General Trend of Lipase to Self-Assemble Giving Bimolecular

January/February 2003

Published by the American Chemical Society

Volume 4, Number 1

© Copyright 2003 by the American Chemical Society

Communications General Trend of Lipase to Self-Assemble Giving Bimolecular Aggregates Greatly Modifies the Enzyme Functionality Jose M. Palomo, Manuel Fuentes, Gloria Ferna´ ndez-Lorente, Cesar Mateo, Jose M. Guisan,* and Roberto Ferna´ ndez-Lafuente* Department of Biocatalysis. Institute of Catalysis, CSIC, Campus Universidad Autonoma, Cantoblanco, 28049 Madrid, Spain Received November 15, 2002

Three microbial lipases (those from Candida rugosa, Humicola lanuginosa, and Mucor miehei) have been found to exhibit a tendency to form bimolecular aggregates in solution even at very low enzyme concentrations (44 µg/mL) in the absence of a detergent, as detected by gel filtration. The monomolecular form of the enzymes was found as unique only at low enzyme concentration and in the presence of detergents. However, in the case of the lipase B from Candida antarctica, no bimolecular form could be identified even at enzyme concentrations as high as 1.2 mg/mL in the absence of detergent. It has been stated that bimolecular and monomolecular structures display very different functional properties: (i) the enzyme specific activity decreased when the lipase concentration increased; (ii) the bimolecular form was much more stable than the monomeric one yielding a higher optimal T (increasing between 5 and 10 °C) and higher stability in inactivation experiments (the dimer half-life became several orders of magnitude higher than that of the monomer); (iii) the enantioselectivity depended on the enzyme concentration even after immobilization. For example, with use of the lipase from H. lanuginosa, the enantiomeric excess of the remaining ester in the hydrolysis of fully soluble ethyl ester of (R,S)-2-hydroxy-4-phenylbutanoic acid varied from 4 to 57 when the concentrated or diluted enzyme immobilized on PEI support, respectively, was used. It seems that the bimolecular structure of lipases might be formed by two open lipase molecules (interfacially activating each other) in very close contact and hence with a very altered active center. Introduction Lipases are relevant enzymes from both a physiological and a biotechnological point of view. In addition to their natural function (hydrolysis of triglycerides), lipases are also able to recognize very different substrates catalyzing regioand enantioselective hydrolysis or synthesis of many esters.1-6 Several authors have reported the identification of unspecific aggregated forms of various lipases, most of them * To whom correspondence should be addressed. Mailing address: Departamento de Biocatalisis, Instituto de Cata´lisis, CSIC, Campus Universidad Auto´noma, 28049 Madrid, Spain. Fax: 34 91 585 47 60. Tel: 34 91 585 48 09. E-mail addresses: [email protected]; [email protected].

being due to the use of drastic experimental conditions.7-18 However, dimers of several lipases have been identified by X-ray studies, showing that lipases may crystallize as dimers in their open conformation, showing the hydrophobic areas surrounding the active center in close relation with those areas of other lipase.19-21 This could be based on the very complex mechanism of action of lipases (“interfacial activation”).22-31 This interfacial activation may be promoted by different hydrophobic interfaces: drops of oil (its natural substrate),32 hydrophobic surfaces of supports,33-40 gas bubbles,41 hydrophobic proteins,42 lipopolysaccharides,43 etc. We hypothesize that it is possible that two open lipase molecules may also interact with each other via the large

10.1021/bm025729+ CCC: $25.00 © 2003 American Chemical Society Published on Web 01/13/2003

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Scheme 1. Likely Mechanism of Aggregation of Lipases

hydrophobic surface surrounding the active center of lipases. In this paper, we propose that lipases may have a specific trend to self-assemble even in diluted solutions giving lipase structures with specific properties that may promote certain problems in the biochemical characterization of lipases (Scheme 1) because this hypothetic “dimer” might present different stability, activity, or enantioselectivity (having an open but partially blocked active center of the lipase).44-49 In this paper, we have studied this possibility using four different lipases chosen among the most used ones: lipases from Candida antarctica B, a lipase lacking a lid,50-51 from Mucor miehei,52 from Humicola lanuginosa,53 and from C. rugosa.54 Experimental Section Materials. The lipases from Candida antarctica B (CAL-B, Novozym 525L), Humicola lanuginosa (HLL, Novozym 871), and Mucor miehei (MML, Novozym 388) were supplied by Novo Nordisk (Denmark). The lipase from Candida rugosa (Type VII, CRL, specific activity 875 U/mg solid), racemic mixture of (R,S)-mandelic acid methyl ester, Triton X-100, polyethyleneimine (PEI) with Mr ) 60 kD, p-nitrophenyl propionate (pNPP), and p-nitrophenyl butyrate (pNPB) were purchased from Sigma Chemical Co. Sepabeads resin was generously donated by Resindion Srl (Mitsubishi Chem. Corp., Milan, Italy). PEI-Sepabeads was prepared as previously described.55 Octyl-Sepharose 4BCL was purchased from Pharmacia Biotech (Uppsala, Sweden). The racemic mixture of (R,S)-2-hydroxy-4-phenylbutanoic acid ethyl ester (HPBEt) was a kind gift from VITA INVEST SA. Other reagents and solvents used were also commercially available. Methods. All experiments were carried out in triplicate, and experimental error was never over 5%. Enzymatic Activity Determination Assay. Esterase standard activities assays were determined spectophotometrically using pNPP or pNPB as previously reported (ref 33). Occasionally, 0.1% Triton X-100 was added to the reaction medium. One international unit of pNPP or pNPB activity was defined as the amount of enzyme that is necessary to hydrolyze 1 µmol of pNPP or pNPB per minute (IU) under the conditions described above. Purification of the Different Lipases. To purify the lipases from any other contaminating protein (e.g., esterases),

Communications

the different lipase preparations were adsorbed on octylagarose following the procedure previously described.33 The adsorbed lipases were washed with Triton X-100 (1% for CAL-B, 0.5% for CRL, 0.6% for MML, and 2% for HLL) in 5 mM sodium phosphate buffer at pH 7 and 4 °C to desorb them. The enzyme/detergent mixture was diluted and adsorbed on DEAE-agarose at pH 8.0, and it was washed with distilled water to eliminate all of the detergent. After that, the adsorbed enzyme was released from the support with 200 mM sodium phosphate buffer at pH 8, dialyzed against 5 mM sodium phosphate buffer at pH 7, and concentrated using Centrikon ultrafiltration devices (Millipore, Milford, MA). Gel Filtration of Purified Lipases. Gel filtration analyses were carried out using a 10 mm × 510 mm column packed with agarose beads; the bed volume of the column was 40 mL. The elution buffer used was 100 mM sodium phosphate (occasionally with 0.1% Triton X-100) at pH 7.0, and the experiments were performed at 25 °C and a flow rate of 0.25 mL/min. The column was equilibrated by passing 400 mL of the corresponding buffer. In all cases, 0.2 mL of enzyme at different concentrations were loaded on the column. The samples were fractionated in 1 mL aliquots, and their enzymatic activities were analyzed using pNPP/pNPB assay using very low enzyme concentration and 0.1% Triton X-100 to avoid enzyme aggregation during activity determination. Immobilization of HLL at Different Enzyme Concentrations on PEI-Coated Sepabeads. Immobilization was carried out using a support coated with PEI (60 kDa). For the concentrated enzyme, a volume of 15 mL of 5 mM sodium phosphate buffer containing 0.74 mg/mL of lipase was offered to 3 g of support. For the diluted enzyme, 120 mL of 0.0925 mg/mL of lipase in 0.6% Triton X-100 was offered to 3 g of support. The enzymatic load was 3.7 mg of lipase/mL of support in both cases. The immobilizations were carried out at pH 7 and 25 °C, immobilization yields being near 100% in all cases. Hydrolysis Reactions. (R,S)-2-Hydroxy-4-phenylbutanoic acid ethyl ester (HPBE) (2mM, 20 mL) or (R,S)-mandelic acid methyl ester solutions (10mM, 10 mL) were prepared in 25 mM sodium phosphate buffer at pH 7 and 25 °C. Then, 1 mL of PEI-lipase preparations was added, and the reaction mixture was gently stirred. Reactions were analyzed by reverse-phase and chiral HPLC as previously described.46-47 The enantioselectivity was expressed as the E value calculated from the enantiomeric excess (ee) of the remaining ester according to eq 1.57 E ) ln[(1 - c)(1 - ee)]/ln[(1 - c)(1 + ee)]

(1)

where c is the conversion degree. Temperature-Enzyme Activity Profile of Different Lipase Preparations. The effect of the temperature on the enzyme activity was checked in the hydrolysis of ethyl butyrate (50 mM) at pH 7 in 25 mM sodium phosphate buffer using a pH stat. All assays were performed keeping the same stirring speed, while special care was taken to avoid the formation of air bubbles in the reaction vessel. The buffer was preincubated to reach the desired temperature before

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Figure 1. Determination of the molecular weight of different lipase samples by gel filtration. The elution volume of diluted and concentrated solution of lipases was compared to the elution volume of the following standard proteins: PGA (90 kDa), BSA (67 kDa), lysozyme (14 kDa). Experiments were carried out using 100 mM phosphate as eluent for concentrated solutions (1.2 mg/mL of CAL-B, 1.25 mg/mL of MML, 0.92 mg/mL of CRL, 1.4 mg/mL of HLL) and 100 mM phosphate/0.5% Triton X-100 for diluted solutions (0.15 mg/mL of CAL-B, 0.7 mg/mL of MML, 0.115 mg/mL of CRL, 0.46 mg/mL of HLL). In all cases, 0.2 mL of different enzymes concentrations were loaded on the column.

adding the ethyl butyrate and the enzyme. The reaction was started by the addition of the enzyme. Thermal Inactivations. To check the stability of different lipases, solutions of the enzymes at different concentrations were incubated at pH 7 and 50 °C. Samples were withdrawn periodically, and the activity was measured using pNPP/ pNPB assay, ensuring to have the same lipase concentration in the activity assay. Results Analyses of Self-Assembly of Lipases. Figure 1 shows that purified CRL, MML, and HLL presented different apparent molecular weight in gel filtration experiments at low concentration (e.g., 0.044 mg/mL) in the presence of detergent or when increasing the protein concentration by a factor of 8. Diluted preparations in the presence of detergent yielded a single peak, and the molecular weight approximately matched the one calculated by SDS-PAGE. Concentrated preparations originated two peaks, one corresponding to the monomer (minority) and another that gave an apparent Mr approximately doubling that value. Even using very low lipase concentration, a certain percentage of bimolecular aggregates could be detected, although this percentage increased with the lipase concentration. In addition, by use of higher ionic strength, the percentage of bimolecular structures could be increased. Similar results were achieved using unpurified samples of lipases, suggesting that this aggregation is not derived from the presence of detergent adsorbed on the enzyme. However, we have not been able to detect higher oligomers (formed by three or more lipase molecules) even at the maximum enzyme concentration employed. These results suggest that perhaps lipases present one area that can yield a strong lipase-lipase interaction. In addition, considering that ionic strength increased the percentage of bimolecular form while detergent decreased it, it seems that a hydrophobic interaction could play some role in the generation of this “dimer”.

Figure 2. Influence of the concentration (a) of lipases on their specific activity: (9) CAL-B, ([) HLL, (2) CRL, (b) MML. Experiments were carried out using pure enzyme and pNPP or pNPB as substrate at pH 7 and 25 °C as described in the Experimental Section. Panel b shows the specific activity of CRL at growing concentration of enzyme under different conditions. Activity was determined (2) in standard buffer or (9) in the presence of 0.1% Triton X-100. Experiments were carried out as described in the Experimental Section.

Only the monomer for CAL-B in the whole range of concentrations studied (0.15-1.2 mg/mL) was detected. This might be due to the lack of lid that results in a smaller hydrophobic area available for lipase-lipase interaction.50 This result in diluted solutions contrasts with the fact that CAL-B may be crystallized in a dimeric form,20 but we should consider that the ionic strength and protein concentration was much higher in the crystallization experiment. Effect of the Enzyme Concentration on the Activity of Lipases. The specific activity of CRL, HLL, and MML on fully soluble substrates strongly depended on their concentrations (Figure 2a); when the enzyme concentration increased, the specific activity of the enzyme decreased. This fact was observed even at a much lower concentration than the ones used in the gel filtration experiments. However, the specific activity was independent of the lipase concentration for CAL-B. In the presence of detergent, lipases exhibited an almost constant specific activity in the range of concentrations used in this experiment (Figure 2b). Moreover, several experiments were performed using substrate under saturation concentration (substrate fully soluble) and over saturated substrate solutions (that is, in the presence of “droplets” of insoluble substrate) in which the lipases molecules could suffer interfacial activation. When using fully soluble substrate to measure the activity, the lipase concentration presented a significant effect on the apparent specific activity. The presence of drops of substrate greatly decreased this effect of concentration in enzyme activity (Figure 3).

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Figure 3. Effect of the concentration of lipase on the activity/substrate concentration curve: (2) 12.5 µg/mL; (9) 0.042 µg/mL. The lipase was that from MML. Experiments were performed in 100 mM sodium phosphate at pH 7 and 25 °C.

Figure 5. Influence of the concentration of lipase on the stability of the enzyme. Inactivations were performed at 50 °C and pH 7 as described in the Experimental Section. Panel a shows the inactivation of lipase from H. lanuginosa: ([) 0.74 mg/mL; (2) 0.09 mg/mL. Panel b shows the inactivation of lipase from M. miehei: ([) 0.35 mg/mL; (2) 0.044 mg/mL. Panel c shows the inactivation of lipase from C. antarctica B: ([) 1.2 mg/mL; (2) 0.15 mg/mL.

Figure 4. Effect of temperature on enzyme activity of different lipases. Experiments were carried out using ethyl butyrate as substrate as described in the Experimental Section. Panel a shows the effect on the hydrolytic activity of CRL: (2) 0.92 mg/mL; (9) 0.115 mg/mL. Panel b shows the effect on the hydrolytic activity of MML: (2) 0.35 mg/mL; (9) 0.044 mg/mL. Panel c shows the effect on the hydrolysis activity of CAL-B: (2) 1.2 mg/mL; (9) 0.15 mg/mL.

Effect of Lipase Concentration on Their Temperature/ Activity Profile. When fully soluble substrate was used (to prevent interfacial activation of lipases that could break the “dimer”), the optimal temperatures for concentrated MML

and CRL were higher (64 and 55 °C, respectively) than those when diluted lipase solutions were used (56 and 45 °C, respectively) (Figure 4). However, in the case of CAL-B, the maximum activity was independent of the enzyme concentration; thus, the optimal T was 55 °C using either 0.15 or 1.2 mg/mL. Effect of the Enzyme Concentration on the Thermal Stability of Different Lipases. Figure 5 shows that the enzyme was more stable when its concentration increased in all cases, except for CAL-B. The differences in stability between concentrated and diluted preparation become very significant in some instances. For example, the activity of the diluted preparation of HLL (mainly monomeric and closed form) decreased by 40% in 20 h, while the concentrated preparation (enriched in bimolecular forms) maintained more than 80% of initial activity after 30 h of incubation at 50 °C.

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Table 1. Effect of the Enzyme Concentration during Immobilization on the Enantioselectivity of HLL in the Hydrolysis of Different Racemic Estersa

enzyme preparation

enzyme conc in the immobilization (mg/mL)

preferred enantiomer

eeb

PEI PEI

0.09 0.74

R R

0 16

(1)

(2)

Ec

preferred enantiomer

eeb

Ec

1 1.4

R R

4 57.5

1.077 4

a Immobilized preparations were produced as described in the Experimental Section. Other specifications as described in the Experimental Section. Enantiomeric excess was determined by chiral HPLC analysis as described in the Experimental Section. c Enantioselectivity was calculated as described in the Experimental Section.57

b

Enantioselectivity of the Lipases at Different Concentrations. One of the most interesting applications of lipases is the resolution of racemic mixtures, and the industrial use of the lipase should be in its immobilized form to simplify the reaction performance. Thus, concentrated and diluted solutions of HLL were immobilized on PEI-coated supports (a strong anionic exchanger designed to immobilize proteins ionically)55 and used in the hydrolytic resolution of several chiral compounds (HPBEt and mandelic acid methyl ester). The immobilization of the enzyme on porous supports ensures that the enzyme only can act on fully soluble substrates. We have found that results achieved using the immobilized preparation from concentrated and diluted HLL were very different in both examples (Table 1). The immobilized enzyme under diluted conditions was not enantiospecific at all toward mandelic acid methyl ester, while the enzyme immobilized under concentrated conditions gave moderate enantiomeric excess (15%). However, in the resolution of HPBEt, the enzyme immobilized under diluted conditions gave an enantiomeric excess of only 4%, while the concentrated preparation yielded an enantiomeric excess of around 60% under the same conditions. The implications of this result are evident: even the final properties of immobilized lipase preparations can vary when changing the immobilization conditions (altering the dimer/monomer ratio). It has been previously described that immobilization of lipases on octyl-agarose proceeds via interfacial activation on the support.33 We have prepared interfacially activated HLL immobilized preparations using different enzyme concentrations (from 0.09 to 0.74 mg/mL) and assayed their enantioselectivity against mandelic acid methyl ester. In all cases, the specific activity of the immobilized lipase and its enantioselectivity were identical (around 1). Again, it seems that any phenomenon that implied interfacial adsorption of lipases was able to break the bimolecular form. Discussion This paper shows that, even at moderate enzyme concentrations, lipases in solution exhibited a strong tendency

to self-assemble giving a mixture of bimolecular and monomeric structures that may complicate the characterization of the intrinsic properties of the soluble lipase molecule. In fact, this trend provoked dramatic changes in the properties of the lipases: the enzyme properties seem to depend on this pseudo-quaternary structure of the lipase. Some evidences suggest that the mechanism shown in Scheme 1 is the most likely to explain this self-assembling of lipases into bimolecular aggregates: (i) the changes in catalytic and stability properties; (ii) the fact that bimolecular structures are the only aggregates detected even under the most drastic conditions (high concentration, high ionic strength); (iii) the apparent generality of this trend for most lipases having a lid; (iv) the apparent hydrophobic nature of the interaction; (v) the inexistence of large hydrophobic areas outside the hydrophobic pocket surrounding the active center of lipases; (vi) the competition between lipase self-assembling and interfacial activation. The dramatic structure-function alterations of the soluble lipase with the enzyme concentrations should be considered in any study implying lipases (even immobilized preparations). To homogenize the results among different research groups, it seems convenient to compare the same lipase form, but this seems difficult using soluble enzyme. From our results, it is difficult to have only the monomeric form of soluble microbial lipases. The monomeric lipase can be only achieved in the presence of detergents in most cases. Moreover, to have just the bimolecular structure is also very difficult, and this structure did not represent the actual intrinsic properties of the lipase molecule. Considering that lipases are going to be industrially used mainly in immobilized form, we propose to characterize the different lipases using solid-phase systems. To have the open form of fully dispersed monomeric lipases, we propose the interfacial adsorption of lipases on hydrophobic supports (e.g., on octyl-agarose or octadecyl-Sepabeads).33-43 This simple technique seems to be able to break the bimolecular structure, and it is a very fast and simple method to stabilize,

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hyperactivate, concentrate, purify, and immobilize lipases. In our laboratory, we have been able to prepare immobilized enzyme preparations from very different lipase concentrations achieving final immobilized preparation with exactly the same specific activity, stability, and enantioselectivity. If a lipase preparation close to the soluble lipase is desired, one-point immobilization on poorly activated dextran-coated supports followed by washing with detergent may ensure the production of this monomeric lipase form.46 The properties of this lipase should be very similar to the soluble lipase but without possibilities of protein-protein interactions. Acknowledgment. The authors gratefully recognize the support from the Spanish CICYT with the Projects BIO20000747-C05-02 and BIO2001-2259. Authors thank CAM for a Ph.D. fellowship for Mr. Palomo and a postdoctoral fellowship for Dr. Ferna´ndez-Lorente. We also gratefully recognize an I3P-BPG2001 fellowship from CSIC for Mr. Fuentes. The kind donation of HPBEt from Vita Invest, the lipases supply from Novo Nordisk, and Sepabeads from Resindion are gratefully recognized. We thank Dr. Martinez (Novo Nordisk) for his help and support. The help and suggestions from Mr. Angel Berenguer during the writing of this paper are gratefully recognized. References and Notes (1) Wong, C.-H.; Whitesides, G. M. Enzymes in Synthetic Organic Chemistry; Baldwin, J. E., Magnus, P. D., Eds.; Tetrahedron Organic Chemistry Series; Pergamon: Oxford, 1994; Vol. 12. (2) Bornscheuer, U. T.; Kazlauskas, R. J. Hydrolases in Organic synthesis-Regio and StereoselectiVe Biotransformations; WileyVCH: Weinheim, Germany, 1999. (3) Sharma, R.; Chisti Y.; Banerjee U. C. Biotechnol. AdV. 2001, 19, 627-662. (4) Diaz, M.; Ferrero, M.; Ferna´ndez, S.; Gotor, V. Tetrahedron: Asymmetry 2002, 13, 539-546. (5) Reetz, M. T. Curr. Opin. Chem. Biol. 2002, 6, 145-150. (6) Gao, C.; Whitcombe, M. J.; Vulfson, E. N. Enzyme Microb. Technol. 1999, 25, 264-70. (7) Ru´a, M. L.; Schmidt-Dannert, C.; Wahl, S.; Sprauer, A.; Schmid, R. D. J. Biotechnol. 1997, 56, 89-102. (8) Du¨nhaupt, A.; Lang, S.; Wagner, F. Biotechnol. Lett. 1992, 14, 953958. (9) Jaeger, K. E.; Adrian, F. J.; Meyer, H. E.; Hancock, R. E. W.; Winkler, U. K. Biochim. Biophys. Acta 1992, 1120, 315-321. (10) Lesuise, E.; Schanck, K.; Colson, C. Eur. J. Biochem. 1993, 216, 155-160. (11) Schmidt-Dannert, C.; Sztajer, H.; Sto¨cklein, W.; Mengen, U.; Schimd, R. D. Biochim. Biophys. Acta 1994, 1301, 105-114. (12) Sugihara, A.; Tani, T.; Tominaga Y. J. Biochem. 1991, 109, 211216. (13) Sugihara, A.; Ueshima, M.; Shimada, Y.; Tsunasawa, S.; Tominaga, Y. J. Biochem. 1992, 112, 598-603. (14) Liou, Y.-C.; Marangoni, A.; Yada, R. Y. Food Res. Int. 1998, 31, 243-248. (15) Stuer, W.; Jaeger, K.-E.; Winkler, U. K. J. Bacteriol. 1986, 168, 1070-1074. (16) Osborne, J. C., Jr.; Bengtsson-Olivecrona, G.; Lee, N. S.; Olivecrona, T. Biochemistry 1985, 24, 5606-5611. (17) Berryman, D. E.; Mulero, J. J.; Barry Hughes, L.; Brasaemle, D. L.; Bensadoun, A. Biochim. Biophys. Acta 1998, 1120, 315-321. (18) Graupner, M.; Haalck, L.; Spener, F.; Lindner, H.; Glatter, O.; Paltauf, F.; Hermetter, A. Biophys. J. 1999, 77, 493-504. (19) Ghosh, D.; Wawrzak, Z.; Pletnev, V. Z.; Li, N.; Kaiser, R.; Pangborn, W.; Jornvall, H.; Erman, M.; Duax, W. L. Structure 1995, 3, 279288. (20) Uppenberg, J.; Ohrner, N.; Norin, M.; Hult, K.; Kleywegt, G. J.; Patkar, S.; Waagen, V.; Anthonsen, T.; Jones, T. A. Biochemistry 1995, 34, 16838-16851. (21) Pernas, M. A.; Lo´pez, C.; Ru´a, M. L.; Hermoso, J. FEBS Lett. 2001, 501, 87-91.

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BM025729+