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1 Dipartimento di Biotecnologie e Bioscienze, Universita’ degli Studi di Milano-Bicocca, 20126 Milano, Italy
2 Istituto di Chimica del Riconoscimento Molecolare, Consiglio Nazionale delle Ricerche, 20131 Milano, Italy
Reprint requests to: Marina Lotti, Dipartimento di Biotecnologie e Bioscienze, Universita’ di Milano-Bicocca, Piazza della Scienza 2, 20126 Milano, Italy; e-mail: marina.lotti{at}unimib.it; fax: ++39-02-6448-3565.
(RECEIVED January 24, 2003; FINAL REVISION July 12, 2003; ACCEPTED July 15, 2003)
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.0304003.
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Abstract |
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Keywords: Lipase; cholesterol esterase; lid; domain grafting; substrate recognition
Abbreviations: CMC, critical micellar concentration • CRLs, Candida rugosa lipases • TFE, 2,2,2-tifluoroethyl
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Introduction |
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In this work we sought to determine whether the lid might be responsible for different catalytic properties observed in isoforms of the same enzyme. The target of this study was the family of extracellular lipases produced by the Ascomycete Candida rugosa (CRLs). CRLs are the products of a family of related genes that code for proteins with over 80% identity in their amino acid sequence (Longhi et al. 1992; Lotti et al. 1993). The expression of lipase genes has been shown to be tuned by the conditions of growth and in particular by the carbon source present in the culture medium (Lotti et al. 1998; Lee et al. 1999). Crystallographic structures of CRL1 (Grochulski et al. 1993, 1994) and CRL3 (Ghosh et al. 1995) have been reported. Interestingly, and despite their close sequence relatedness, lipase isoforms purified by chromatography or expressed in heterologous systems have been shown to differ in substrate specificity, in particular with regard to substrate chain length (Rua et al. 1993; Brocca et al. 1998; Lopez et al. 2000; Tang et al. 2001; Lee et al. 2002). CRL3 displays high activity towards cholesterol esters and is therefore also known as cholesterol esterase (Ghosh et al. 1995). Comparison of the amino acid sequences reveals that variability among isoenzymes is clustered at the substrate binding sites and in the lid region, which interacts with the substrate molecules in the enzyme open conformation (Grochulski et al. 1993; Lotti et al. 1994).
In an effort to explore the contribution of the lid to the catalytic properties of lipase1 (CRL1, Brocca et al. 1998), we modified the recombinant enzyme by substituting its lid with the corresponding peptides from isoforms CRL3 and CRL4. Activity of the chimeric proteins was then investigated in the hydrolysis of soluble and insoluble substrates as well as in alcoholysis reactions in organic solvent (Carrea and Riva 2000). Activity towards cholesterol esters was conferred through CRL3 lid swapping. Moreover, chimeras were affected in their sensitivity to the reaction environment, suggesting that the lid may play a role in the dynamics of the transition between the closed and open enzyme conformations.
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Results |
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). In particular, LID4 is characterized by an enhanced ratio of hydrophobic residues, yet maintains its amphipathic nature. Protein engineering was performed on plasmid pPICslip1 containing a synthetic sequence coding for CRL1 fused to the 
-factor leader peptide (Brocca et al. 1998). To ease further manipulations, the coding sequence was transferred in pGEM-7Z because of the larger choice of unique restriction sites. To this purpose, a 1742-bp fragment encompassing the whole lip1 gene was excised by HindIII and BamH1 digestion and cloned in the polylinker sequence of plasmid pGEM7Z, yielding the construct pGEM-slip1. In this latter construct, the lid sequence is enclosed in a 541-bp fragment defined by two unique restriction sites for SmaI, located in the plasmid cloning sites and 42 bp downstream of the sequence of interest, respectively. Two cassettes were synthesized in order to replace the natural lid (LID1) with the corresponding sequences of isoform 3 (LID3) and 4 (LID4), using the experimental strategy described in Figure 2. The 541-bp sequence was subdivided into two regions: (1) a 5' half (378 bp) containing a constant sequence which encompasses part of the cloning plasmid and the -factor leader sequence; as this sequence did not need to be mutagenized, it was simply amplified by PCR, and (2) a 3' moiety with sequences unique to either LID3 or LID4 (163 bp) was enzymatically synthesized by PCR overlap extension using overlapping oligonucleotides 88–93-bp long. Being the 3'-end of the invariant half complementary to the 5'-half of the lid-containing fragment, a final step of PCR overlap extension allowed the reconstruction of whole cassettes that could be easily inserted in pGEM-slip1 restricted with SmaI. Sequencing confirmed that chimeric lipases had been constructed on the CRL1 scaffold with the lid replaced by the corresponding sequence in CRL3 or CRL4.
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Four clones of each transformant type were grown on a small scale (25 mL) in YEPS medium. Cultures were induced with methanol, and the presence of lipase in the culture medium and within the cells was assessed by Western blotting with anti-lipase antibodies (Fig. 3). Whereas CRL1-LID3 was secreted in amounts comparable to wild-type (wt) CRL1 (0.2 mg/mL culture broth), CRL1-LID4 was hardly detectable in the culture medium (0.1 µg/mL). However, high amounts of CRL1-LID4 were revealed intracellularly (not shown).
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Activity in aqueous medium
Hydrolytic activity on soluble substrates was measured on 1 mM p-nitrophenylesters of different chain length: p-n-phenylbutyrate (C4), p-n-phenylcaproate (C6), p-n-phenylcaprylate (C8), p-n-phenyllaurate (C12), p-n-phenylpalmitate (C16), and p-n-phenylstearate (C18). Reaction mixtures varied in their composition for the presence of different nonionic detergents (Table 1). Mixture 1 contained only the appropriate buffer, whereas mixture 2 was added with arabic gum and Triton X-100 well above its critical micellar concentration (CMC). In this latter case, only C4, C6, and C8 substrates were tested due to the poor solubility of longer-chain compounds in 0.5% Triton. In addition, three different detergents, namely Triton X-100 (3), Tween 20 (4), and Tween 80 (5) were used in concentrations below the respective CMC, as high detergent concentrations are often inhibitory for lipase activity (Helistö and Korpela 1998).
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, arrow). CRL1LID3 and CRL1LID4 both showed an increase in relative activity toward the C4 substrate compared to esters with longer acyl chain. However, no general trend in lid-dependent substrate preference was apparent. All lipases turned out to be extremely sensitive, but with varied responses, to the presence of detergents. Striking evidence of a differential effect of the detergent on enzymes differing in their lids emerged from the comparison of specific activities of mutant lipases under different experimental conditions, as presented in Figure 4. Due to the complexity of the results, and for the sake of clarity, we will point out only a few significant results. Thus, for example, from the comparison of the activity data obtained with and without detergents, it can be observed that Triton X-100 above its CMC inhibited wt CRL1 but activated the two chimeras on C8 substrates (Fig. 4, stars). In contrast, the activity of CRL1LID4 on the C18 substrate is drastically decreased by the presence of Tween 80 (Fig. 4, arrows).
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). This result clearly points to the lid as a major functional determinant of the cholesterol esterase activity reported for CRL3.
Activity in organic solvent
Table 3 reports the initial rates of alcoholysis of wt CRL1 and CRL1LID3 in toluene using n-octanol as the nucleophile and different activated acyl donors. The specific activity of wt CRL1 was higher than that of CRL1LID3 with most tested acyl donors. We observed that the rates of the two enzymes were affected in the same way by the structure and chain length of the acyl donor used. The highest alcoholysis rate was obtained with activated esters carrying a butyryl moiety. Interestingly, activity on TFE octanoate was higher (about five- and threefold, respectively, for wt CRL1 and CRL1LID3) than on TFE hexanoate. This observation is in good agreement with the low activity on C6 substrates obtained in aqueous medium reactions. These results suggest that in organic solvents, replacement of the lid did affect the specific activity of the enzyme but not its specificity towards activated monoesters employed as acyl donors in synthetic reactions.
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Discussion |
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Here we describe the effect of swapping the lid domains among lipase isoforms. This approach results in the production of chimeric proteins differing in only a few of 534 residues, all of them clustered in the same structural and functional region. However, the lid is the most variable region in sequences which are otherwise very tightly related (Lotti et al. 1994) and is therefore an interesting target for investigating the reported catalytic differences of CRL isoenzymes. Information provided by X-ray analysis supports this choice, indicating that the lid is an important functional structure (Cygler and Schrag 1999). Not only does the lid’s conformation modulate accessibility of the catalytic machinery to substrates, but the lid also contributes to the substrate-binding surface and is likely to be directly involved in substrate recognition. In a number of lipases, but not in CRL, movement of the lid to the open conformation generates the correct positioning of the oxyanion hole, a constellation of amino acids responsible for the stabilization of the reaction tetrahedral intermediate.
Surprisingly, whereas CRL1LID3 was secreted by P. pastoris at levels comparable to those of the wt, the CRL1LID4 mutant was almost fully retained within the cells. This result was unexpected, because this fusion protein was originated by the swapping of a domain from a natural enzyme (CRL4) that can be correctly expressed, folded, and transported by P. pastoris cells (Tang et al. 2001). However, we also observed in other cases that a single mutation in CRL may result in defects of its transport (S. Brocca et al. 2000, unpubl.). A recent series of papers by Sagt and colleagues put emphasis on the sensitivity of yeast expression systems to changes in the sequence of heterologous proteins. Those authors showed that introduction of hydrophobic patches in a cutinase enzyme resulted in retention of the protein in the endoplasmic reticulum (ER), association with chaperones, and impaired secretion (Sagt et al. 1998) and evoked the unfolded protein response (UPR) with consequent proliferation of ER membranes, oxidative stress, and proteasomal degradation of the modified recombinant enzyme (Sagt et al. 2002). It might be hypothesized that the observed block in protein secretion depends on the increased hydrophobicity introduced in the CRL1-based protein by the LID4 peptide. As recombinant CRL4 can be correctly transported (Tang et al. 2001), one should assume that the lid features are counterbalanced by other as yet unknown properties of the wt CRL4 protein, that make it compatible with the P. pastoris secretion machinery.
We did not notice a specific shift in the chain-length specificity as a consequence of the lid swapping. Accordingly, a recent site-directed mutagenesis study unambiguously demonstrated the role of specific residues located in the substrate-binding tunnel of CRL1 (Schmitt et al. 2002). Subtle changes—if any—related to the substrate–lid interaction might be more difficult to assess.
On the other hand, replacement of the loop building the lid with the LID3 peptide was sufficient to confer to CRL1, a protein completely inactive towards cholesterol esters, activity in the hydrolysis of cholesteryl linoleate. This observation is of interest, as it confronts the issue of how CRL1 and CRL3, which share 89% sequence identity and are very similar in their structure, can display completely different behaviors in the hydrolysis of cholesteryl esters. Complexes of CRL1 with substrate-like inhibitors and of linoleate-bound CRL3 disclosed molecular details of substrate–enzyme interactions (Grochulski et al. 1994; Ghosh et al. 1995). CRL1 and CRL3 differ in 55 amino acids, 23 of them located near the active site and at the active-site gorge region, to which the lid contributes. The structural analysis reported by Ghosh and colleagues (1995), besides focussing on substitutions within the substrate-binding pocket, suggested a specific role of the lid residues Phe69, Gly74, Thr76, and Gln88 (Tyr69, Pro74, Ala76, and Glu88 in CRL1) in enhancing hydrophobicity at the entrance of the substrate-binding pocket. This effect should favor binding of very hydrophobic substrates as cholesteryl esters. We have shown that the LID3 sequence alone was sufficient to obtain activity toward cholesteryl linoleate. This result therefore identifies the lid as one of the major structural and functional determinants of the cholesterol esterase activity typical of CRL3. However, the information obtained does not allow us to unambiguously conclude whether specificity is conferred by specific amino-acid side chains, as suggested by Ghosh et al. (1995). Lid-dependent conformational effects as well as the contribution of other protein regions cannot be excluded. We addressed, by site-directed mutagenesis, position 344 within the substrate-binding pocket (Phe in CRL1), because it was suggested as a possible critical residue (Ghosh et al. 1995). However, the mutant did not display any activity on cholesteryl linoleate (data not shown).
Lid replacement was found to produce a striking and substrate-independent change in the enzyme response to detergents added to the reaction mixture in reactions of hydrolysis of p-nitrophenolesters carried out under different experimental conditions. Because three nonionic detergents were used, a differential action on proteins differing from each other in only a 26-residue stretch was hardly to be expected. A clue to this observation is provided by a recent study by Gonzalez-Navarro and colleagues (2001), who addressed the issue of how the conformational state of CRL contributes to substrate preference. In that work, different enzyme conformations present in aqueous solution in the presence of detergents/interfaces were trapped by freeze-drying and used for reactions in a water-restricted environment. It was shown that different conformational states are endowed with different activity on substrates of growing chain length. These results were interpreted as the lipase existing in solution as a continuum of conformations concerning the position of the lid, whose equilibrium can be shifted by the environment (i.e., the detergent concentration). Accordingly, different opening of the lid would make the enzyme accessible to substrates of different sizes. In this conceptual frame, the effects of detergents on enzymes that differ only in their lid structures might be interpreted in terms of specific lid/detergent interactions as well as of a modulation of the conformational flexibility introduced by lid swapping. LID3 differs from the CRL1 sequence in six residues, of which tyrosine 69 forms a hydrogen bond with the sugar linked to Asn 351, which has been shown to stabilize the lid in its open conformation (Grochulski et al. 1993; Brocca et al. 2000). The substitution of Tyr with Phe removes the hydroxyl function and exposes a hydrophobic side chain to the solvent. Moreover, replacement of Pro 74 by Gly might enhance the lid conformational flexibility. The LID4 sequence differs markedly from the wt, with 16 residues substituted. Besides the significant increase in the hydrophobicity of this sequence, replacement of a few residues appears worthy of discussion. In LID4, Glu66, which was recognized by Grochulski et al. (1994) as one of the hinge points for the swinging of the lid, is replaced by leucine. Trp and Asp also replace, respectively, Tyr 69 and Glu70, which have been shown to stabilize the open lid by interactions with sugars. In particular, the lack of the Tyr hydroxyl function and the presence of a bigger side chain might be of some hindrance for the closing of the lid. This might contribute to making the open conformation more populated. In both mutants, the two substitutions Ser91Leu and Ser93Gln/Asn locate larger side chains at the two sides of Pro92, whose cis-trans isomerization determines the opening of the lid (Grochulski et al. 1993).
In conclusion, the role of the lid in affecting CRL activity appears to be very complex and might involve both specific interactions with the substrate molecules and the equilibrium between the active and inactive enzyme conformations.
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Materials and methods |
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Strains, plasmids, and growth conditions
Escherichia coli strain DH5 (Promega) was used as the host for plasmid amplification, and Pichia pastoris X-33 (Cregg et al. 1993; Invitrogen) was used for the expression of recombinant lipases. The plasmids used were pPICB (Invitrogen) for expression in P. pastoris and pGEM-7Zf(+) for intermediate cloning steps (Promega). E. coli was grown at 37°C in low-salt Luria-Bertani (LB) medium (DIFCO) containing either 50 µg/mL ampicillin for selection of clones containing pGEM-7Z or 25 µg/mL zeocin for selection of clones carrying pPICZB. P. pastoris was grown in shaking flasks at 30°C in buffered YEPS medium containing 1% LB yeast extract, 2% peptone, and 1% sorbitol. Selection of transformants was on the appropriate medium containing 25 µg/mL zeocin.
DNA synthesis and generation of chimeric lipases
Plasmid DNA was purified using either the alkaline lysis procedure for mini-preparations (Birnboim and Doly 1979) or kits for mini- or maxi-preparations (QIAGEN). Other standard DNA manipulations were carried out according to Sambrook et al. (1989).
The parent plasmid used for the generation of chimeric lipases was pPICslip1 (Brocca et al. 1998). A 541-bp sequence containing the lid region and delimited by two SmaI restriction sites was replaced with the corresponding sequence of the CRL3 or CRL4 isoenzymes obtained as described in Results. PCR was performed in an automated DNA Thermal Cycler (Perkin Elmer) using 50 pmoles of each primer and 2.5 Units of Pfu TurboTM Polymerase (Stratagene) in a total volume of 100 µL. Samples were subjected to 25 cycles of 1-min denaturation at 94°C, 1-min annealing at 77°C, and 1-min extension at 72°C, followed by a final step of extension of 10 min at 72°C. After validation by sequencing (M-Medical), synthetic sequences were used to replace the corresponding SmaI-SmaI fragment in pPICslip1.
Expression of recombinant lipases
Pichia pastoris X-33 cells were transformed by the Bicine method (Klebe et al. 1983) with 10 µg of SacI-linearized plasmid DNA carrying wt or mutated lipase-encoding genes. Freshly transformed cells were plated onto solid YEPS medium containing 25 µg/mL zeocin. Positive transformants were checked for lipase activity by transferring colonies onto YEPS plates containing 1% tributyrin. After overnight incubation at 30°C, lipase activity was detected by the formation of a clear halo on the opaque tributyrin emulsion. Recombinant cultures were grown in flasks (shaking rate 150 rpm) at 30°C in buffered YEPS medium at pH 6.0 for 5 d, reaching an OD600 of 60–80. The cultures were maintained at constant pH by adding 1 M phosphate buffer pH 6.0 and fed by the addition of 1% v/v methanol twice a day. Wt and chimeric lipases were recovered from the culture supernatants by centrifugation at 4000 rpm. Concentration was carried out by tangential flow filtration against 4–5 vol of 5 mM Tris-HCl pH 7.5 with a Minitan system (Millipore) using filter plates at 60 kD exclusion.
The wild-type and mutant enzymes were identified by electrophoresis on 10% SDS-polyacrylamide gel (Laemmli 1970), with the wild-type LIP1 as a molecular mass reference. The purity degree of the different enzymes was assessed by Coomassie Blue staining of SDS-polyacrylamide gels after electrophoresis. Culture supernatant samples and rough cellular extracts were subjected to Western blotting analysis with a polyclonal antibody raised against G. candidum lipase (GCL). Lipase was quantified by densitometry analysis.
Activity assays
Activity on p-nitrophenyl-esters was determined by following the increase in absorbance at 410 nm due to the release of p-nitrophenol, with a Jasco V-530 UV/Vis spectrophotometer. In 1 mL final volume, 100 µL of 10 mM substrates dissolved in isopropanol were mixed with 100 mM Tris-HCl pH 7.5 and detergents as specified. Nonionic detergents used were Triton X-100 (polyoxyethylene mono p-tert-octylphenylether, CMC 0.015%), Tween 20 (polyoxyethylene sorbitan monolaurate, CMC 0.007%), and Tween 80 (polyoxyethylene sorbitan monooleate, CMC 0.001%).
Activity on triglycerides was determined by titrating released fatty acids with 0.01 N sodium hydroxide using a STAT TITRINO (Metrohm). Emulsions of 20 mM triacylglycerols with 2% arabic gum were used as a substrate in 20 mM potassium phosphate buffer, pH 7.5, in a volume of 20 mL.
Cholesterol esterase activity was determined as follows: 5.3 mg of solid cholesteryl linoleate was dissolved in 0.2 mL polyoxyethylene 9 lauryl ether (Sigma) and mixed under heating with 0.8 mL 0.9 % (w/w) sodium chloride in 50 mM potassium phosphate buffer, pH 7.2 (substrate concentration 8.2 mM). Hydrolysis of cholesteryl linoleate was carried out at 25°C, adding a proper amount of lipase to 1 mL of substrate solution, and the reaction progress was monitored by measuring the release of cholesterol by GLC (HP-1 Crosslinked Methyl Silicone Gum, 25 m, 0.32 mm ID, Hewlett-Packard) under the following operating conditions: oven temperature from 270°C (initial time 5 min) to final temperature 278°C with a heating rate of 0.5°C/min, H2 as the carrier gas. Control CRL3 was the commercial cholesterol esterase, from Roche (n.393916).
Activity in toluene was determined by measuring the initial rate of alcoholysis by n-octanol of vinyl esters or trifluorethylesters or triglycerides. The lipase samples were prepared by co-lyophilizing 0.03 mg of lipase protein with 5 mg PEG (Secundo et al. 1999). Before reaction, solvent, reagents, and enzyme samples were brought separately to the water activity value of 0.11 by equilibration (for at least 48 h, at 25°C) in sealed vessels, with the vapor phase of a saturated LiCl solution. In a typical experiment, a sample of lipase was added to 1 mL toluene containing 19 mM n-octanol and 39 mM activated ester or 20 mM triglycerides, and the reaction mixture was shaken at 150 rpm, at 25°C. The reaction progress was monitored by GLC as described above, but with an oven temperature from 35°C (initial time 10 min) to final temperature 300°C (depending on the chain length of the obtained ester) with a heating rate of 15°C/min.
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The publication costs of this article were defrayed in part by payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 USC section 1734 solely to indicate this fact.
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References |
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Brocca, S., Schmidt-Dannert, C., Lotti, M., Alberghina, L., and Schmid, R.D. 1998. Design, total synthesis, and functional overexpression of the Candida rugosa lip1 gene coding for a major industrial lipase. Protein Sci. 7: 1415–1422.[Abstract]
Brocca, S., Persson, M., Wehtje, E., Adlercreutz, P., Alberghina, L., and Lotti, M. 2000. Mutants provide evidence of the importance of glycosydic chains in the activation of lipase 1 from Candida rugosa. Protein Sci. 9: 985–990.[Abstract]
Carrea, G. and Riva, S. 2000. Properties and synthetic applications of enzymes in organic solvents. Angew. Chem. Int. Ed. Engl. 39: 2226–2254.[CrossRef][Medline]
Carriére, F., Withers-Martinez, C., van Tilbeurgh, H., Roussel, A., Cambillau, C., and Verger, R. 1998. Structural basis for the substrate selectivity of pancreatic lipases and some related proteins. Biochim. Biophys. Acta 1376: 417–432.[Medline]
Cregg, J.M., Vedvick, T.S., and Raschke, W.C. 1993. Recent advances in the expression of foreign genes in Pichia pastoris. Biotechnology 11: 905–910.[CrossRef][Medline]
Cygler, M. and Schrag, J.D. 1999. Structure and conformational flexibility of Candida rugosa lipase. Biochim. Biophys. Acta 1441: 205–214.[Medline]
Dugi, K.A., Dichek, H.L., Talley, G.D., Brewer Jr., H.B., and Santamarina-Fojo, S. 1992. Human lipoprotein lipase: The loop covering the catalytic site is essential for interaction with lipid substrates. J. Biol. Chem. 267: 25086–25091.
Dugi, K.A., Dichek, H.L., and Santamarina-Fojo, S. 1995. Human hepatic and lipoprotein lipase: The loop covering the catalytic site mediates substrate specificity. J. Biol. Chem. 270: 25396–25401.
Ghosh, D., Wawrzak, Z., Pletnev, V.Z., Li, N., Kaiser, R., Pangborn, W., Jörnvall, H., Erman, M., and Duax, W.L. 1995. Structure of uncomplexed and linoleate-bound Candida cylindracea cholesteol esterase. Structure 3: 279–288.[Medline]
Gonzalez-Navarro, H., Bano, M.C., and Abad, C. 2001. The close/open model for lipase activation. Addressing intermediate active forms of fungal enzymes by trapping of conformers in water-restricted environments. Biochemistry 40: 3174–3183.[CrossRef][Medline]
Grochulski, P., Li, Y., Schrag, J.D., Douthillier, F., Smith, P., Harrison, D., Rubin, B., and Cygler, M. 1993. Insights into interfacial activation form an open structure of Candida rugosa lipase. J. Biol. Chem. 268: 12843–12847.
Grochulski, P., Li, Y., Schrag, J.D., and Cygler, M. 1994. Two conformational states of Candida rugosa lipase. Protein Sci. 3: 82–91.
Helistö, P. and Korpela, T. 1998. Effects of detergents on activity of microbial lipases as measured by the nitrophenyl alkanoate esters method. Enzyme Microb. Technol. 22: 113–117.[CrossRef]
Holmquist, M., Clausen, I.G., Patkar, S., Svendsen, A., and Hult, K. 1995a. Probing a functional role of Glu87 and Trp89 in the lid of Humicola lanuginosa lipase through transesterification reactions in organic solvent. J. Protein Chem. 14: 217–224.[CrossRef][Medline]
Holmquist, M., Martinelle, M., Berglund, P., Clausen, I.G., Patkar, S., Svendsen, A., and Hult, K. 1995b. Lipases from Rhizomucor miehei and Humicola lanuginosa: modification of the lid covering the active site alters enantioselectivity. J. Protein Chem. 12: 749–757.
Klebe, R.J., Harriss, J.V., Sharp, Z.D., and Douglas, M.G. 1983. A general method for polyethylene-glycol induced genetic transformation of bacteria and yeast. Gene 25: 333–341.[CrossRef][Medline]
Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of the bacteriophage T4. Nature 227: 680–685.[CrossRef][Medline]
Lee, G.C., Tang, S.J., Sun, K.H., and Shaw, J.F. 1999. Analysis of the gene family encoding lipases in Candida rugosa by competitive reverse transcription-PCR. Appl. Environ. Microbiol. 65: 3888–3895.
Lee, G.C., Lee, L.C., Sava, V., and Shaw, J.F. 2002. Multiple mutagenesis of non-universal serine codons of the Candida rugosa LIP2 gene and biochemical characterization of purified recombinant LIP2 lipase overexpressed in Pichia pastoris. Biochem. J. 366: 603–611.[CrossRef][Medline]
Longhi, S., Lotti, M., Fusetti, F., Pizzi, E., Tramontano, A. and Alberghina, L. 1992. Homology-derived three-dimensional structure prediction of Candida cylindracea lipase. Biochim. Biophys. Acta 1165: 129–133.[Medline]
Lopez, C., Guerra, N.P., and Rua, M.L. 2000. Purification and characterization of two isoforms form Candida rugosa lipase B. Biotech. Lett. 22: 1291–1294.[CrossRef]
Lotti, M., Grandori, R., Fusetti, F., Longhi, S., Brocca, S., Tramontano, A., and Alberghina, L. 1993. Cloning and analysis of Candida cylindracea lipase sequences. Gene 124: 45–55.[CrossRef][Medline]
Lotti, M., Tramontano, A., Longhi, S., Fusetti, F., Brocca, S., Pizzi, E., and Alberghina, L. 1994. Variabilità within the Candida rugosa lipases family. Protein Eng. 7: 531–534.
Lotti, M., Monticelli, S., Montesinos, J.L., Brocca, S., Valero, F., and Lafuente, J. 1998. Physiological control on the expression and secretion of Candida rugosa lipase. Chem. Phys. Lipids 93: 143–148.[CrossRef][Medline]
Martinelle, M., Holmquist, M., Clausen, I.G., Patkar, S., Svendsen, A., and Hult, K. 1996. The role of Glu87 and Trp89 in the lid of Humicola lanuginosa lipase. Protein Eng. 9: 519–524.
Rua, M.L., Diaz-Maurino, T., Fernandez, V.M., Otero, C., and Ballesteros, A. 1993. Purification and characterization of two distinct lipases from Candida cylindracea. Biochim. Biophys. Acta 1156: 181–189.[Medline]
Rubin, B. and Dennis, E.A., eds. 1997. Lipases. Methods in enzymology, Vol. 284. Academic Press, New York.
Sagt, C.M.J., Mueller, W.H., Boonstra, J., Verkleij, A.J., and Verrips, C.T. 1998. Impaired secretion of a hydrophobic cutinase by Saccharomyces cerevisiae cells correlates with an increased association with immunoglobulin heavy-chain binding protein (BiP). Appl. Environ. Microbiol. 64: 316–324.
Sagt, C.M.J., Mueller, W.H., van der Heide, L., Boonstra, J., Verkleij, A.J., and Verrips, C.T. 2002. Impaired cutinase secretion in Saccharomyces cerevisiae induces irregular endoplasmic reticulum (ER) membrane proliferation, oxidative stress, and ER-associated degradation. Appl. Environ. Microbiol. 68: 2155–2160.
Sambrook, K.J., Fritsch, E.F., and Maniatis T. 1989. Molecular cloning: A laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
Schmitt, J., Brocca, S., Schmid, R.D., and Pleiss, J. 2002. Blocking the tunnel: Engineering of Candida rugosa lipase mutants with short chain length specificity. Protein Eng. 15: 595–601.
Secundo, F., Spadaro, S., Carrea, G., and Overbeeke, P.L.A. 1999. Optimization of Pseudomonas cepacia lipase preparations for catalysis in organic solvents. Biotechnol. Bioeng. 62: 554–561.[CrossRef][Medline]
Tang, S.J., Shaw, J.F., Sun, K.H., Sung, G.H., Chang, T.Y., Lin, C.K., Lo, Y.C., and Lee, G.C. 2001. Recombinant expression and characterization of the Candida rugosa lip4 lipase in Pichia pastoris: Comparison of glycosylation, activity and stability. Arch. Biochem. Biophys. 387: 93–98.[CrossRef][Medline]
Verger, R. 1997. "Interfacial activation" of lipases: Facts and artifacts. Trends Biotechnol. 15: 32–38.[CrossRef]
Yang, Y. and Lowe, M.E. 2000. The open lid mediates pancreatic lipase function. J. Lipid Res. 41: 48–57.
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