-min extension at 72°C in 1 × buffer 2 (Stratagene Inc.), 200 μM each dATP, dCTP, dGTP, and dTTP (Boehringer Mannheim), and 0.5 μM each primer. The exchange of the lids was performed with partially complementary oligonucleotide primers spanning the entire noncomplementary region of the 22-amino acid lid. Generation of chimeric proteins by domain exchange was performed as described previously(40Dichek H.L. Parrott C. Ronan R. Brunzell J.D. Brewer Jr., H.B. Santamarina-Fojo S. J. Lipid Res. 1993; 34: 1393-1401Abstract Full Text PDF PubMed Google Scholar). The mutant cDNAs were subcloned into the pCMV expression vector and amplified using competent DH5α cells (Life Technologies Inc.). Clones carrying the mutant cDNA were grown overnight at 37°C in LB broth (Biofluids, Inc., Rockville, MD), and DNA was isolated by one-tube minipreparation(52Del Sal G. Mantioletti G. Schneider C. Nucleic Acids Res. 1988; 16: 9878-9880Crossref PubMed Scopus (229) Google Scholar). All constructs were examined by sequence analysis of the complete cDNA insert. Oligonucleotide primers for overlap extension polymerase chain reaction and sequencing were synthesized by the phosphoramidite method on a DNA synthesizer (Model 380B, Applied Biosystems, Inc., Foster City, CA).In Vitro Expression of cDNA in Human Embryonal Kidney 293 CellsPlasmids used for transfection were purified by the cesium chloride double-banding method(53Radloff R. Bauer W. Vinograd J. Proc. Natl. Acad. Sci. U. S. A. 1967; 57: 1514-1521Crossref PubMed Scopus (856) Google Scholar). Transfections were performed using the calcium phosphate coprecipitation method (54Chen C. Okayama H. Mol. Cell. Biol. 1987; 7: 2745-2752Crossref PubMed Scopus (4809) Google Scholar) by adding 40 μg of plasmid DNA to each 100-mm plate of subconfluent human embryonal kidney 293 cells (American Type Culture Collection, Rockville, MD). Twenty-four hours after addition of DNA, the cells were washed, and medium containing 10% (v/v) fetal calf serum and 2 units/ml heparin sodium (Elkins-Sinn, Cherry Hill, NJ) was added. Medium for activity determination was harvested 12-16 h after washing and supplemented with glycerol to a final concentration of 30% (v/v). Intracellular protein was harvested as described by Chait et al.(55Chait A. Iverius P.H. Brunzell J.D. J. Clin. Invest. 1982; 69: 490-493Crossref PubMed Scopus (117) Google Scholar). Aliquots of media and intracellular extracts were kept at −70°C until lipase assays were performed. Each plasmid was transfected in triplicate. Wild-type HL and LPL were used as positive controls, and the pCMV vector without insert was used as negative control.Determination of HL and LPL ActivitiesEsterase activity was quantitated in triplicate using [14C]tributyrin(56Shirai K. Saito Y. Yoshida S. Biochim. Biophys. Acta. 1984; 795: 9-14Crossref PubMed Scopus (13) Google Scholar), and triglyceride lipase activity was determined in triplicate using [14C]triolein as previously published(57Iverius P.H. Brunzell J.D. Am. J. Physiol. 1985; 249: E107-E114PubMed Google Scholar). Phospholipase activities were measured in triplicate utilizing three different substrates. Phospholipid vesicles were generated by modifying the synthesis of triolein emulsion (57Iverius P.H. Brunzell J.D. Am. J. Physiol. 1985; 249: E107-E114PubMed Google Scholar) as follows. Dioleoylphosphatidylcholine (DOPC) (1 mM; Sigma) was used instead of egg yolk extract. Labeled triolein was substituted with [14C]DOPC (Amersham Corp.) at an activity of 0.1 μCi/ml of substrate. Proteoliposomes were synthesized according to a previously published protocol (58Chen C.H. Albers J.J. J. Lipid Res. 1982; 23: 680-691Abstract Full Text PDF PubMed Google Scholar) with the following modifications. DOPC was used as the unlabeled phospholipid, and the proteoliposomes were labeled with 1 μCi of [14C]DOPC. One-hundred microliters of substrate were added to 200 μl of medium from transfected cells in a total volume of 500 μl (0.15 M NaCl, 0.1 M Tris-HCl, pH 8.5, 2.5% bovine serum albumin, 25 μl of human plasma (as source of apoC-II), 2 units/ml heparin). Samples were incubated at 37°C in a shaking water bath for 1-4 h, and oleic acid was extracted by the method of Belfrage and Vaughan (59Belfrage P. Vaughan M. J. Lipid Res. 1969; 10: 341-344Abstract Full Text PDF PubMed Google Scholar). Mixed liposome substrate was prepared by a modification of a previously published protocol (60Rojas C. Olivecrona T. Bengtsson-Olivecrona G. Eur. J. Biochem. 1991; 197: 315-321Crossref PubMed Scopus (30) Google Scholar) using unlabeled DOPC, unlabeled triolein (Sigma), and 2 μCi of [14C]DOPC. Assay conditions used were the same as in the proteoliposome protocol.Determination of LPL MassLPL mass was determined six times by an enzyme-linked immunosorbent assay using the 5D2 monoclonal antibody (kindly provided by Dr. J. D. Brunzell, University of Washington, Seattle) for capture and a chicken polyclonal antibody (kindly provided by Dr. I. J. Goldberg, Columbia University, New York) for measurement.RESULTSTo evaluate a potential role of the lipase lid in modulating the substrate specificities of LPL and HL, we first established the importance of the lid region in the hydrolysis of different lipase substrates. We had previously demonstrated that the integrity of the amphipathic helices in the lipase lid is essential for the hydrolysis of water-insoluble long chain fatty acid triglycerides(42Dugi K.A. Dichek H.L. Talley G.D. Brewer Jr., H.B. Santamarina-Fojo S. J. Biol. Chem. 1992; 267: 25086-25091Abstract Full Text PDF PubMed Google Scholar). In a similar manner, we now investigated the role of the amphipathic helices in the hydrolysis of water-insoluble phospholipid substrates. Table 1 summarizes the concentration as well as activity in the medium of cells transfected with wild-type LPL, wild-type HL, or lipase-lid mutant plasmids. Disruption of the amphipathicity of helix 1 or helix 2 of the lid or deletion of the helices destroyed the ability of the lipases to hydrolyze water-insoluble triglycerides, while the esterase activity remained intact. Extensive rearrangement of both helices in HL and LPL markedly reduced esterase activity, possibly by reducing the ability of the lid to change to the open conformation upon interfacial activation(42Dugi K.A. Dichek H.L. Talley G.D. Brewer Jr., H.B. Santamarina-Fojo S. J. Biol. Chem. 1992; 267: 25086-25091Abstract Full Text PDF PubMed Google Scholar). The tributyrin activity, however, was still 5 times above background, whereas the more sensitive triolein assay revealed absent activity. These results indicate that the disruption of both helices in HL and LPL selectively prevented the hydrolysis of liposoluble substrate. Similarly, disruption of the amphipathic properties of the lipase lid abolished the ability of all mutant enzymes to hydrolyze phospholipid substrate presented as a DOPC vesicle (Table 1). Taken together, these studies demonstrate that the lipase lid is essential for the hydrolysis of not only liposoluble triglycerides, but also of liposoluble phospholipid substrates.Tabled 1View Large Image Figure ViewerDownload Hi-res image Download (PPT) Open table in a new tab To investigate a potential role of the lid in conferring substrate specificity, we generated several mutant lipases. As illustrated in Fig. 2, in Mut I, the lid of LPL was replaced by the lid of HL, and in the reciprocal mutant (Mut II), the lid of HL was replaced by the lid of LPL. Mut III is a chimeric lipase in which the carboxyl-terminal 134 amino acids of human LPL were replaced by the carboxyl-terminal 146 amino acids of human HL. Mut IV is a chimeric lipase that contains the NH2-terminal domain of LPL and the COOH-terminal domain as well as the lid of HL. In addition to these four chimeric constructs, plasmids containing wild-type LPL and HL cDNAs and a negative control consisting of the parent vector were transfected into 293 cells. Greater than 95% of the total triolein hydrolyzing activity was found in the culture medium of all transfected cells, indicating that the mutant lipases were secreted to a similar extent as wild-type LPL and HL (data not shown).Figure 2Schematic representation of wild-type and mutant LPL and HL constructs used for the analysis of substrate specificity. The name as well as a general description of the constructs utilized in this study are listed. LPL sequences are shown in white, and HL sequences are shown in black. The numbers indicate the amino acid residues of the particular lipase comprising the NH2-terminal domain of the respective construct.View Large Image Figure ViewerDownload Hi-res image Download (PPT)Table 2 summarizes the tributyrin, triolein, and phospholipid hydrolyzing activities in the medium of 293 cells transfected with the different plasmids. Tributyrin is water-soluble at the concentrations used in the assay and thus measures the esterase function of the lipases. Table 2 shows that substitution of the COOH-terminal domain of LPL with that of HL (Mut III) leads to a parallel reduction of tributyrin and triolein activities compared with LPL WT possibly due to a destabilization of the active dimer(40Dichek H.L. Parrott C. Ronan R. Brunzell J.D. Brewer Jr., H.B. Santamarina-Fojo S. J. Lipid Res. 1993; 34: 1393-1401Abstract Full Text PDF PubMed Google Scholar). Comparison of Mut I with LPL WT, Mut II with HL WT, and Mut IV with Mut III indicates that despite the exchange of the lids, the catalytic domain and esterase function of the mutant lipases are preserved. Further analysis reveals that the presence of the HL lid (Mut I versus LPL WT and Mut IV versus Mut III) increased phospholipid and lowered triolein hydrolysis, whereas the presence of the LPL lid (Mut II versus HL WT and Mut III versus Mut IV) had the opposite effect.Tabled 1View Large Image Figure ViewerDownload Hi-res image Download (PPT) Open table in a new tab Similar results were obtained when the triacylglycerol hydrolase and phospholipase activities of the different lipases were directly compared after normalization for tributyrin (esterase) activity (Fig. 3). The ability of wild-type LPL and HL as well as Mut I-IV to hydrolyze triglyceride (triolein) versus phospholipid (DOPC vesicles, DOPC proteoliposomes, and DOPC-mixed liposomes) substrates is summarized in Fig. 3(A-E). Replacement of the LPL lid with the lid of HL (Mut I) resulted in a 51% reduction in the ability of mutant LPL to hydrolyze triolein as well as a 317-618% increase in the phospholipid hydrolysis relative to that of wild-type LPL (Fig. 3A). Conversely, replacement of the HL lid with that of LPL (Mut II) led to a 23% increase in triolein hydrolysis and a reduction of phospholipase activity to <30% of wild-type HL (Fig. 3B).Figure 3Triolein and DOPC hydrolyzing activities of wild-type and mutant lipases. Activities are normalized for esterase (tributyrin hydrolyzing) activity and presented as percent of wild-type lipase activity. Wild-type lip
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