Rapid Subunit Exchange in Dimeric Lipoprotein Lipase and Properties of the Inactive Monomer
2004; Elsevier BV; Volume: 279; Issue: 48 Linguagem: Inglês
10.1074/jbc.m407419200
ISSN1083-351X
AutoresAivar Lõokene, Liyan Zhang, Magnus Hultin, Gunilla Olivecrona,
Tópico(s)Cholesterol and Lipid Metabolism
ResumoLipoprotein lipase (LPL), a key enzyme in the metabolism of triglyceride-rich plasma lipoproteins, is a homodimer. Dissociation to monomers leads to loss of activity. Evidence that LPL dimers rapidly exchange subunits was demonstrated by fluorescence resonance energy transfer between lipase subunits labeled with Oregon Green and tetrametylrhodamine, respectively, and also by formation of heterodimers composed of radiolabeled and biotinylated lipase subunits captured on streptavidine-agarose. Compartmental modeling of the inactivation kinetics confirmed that rapid subunit exchange must occur. Studies of activity loss indicated the existence of a monomer that can form catalytically active dimers, but this intermediate state has not been possible to isolate and remains hypothetical. Differences in solution properties and conformation between the stable but catalytically inactive monomeric form of LPL and the active dimers were studied by static light scattering, intrinsic fluorescence, and probing with 4,4′-dianilino-1,1′-binaphtyl-5,5′-disulfonic acid and acrylamide. The catalytically inactive monomer appeared to have a more flexible and exposed structure than the dimers and to be more prone to aggregation. By limited proteolysis the conformational changes accompanying dissociation of the dimers to inactive monomers were localized mainly to the central part of the subunit, probably corresponding to the region for subunit interaction. Lipoprotein lipase (LPL), a key enzyme in the metabolism of triglyceride-rich plasma lipoproteins, is a homodimer. Dissociation to monomers leads to loss of activity. Evidence that LPL dimers rapidly exchange subunits was demonstrated by fluorescence resonance energy transfer between lipase subunits labeled with Oregon Green and tetrametylrhodamine, respectively, and also by formation of heterodimers composed of radiolabeled and biotinylated lipase subunits captured on streptavidine-agarose. Compartmental modeling of the inactivation kinetics confirmed that rapid subunit exchange must occur. Studies of activity loss indicated the existence of a monomer that can form catalytically active dimers, but this intermediate state has not been possible to isolate and remains hypothetical. Differences in solution properties and conformation between the stable but catalytically inactive monomeric form of LPL and the active dimers were studied by static light scattering, intrinsic fluorescence, and probing with 4,4′-dianilino-1,1′-binaphtyl-5,5′-disulfonic acid and acrylamide. The catalytically inactive monomer appeared to have a more flexible and exposed structure than the dimers and to be more prone to aggregation. By limited proteolysis the conformational changes accompanying dissociation of the dimers to inactive monomers were localized mainly to the central part of the subunit, probably corresponding to the region for subunit interaction. Lipoprotein lipase (LPL) 1The abbreviations used are: LPL, lipoprotein lipase; bis-ANS, 4,4′-dianilino-1,1′-binaphtyl-5,5′-disulfonic acid; bis-Tris, 2-[bis(2-hydroxyethyl)amino]-2-(hydroxymethyl)propane-1,3-diol; BSA, bovine serum albumin; endo, endoproteinase; LDL, low density lipoprotein; MALDI-TOF, matrix-assisted laser desorption/ionization time-of-flight; OG, Oregon Green 488 carboxylic acid, succinimidyl ester; TMRA, 5-carboxytetramethylrhodamine succinimidyl ester; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine; VLDL, very low density lipoprotein. plays a fundamental role in the metabolism of blood lipoproteins (1Merkel M. Eckel R.H. Goldberg I.J. J. Lipid Res. 2002; 43: 1997-2006Abstract Full Text Full Text PDF PubMed Scopus (453) Google Scholar, 2Mead J.R. Irvine S.A. Ramji D.P. J. Mol. Med. 2002; 80: 753-769Crossref PubMed Scopus (670) Google Scholar, 3Preiss-Landl K. Zimmermann R. Hammerle G. Zechner R. Curr. Opin. Lipidol. 2002; 13: 471-481Crossref PubMed Scopus (197) Google Scholar). Lowered LPL activity results in dyslipidemia (4Stein Y. Stein O. Atherosclerosis. 2003; 170: 1-9Abstract Full Text Full Text PDF PubMed Scopus (73) Google Scholar). The catalytically active form of LPL is a noncovalent homodimer of 55 kDa subunits (5Iverius P.-H. Östlund-Lindqvist A.M. J. Biol. Chem. 1976; 251: 7791-7795Abstract Full Text PDF PubMed Google Scholar, 6Osborne Jr., J.C. Bengtsson-Olivecrona G. Lee N.S. Olivecrona T. Biochemistry. 1985; 24: 5606-5611Crossref PubMed Scopus (121) Google Scholar). The functional site of LPL is at the luminal surface of the vascular endothelium, where the enzyme is bound to heparan sulfate proteoglycans (1Merkel M. Eckel R.H. Goldberg I.J. J. Lipid Res. 2002; 43: 1997-2006Abstract Full Text Full Text PDF PubMed Scopus (453) Google Scholar, 2Mead J.R. Irvine S.A. Ramji D.P. J. Mol. Med. 2002; 80: 753-769Crossref PubMed Scopus (670) Google Scholar, 3Preiss-Landl K. Zimmermann R. Hammerle G. Zechner R. Curr. Opin. Lipidol. 2002; 13: 471-481Crossref PubMed Scopus (197) Google Scholar, 4Stein Y. Stein O. Atherosclerosis. 2003; 170: 1-9Abstract Full Text Full Text PDF PubMed Scopus (73) Google Scholar). Inactive monomers are less tightly bound to heparan sulfate (7Lookene A. Chevreuil O. Østergaard P. Olivecrona G. Biochemistry. 1996; 35: 12155-12163Crossref PubMed Scopus (113) Google Scholar) and are therefore the dominant form of LPL in the circulating blood (8Tornvall P. Olivecrona G. Karpe F. Hamsten A. Olivecrona T. Arterioscler. Thromb. Vasc. Biol. 1995; 15: 1086-1093Crossref PubMed Scopus (150) Google Scholar). In vitro studies show that the dimeric state of LPL is unstable (9Olivecrona T. Bengtsson G. Borgström B. Brockman H. Lipases. Elsevier, Amsterdam1984: 205-261Google Scholar). Under physiological conditions (pH, temperature, and concentration of salt), the dimers spontaneously dissociate into monomers within a few minutes. The formed monomers may reassociate but into inactive aggregates (9Olivecrona T. Bengtsson G. Borgström B. Brockman H. Lipases. Elsevier, Amsterdam1984: 205-261Google Scholar). It is believed that this process is irreversible. The inactivation is strongly prevented by heparan sulfate and heparin (7Lookene A. Chevreuil O. Østergaard P. Olivecrona G. Biochemistry. 1996; 35: 12155-12163Crossref PubMed Scopus (113) Google Scholar). The dimeric form of LPL is also relatively stable in the presence of high concentrations of salt (at low temperature) (9Olivecrona T. Bengtsson G. Borgström B. Brockman H. Lipases. Elsevier, Amsterdam1984: 205-261Google Scholar, 10van Tilbeurgh H. Roussel A. Lalouel J.-M. Cambillau C. J. Biol. Chem. 1994; 269: 4626-4633Abstract Full Text PDF PubMed Google Scholar). In the early 1970s this discovery allowed purification of the active enzyme from bovine milk by using adsorption to heparin-Sepharose followed by elution with a salt gradient (5Iverius P.-H. Östlund-Lindqvist A.M. J. Biol. Chem. 1976; 251: 7791-7795Abstract Full Text PDF PubMed Google Scholar, 11Egelrud T. Olivecrona T. J. Biol. Chem. 1972; 247: 6212-6217Abstract Full Text PDF PubMed Google Scholar). Osborne et al. (6Osborne Jr., J.C. Bengtsson-Olivecrona G. Lee N.S. Olivecrona T. Biochemistry. 1985; 24: 5606-5611Crossref PubMed Scopus (121) Google Scholar) showed that the inactivation rate is dependent on the concentration of LPL. On this basis, they proposed that inactivation of LPL includes a step of reversible dissociation of the dimer followed by a step of irreversible unfolding of the dissociated monomers (6Osborne Jr., J.C. Bengtsson-Olivecrona G. Lee N.S. Olivecrona T. Biochemistry. 1985; 24: 5606-5611Crossref PubMed Scopus (121) Google Scholar). LPL, pancreatic lipase, hepatic lipase, and endothelial lipase share sequence homologies and together form the mammalian triacylglycerol lipase gene family (12Wong H. Schotz M.C. J. Lipid Res. 2002; 43: 993-999Abstract Full Text Full Text PDF PubMed Scopus (241) Google Scholar). A model of the three-dimensional structure of LPL has been constructed based on the x-ray structure of pancreatic lipase (10van Tilbeurgh H. Roussel A. Lalouel J.-M. Cambillau C. J. Biol. Chem. 1994; 269: 4626-4633Abstract Full Text PDF PubMed Google Scholar). The model is supported by results obtained by other techniques, including limited proteolysis and mutagenesis (12Wong H. Schotz M.C. J. Lipid Res. 2002; 43: 993-999Abstract Full Text Full Text PDF PubMed Scopus (241) Google Scholar, 13Lookene A. Bengtsson-Olivecrona G. Eur. J. Biochem. 1993; 213: 185-194Crossref PubMed Scopus (44) Google Scholar, 14Santamarina-Fojo S. Dugi K.A. Curr. Opin. Lipidol. 1994; 5: 117-125Crossref PubMed Scopus (105) Google Scholar). Thus, the overall three-dimensional structures of the lipases appears to be quite similar. In contrast to LPL, pancreatic lipase can exist as a catalytically active monomer (10van Tilbeurgh H. Roussel A. Lalouel J.-M. Cambillau C. J. Biol. Chem. 1994; 269: 4626-4633Abstract Full Text PDF PubMed Google Scholar). Thus, some important differences in folding or in conformational flexibility between these two proteins are expected. Because of instability and low solubility, the possibilities of using physical techniques for structural studies of LPL are limited. Little is known about the conformational differences between the active dimer and the catalytically inactive monomer. By continuous measurement of circular dichroism, under conditions in which the enzyme slowly lost its catalytic activity, a decrease in the content of secondary structure has been demonstrated (6Osborne Jr., J.C. Bengtsson-Olivecrona G. Lee N.S. Olivecrona T. Biochemistry. 1985; 24: 5606-5611Crossref PubMed Scopus (121) Google Scholar). Conformational differences between active and inactive forms of LPL are also implicated by differences in their abilities to react with monoclonal antibodies (15Peterson J. Fujimoto W.Y. Brunzell J.D. J. Lipid Res. 1992; 33: 1165-1170Abstract Full Text PDF PubMed Google Scholar). Inactive LPL monomers can be demonstrated by sucrose density gradient centrifugation (16Olivecrona G. Lookene A. Methods Enzymol. 1997; 286: 102-116Crossref PubMed Scopus (22) Google Scholar, 17Zhang L. Wu G. Tate C.G. Lookene A. Olivecrona G. J. Biol. Chem. 2003; 278: 29344-29351Abstract Full Text Full Text PDF PubMed Scopus (59) Google Scholar). The affinity of the monomer to heparin is ∼6000-fold weaker than that of the dimer (7Lookene A. Chevreuil O. Østergaard P. Olivecrona G. Biochemistry. 1996; 35: 12155-12163Crossref PubMed Scopus (113) Google Scholar), and the difference in heparin affinities can readily be used for separation of dimers and monomers by affinity chromatography on heparin columns (16Olivecrona G. Lookene A. Methods Enzymol. 1997; 286: 102-116Crossref PubMed Scopus (22) Google Scholar, 17Zhang L. Wu G. Tate C.G. Lookene A. Olivecrona G. J. Biol. Chem. 2003; 278: 29344-29351Abstract Full Text Full Text PDF PubMed Scopus (59) Google Scholar, 18Hata A. Ridinger D.N. Sutherland S.D. Emi M. Kwong L.K. Shuhua J. Lubbers A. Guy-Grand B. Basdevant A. Iverius P.-H. Wilson D.E. Lalouel J.-M. J. Biol. Chem. 1992; 267: 20132-20139Abstract Full Text PDF PubMed Google Scholar). Functional studies have shown that, although the monomers are catalytically inactive, they retain several properties of the active dimer. Monomers are still able to interact with lipoproteins (19Pentikäinen M.O. Oorni K. Kovanen P.T. J. Biol. Chem. 2000; 275: 5694-5701Abstract Full Text Full Text PDF PubMed Scopus (44) Google Scholar). In blood the monomers are mainly found together with LDL particles (20Vilella E. Joven J. Fernández M. Vilaró S. Brunzell J.D. Olivecrona T. Bengtsson-Olivecrona G. J. Lipid Res. 1993; 34: 1555-1564Abstract Full Text PDF PubMed Google Scholar). The monomer interacts, although with reduced affinity, with receptors of the LDL-receptor family (21Nykjær A. Bengtsson-Olivecrona G. Lookene A. Moestrup S.K. Petersen C.M. Weber W. Beisiegel U. Gliemann J. J. Biol. Chem. 1993; 268: 15048-15055Abstract Full Text PDF PubMed Google Scholar). Compared with dimeric LPL, the receptor-bound monomers are less effective in binding of lipoproteins, probably because the binding sites for lipoproteins and receptors partly overlap (21Nykjær A. Bengtsson-Olivecrona G. Lookene A. Moestrup S.K. Petersen C.M. Weber W. Beisiegel U. Gliemann J. J. Biol. Chem. 1993; 268: 15048-15055Abstract Full Text PDF PubMed Google Scholar). Altogether these data demonstrate that the inactive LPL monomer is still folded and rather stable. LPL monomers are found, for example, in blood (22Olivecrona G. Hultin M. Savonen R. Skottova N. Lookene A. Tugrul Y. Olivecrona T. Woodford F.P. Davignon J. Sniderman A.D. Atherosclerosis X. Elsevier, Amsterdam1995: 250-253Google Scholar) and adipose tissue (23Bergö M. Olivecrona G. Olivecrona T. Biochem. J. 1996; 313: 893-898Crossref PubMed Scopus (111) Google Scholar) and are also present in culture media from the expression of recombinant LPL in vitro (17Zhang L. Wu G. Tate C.G. Lookene A. Olivecrona G. J. Biol. Chem. 2003; 278: 29344-29351Abstract Full Text Full Text PDF PubMed Scopus (59) Google Scholar, 24Krapp A. Zhang H.F. Ginzinger D. Liu M.S. Lindberg A. Olivecrona G. Hayden M.R. Beisiegel U. J. Lipid Res. 1995; 36: 2362-2373Abstract Full Text PDF PubMed Google Scholar). It has been shown that the monomer/dimer ratio in adipose tissue is dependent on the nutritional state (23Bergö M. Olivecrona G. Olivecrona T. Biochem. J. 1996; 313: 893-898Crossref PubMed Scopus (111) Google Scholar). In the fasted state more LPL is found in the inactive form (25Bergö M. Wu G. Ruge T. Olivecrona T. J. Biol. Chem. 2002; 277: 11927-11932Abstract Full Text Full Text PDF PubMed Scopus (71) Google Scholar), and therefore less of the plasma lipids are taken up for storage, whereas more is used in other tissues for energy production. Studies with actinomycin D indicate that some other protein with a short-lived mRNA regulates the dissociation of LPL into inactive monomers or prevents inactive LPL subunits from forming active dimers (25Bergö M. Wu G. Ruge T. Olivecrona T. J. Biol. Chem. 2002; 277: 11927-11932Abstract Full Text Full Text PDF PubMed Scopus (71) Google Scholar, 26Wu G. Olivecrona G. Olivecrona T. J. Biol. Chem. 2003; 278: 11925-11930Abstract Full Text Full Text PDF PubMed Scopus (38) Google Scholar). The details of this important control mechanism are still unknown, but the fact remains that monomeric LPL is present in most tissues and that monomeric LPL is the dominant form of LPL in blood. Recent studies indicate that the levels of this form in plasma correlate to disease (27Hanyu O. Miida T. Obayashi K. Ikarashi T. Soda S. Kaneko S. Hirayama S. Suzuki K. Nakamura Y. Yamatani K. Aizawa Y. Atherosclerosis. 2004; 174: 385-390Abstract Full Text Full Text PDF PubMed Scopus (58) Google Scholar). Many single-point mutations in the LPL gene result in the production of inactive LPL monomers (12Wong H. Schotz M.C. J. Lipid Res. 2002; 43: 993-999Abstract Full Text Full Text PDF PubMed Scopus (241) Google Scholar, 18Hata A. Ridinger D.N. Sutherland S.D. Emi M. Kwong L.K. Shuhua J. Lubbers A. Guy-Grand B. Basdevant A. Iverius P.-H. Wilson D.E. Lalouel J.-M. J. Biol. Chem. 1992; 267: 20132-20139Abstract Full Text PDF PubMed Google Scholar). It is therefore important to investigate the properties of the folded monomer and to understand its relation to the active dimeric form of LPL. In the present investigation we have focused on mechanisms responsible for the spontaneous inactivation of LPL. We demonstrate that the subunits of dimeric LPL rapidly exchange partners. This may be a first step in the process that leads to inactivation of the enzyme. We have used compartmental modeling to study the inactivation mechanism, static light scattering to study aggregation, limited proteolysis to study differences in folding between inactive monomers and active dimmers, and bis-ANS to probe exposure of hydrophobic regions due to unfolding. The largest conformational changes on dissociation of the dimer were located to the middle of the LPL subunit and probably involve areas of subunit interactions. Materials—Bovine LPL was purified from milk (28Bengtsson-Olivecrona G. Olivecrona T. Methods Enzymol. 1991; 197: 345-356Crossref PubMed Scopus (100) Google Scholar). It was iodinated by the lactoperoxidase method and separated from free Na125I by chromatography on heparin-Sepharose as described (16Olivecrona G. Lookene A. Methods Enzymol. 1997; 286: 102-116Crossref PubMed Scopus (22) Google Scholar). The amino coupling kit and CM5 sensor chips were from BIAcore (Uppsala, Sweden). Heparin was from Leo Pharma AB (Malmö, Sweden). Oregon Green 488 carboxylic acid, succinimidyl ester (OG), 5-carboxytetramethylrhodamine, succinimidyl ester (TMRA), and bis-ANS (4,4′-dianilino-1,1′-binaphtyl-5,5′-disulfonic acid) were from Molecular Probes. LPL was biotinylated as described previously (7Lookene A. Chevreuil O. Østergaard P. Olivecrona G. Biochemistry. 1996; 35: 12155-12163Crossref PubMed Scopus (113) Google Scholar). Lipoproteins (very low density lipoproteins (VLDL) and low density lipoproteins (LDL) were isolated from fresh human plasma by sequential ultracentrifugation. Trypsin, chymotrypsin, endoproteinase Glu-C (endo-Glu-C), bovine serum albumin (BSA), streptavidin-agarose, tributyrin, gum Arabic, and acrylamide were from Sigma. Labeling of LPL with OG and TMRA—LPL (0.2–0.5 mg/ml) in 0.2 m NaHCO3, 1 m NaCl, pH 8.4, was incubated with OG or TMRA at 4 °C in the dark. After 1 h, the reaction was stopped by the addition of lysine to a final concentration of 1 mm. After 15 min, the mixture was diluted ∼5× with 20 mm bis-Tris, pH 6.5, and immediately applied onto a column of heparin-Sepharose. Labeled LPL was eluted by a linear gradient of NaCl in the bis-Tris buffer. Most of the active labeled LPL eluted with a peak at 1.1 m NaCl. The fractions with high LPL activity were pooled. The incorporation of OG and TMRA into LPL was characterized by the degree of labeling (DOL),DOL=Amax[LPL]×ϵdye where Amax is absorbance of OG-LPL or TMRA-LPL at their absorbance maximum wavelength, ϵdye is extinction coefficient of OG or TMRA, and [LPL] is the concentration of the labeled LPL. The ϵdye values used were 70,000 for OG and 65,000 for TMRA. These values were from the website of Molecular Probes. The concentration of LPL was calculated from absorbance, A280 (1%) = 1.68 (29Olivecrona T. Bengtsson G. Osborne Jr., J.C. Eur. J. Biochem. 1982; 124: 629-633Crossref PubMed Scopus (25) Google Scholar). Fluorescence and Light-scattering Measurements—These experiments were performed on a Fluoro-Max spectrofluorimeter. The conditions are detailed in the legends to Figs. 2, 5, and 6.Fig. 5Differences in inactivation curves when different assay systems were used.A, inactivation time courses recorded in parallel by using tributyrin (•) or Intralipid (○) as substrates. In both cases 8.5 nm LPL was incubated in a solution of 0.4 m NaCl, 1 mg BSA/ml, 10 mm Tris, pH 7.4, at 20 °C. The remaining activity is expressed in percent of LPL activity in a sample assayed immediately after dilution in ice-cold buffer. B, decrease of tryptophan fluorescence at 340 nm on excitation at 280 nm under the conditions used in the experiment in A but in the absence of BSA.View Large Image Figure ViewerDownload (PPT)Fig. 6Aggregation of LPL as studied by static light scattering. The experiments were carried out on a spectrophotofluorimeter. Both the excitation and emission wavelengths were set to 500 nm. The intensity of the scattered light was measured at a 90° angle to the direction of the excitation light. The relative values presented were obtained after subtraction of scattering intensity of the incubation solution without LPL. All measurements were performed on 8.5 nm LPL in solutions of 20 mm Tris, pH 7.4, at 22 °C, with the indicated additions.View Large Image Figure ViewerDownload (PPT) Binding to Streptavidin-Agarose—Equal volumes (0.1 ml) of radiolabeled LPL (0.05 mg/ml, 4000 cpm/μl) and biotinylated LPL (0.05–0.15 mg/ml) were incubated together for 1 min in 10 mm Tris, 0.5 m NaCl, 1 mg/ml BSA, pH 7.4 at 4 °C. Of this incubation mixture, 0.05 ml was added to 2 ml of the same buffer containing 0.3 ml of streptavidinagarose. The mixture was gently shaken for 5 min at 4 °C and centrifuged at 3000 × g for 5 min. The radioactivity was determined in the top phase (containing unbound LPL) and in the lower phase (containing sedimented streptavidin-agarose and the bound LPL). Data are expressed as ratios of radioactivity in the sedimented agarose to the total radioactivity added to the system. Nonspecific binding of radioactive LPL to streptavidin-agarose in the absence of biotinylated LPL was subtracted from the data. Binding Studies on BIAcore—Details concerning conditions for these studies are described previously (7Lookene A. Chevreuil O. Østergaard P. Olivecrona G. Biochemistry. 1996; 35: 12155-12163Crossref PubMed Scopus (113) Google Scholar, 30Lookene A. Savonen R. Olivecrona G. Biochemistry. 1997; 36: 5267-5275Crossref PubMed Scopus (67) Google Scholar). Experiments were performed on a BIAcore 2000 instrument. Biotinylated heparin or biotinylated LPL was bound to the matrix-coupled streptavidin. LDL and VLDL were then injected into these sensorchips in 10 mm Hepes, 0.15 m NaCl, pH 7.4, at 25 °C. Activity Measurements—Activity measurements with tributyrin as substrate were performed by continuous titration of butyric acid by a pH-stat (Methrome, Herisau, Switzerland) at 25 °C (13Lookene A. Bengtsson-Olivecrona G. Eur. J. Biochem. 1993; 213: 185-194Crossref PubMed Scopus (44) Google Scholar). Stock solutions of the substrate were prepared by sonication of tributyrin in a solution of gum Arabic/water (Soniprep, MSE Technology Application Inc.). The enzymatic activity was recorded at pH 8.5 in 0.15 m NaCl containing 30 mm tributyrin and 0.2% gum Arabic. Titration was performed with 25 mm KOH. LPL activity on phospholipid stabilized emulsions of long acyl chain triacylglycerols was measured as described in a previous study (23Bergö M. Olivecrona G. Olivecrona T. Biochem. J. 1996; 313: 893-898Crossref PubMed Scopus (111) Google Scholar). Briefly, 5 μl of each sample was incubated in 200 μl of a mixture containing a phospholipid-stabilized emulsion of soy bean triacylglycerols with the same composition as Intralipid 10% (Fresenius-KABI, Uppsala, Sweden), into which tri-[9,10-3H]oleoylglycerol had been incorporated on manufactory. Heat-inactivated rat serum was present as a source of apolipoprotein CII, BSA as fatty acid acceptor, and heparin to stabilize the lipase. One milliunit of lipase activity corresponded to the release of 1 nmol of fatty acid/min at 25 °C. All samples were assayed in triplicate. Mathematical Modeling of the Inactivation Process—Mathematical modeling of the transfer of LPL between different states was done using SAAMII (simultaneous analysis and modeling), version 1.1 (SAAM Institute, Seattle, WA). This software allows compartmental modeling of complex systems using linear as well as nonlinear models. The error of the data was set to a fractional standard deviation of 0.1. Proteolytic Cleavage—Limited cleavage of LPL by chymotrypsin and trypsin was carried out in 0.1 m Tris, 0.5 m NaCl, pH 7.4, at 10 °C. In the case of monomeric LPL the cleavage reactions also contained 1.0 m guanidinium chloride. The protease/LPL mass ratio was 5%, and the cleavage reactions were stopped by adding phenylmethylsulfonyl fluoride to a 10-fold molar excess over trypsin or chymotrypsin. Digestion by endo-Glu-C was performed in 0.1 m phosphate buffer, pH 7.8, with or without 1.0 m guanidinium chloride. The reactions were stopped by heating the samples at 90 °C for 15 min. The cleavage products were analyzed on Tricine-SDS-PAGE using 15.5% polyacrylamide gels. Mass Spectrometry and Sequence Analyses—Matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrometry was used to analyze the products of the proteolytic cleavage reactions. The analyses were performed on a home-made mass spectrometer at the National Institute of Chemical Physics and Biophysics (Tallinn, Estonia). The matrix dihydrobenzoic acid was dissolved in 1 ml of a 1:1 mixture of 0.1% trifluoroacetic acid and acetonitrile. A 0.5-μl aliquot of the cleavage mixture was deposited on a stainless steel probe. The mass determination was accurate enough only for peptides smaller than 10 kDa. For larger peptides, N-terminal sequences were determined on an Applied Biosystems (Foster City, CA) 477A pulsed liquid-phase sequencer. Monomer Exchange Kinetics Studied by Fluorescence Resonance Energy Transfer— To investigate whether the subunits of dimeric LPL are stably bound to each other or whether they rapidly exchange partners, we studied fluorescence resonance energy transfer between two differently fluorescence-labeled LPL dimers. One portion of LPL was labeled with OG and another portion with TMRA. Control experiments showed that both variants, named OG-LPL and TMRA-LPL, maintained 70–80% of the activity of the unmodified enzyme, that their activities were stimulated by apolipoprotein CII, and that they eluted from heparin-Sepharose at around 1.1 m NaCl, which corresponded to the elution pattern of unlabeled, dimeric LPL. BIAcore studies showed that both labeled forms of LPL were able to interact with lipoproteins (VLDL and LDL) similarly to unmodified LPL (data not shown). Taken together these results showed that the labeling of LPL by TMRA or OG did not change the functional properties of the dimeric LPL. Increasing the ratio of dye to LPL led to an increased degree of labeling at low ratios ( 4 mol/mol) the incorporation of label approached saturation (Fig. 1). The maximal degrees of labeling were similar, 2.1 mol/mol for the OG-LPL and 2.2 mol/mol for the TMRA-LPL. In the following experiments the labeling was between 1.9 and 2.1 mol/mol LPL dimer. When mixed together (Fig. 2A), TMRA-LPL significantly quenched the fluorescence emission of OG-LPL, as measured between 510 and 530 nm on excitation at 480 nm. A rapid decrease (half-life less than 5 s) in fluorescence of the OG-LPL variant was observed after the addition of TMRA-LPL, but after that the fluorescence was stable for hours (Fig. 2B). Fig. 2C presents the fluorescence spectra of OG-LPL in the presence of increasing concentrations of TMRA-LPL. The effect was concentration-dependent with a tendency to saturation at higher concentrations of TMRA-LPL (Fig. 2D). When TMRA-LPL was titrated with OG-LPL, the fluorescence of TMRA groups increased between 540 and 570 nm on excitation at 480 nm (data not shown). These observations demonstrated that fluorescence resonance energy transfer occurred between OG and TMRA groups when the labeled variants of LPL were mixed, suggesting formation of heterodimers of subunits from OG-LPL and TMRA-LPL, respectively. Fig. 2D demonstrates that fluorescence energy transfer did not occur when the experiments were carried out under denaturating conditions (in 1 or 6 m guanidinium chloride). The fluorescence resonance energy transfer between the labeled proteins was not affected by the presence of heparin (15 IU/ml) or by lipoproteins (LDL, 0.1 μg protein/ml) in the incubation mixture (data not shown). Subunit exchange between LPL dimers was also demonstrated by binding experiments to streptavidin-agarose. In this case 125I-labeled LPL was incubated with biotinylated LPL, and then streptavidin-agarose beds were added. After centrifugation, the radioactivity bound to the gel was determined. Despite the high nonspecific binding of 125I-labeled LPL to the matrix (20–30% of the total), there was a significant increase of radioactivity after mixing of the radiolabeled enzyme with biotinylated LPL (Table I). Because biotinylation or iodination has previously been shown not to affect the activity or the dimeric structure of LPL (16Olivecrona G. Lookene A. Methods Enzymol. 1997; 286: 102-116Crossref PubMed Scopus (22) Google Scholar), the increased binding of radioactivity to the gel suggested formation of heterodimers of subunits from 125I-labeled LPL and biotinylated LPL, respectively.Table IBinding of 125I-labeled LPL to streptavidin-agarose in the presence of different concentration of biotinylated LPLAdded biotin-LPLBound radioactivitymg/ml% of total0.0623 ± 50.1133 ± 70.1544 ± 6 Open table in a new tab Studies of Inactivation of LPL and Modeling of the Inactivation Process—Osborne et al. (6Osborne Jr., J.C. Bengtsson-Olivecrona G. Lee N.S. Olivecrona T. Biochemistry. 1985; 24: 5606-5611Crossref PubMed Scopus (121) Google Scholar) reported that the relative inactivation rate is faster at lower concentrations of LPL and proposed that the inactivation is a two-step process including a reversible dissociation of LPL dimers followed by an irreversible unfolding of the monomers. In the present study we revisited the inactivation of LPL with the aim of analyzing the inactivation process by compartmental modeling and also of obtaining estimates for rate constants. The inactivation kinetics was measured for a wide variation of LPL concentrations under different conditions (Fig. 3). The effect of the concentration of the LPL protein was observed only under conditions in which the enzyme was relatively stable, as in 0.3 m NaCl at 20 °C (Fig. 3A) or in 0.15 m NaCl with 15 IU of heparin/ml at 37 °C (Fig. 3B). In 0.15 m NaCl without heparin, where the inactivation rate was fast, the effect of LPL concentration was less pronounced (Fig. 3C). In this case, inactivation followed simple monoexponential decay kinetics. It was pos
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