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Novel microbial lipases: catalytic activity in reactions in organic media
Enzyme and Microbial Technology 28 (2001) 145–154
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Novel microbial lipases: catalytic activity in reactions in organic media F. Cardenasa,, M.S. de Castrob, J.M. Sanchez-Monterob, J.V. Sinisterrab, M. Valmasedaa, S.W. Elsona, E. Alvareza,* a
SmithKline Beecham, Centro de Investigacio´n Ba´sica, Santiago Grisolı´a, 4, Parque Tecnolo´gico de Madrid, 28760 Tres Cantos, Madrid, Spain b Departamento de Quı´mica Orga´nica y Farmace´utica, Facultad de Farmacia, Universidad Complutense, 28040 Madrid, Spain Received 30 May 2000; accepted 17 July 2000
Abstract 2,000 microbial strains were isolated from soil samples and tested to determine their lipolytic activity by employing screening techniques on solid and in liquid media. Culture broths were initially tested with 1,2-O-dilauryl-rac-glycero-3-glutaric acid-resorufinyl ester, a chromogenic substrate specific for lipases. Fourteen lipase-producing microorganisms were selected and their taxonomic identification was carried out. Hydrolysis of tributyrin or olive oil and the esterification of oleic acid with heptanol were selected to preliminary evaluate the catalytic activity of these lipases. All the selected lipases catalysed this esterification reaction with good yields. Resolution of (R,S)-2-(4isobutylphenyl) propionic acid, (R,S)-1-phenylethanol, (R,S) 1-phenylethylamine and of (R) or (S) glycidol were performed to evaluate the stereoselectivity of these novel enzymes as biocatalysts in reactions in organic media. Lipases from the fungi Fusarium oxysporum and Ovadendron sulphureo-ochraceum gave the best yields and enantioselectivities in the resolution of racemic ibuprofen and 1-phenylethanol. Several lipases displayed a high stereoselectivity in the resolution of chiral amines by an alcoxycarbonylation reaction. © 2001 Elsevier Science Inc. All rights reserved. Keywords: Screening; Microbial lipases; Lipase-catalysed reactions; Resolution of racemic acids; Alcohols and amines
1. Introduction Lipases (triacylglycerol hydrolases EC 3.1.1.3) are enzymes that catalyse the hydrolysis of triglycerides at the oil-water interface. They are produced by animals, plants and microorganisms [1–3]. Microbial lipases have a great potential for commercial applications due to their stability, selectivity and broad substrate specificity [4]. To date, a large number of lipases from filamentous fungi have been extensively studied, both from the biochemical and from the genetic point of view. The most productive species belong to the genera Geotrichum, Penicillium, Aspergillus and Rhizomucor [5,6]. There are also a certain number of lipases produced by yeasts, most of them belonging to the Candida genus, that have been used for biotechnological purposes [7,8]. Lipases from unicellular bacteria, mainly those produced by various species of the genus Pseudomonas, have also proved to be useful both in organic reactions and in the detergent industry. Many of them have been purified, char-
* Corresponding author. Tel.: ⫹34 1 807 4000. E-mail address: [email protected] (E. Alvarez).
acterised and their encoding genes cloned [9]. By contrast, and despite its high biotechnological interest for the production of secondary metabolites and enzymes, the actinomycetes have not been widely exploited for lipase studies, and there is little information available concerning lipaseproducing actinomycetes [1,10]. Among the high number of lipases described in the literature, only the enzymes belonging to a few species have been demonstrated to have adequate stability and biosynthetic capabilities to allow routine use in organic reactions, and hence their consideration as industrially relevant enzymes [11,12]. The application of different techniques of enzyme immobilisation and of chemical modification of enzymes has introduced considerable improvements in the efficiency of certain processes [13,14]. Similarly, a more detailed study on the nature of the reaction medium, involving parameters such as water activity and log P, and the use of optically active solvents has also made important contributions to both the increase in reaction yields, and the selectivity of the lipases [15–17]. However, despite the above mentioned advances in modification and improvement of enzyme activity, there still remain many important problems to be solved in the application of lipases to indus-
0141-0229/01/$ – see front matter © 2001 Elsevier Science Inc. All rights reserved. PII: S 0 1 4 1 - 0 2 2 9 ( 0 0 ) 0 0 2 7 8 - 7
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trial processes. Therefore, access to a wider range of lipase types would be beneficial [18]. In the present study we describe the selection by screening techniques of fourteen lipaseproducing microorganisms and the preliminary characterisation of the lipases as catalysts in reactions in organic media.
2. Materials and methods 2.1. Chemicals The lipase chromogenic substrate 1,2-O-dilauryl-racglycero-3-glutaric acid resorufinyl ester was purchased from Boehringer Mannheim Biochemicals. (R,S)-2-(4-isobutylphenyl)propionic acid (ibuprofen) was obtained from Boots Pharmaceuticals. Yeast extract and Bacto-peptone were purchased from Difco. The other components of the culture media and the chemical reagents were obtained from Merck and from Sigma in the highest purity available. All the reaction products were chemically synthesised and characterised by IR, 1H-NMR, 13C-NMR and elemental analysis. The data obtained agreed with those reported in the literature. These compounds were used as the reference in HPLC, GC and 1H-NMR-Eu analysis. 2.2. Isolation of microorganisms and screening of lipolytic activity All the microorganisms used in this study were isolated from soil samples collected from around the world. The isolation process was performed by serial dilution of the samples according to standard techniques [19]. Taxonomic identification of unicellular bacteria, actinomycetes and yeasts was performed by the Spanish Type Culture Collection (C.E.C.T., Valencia, Spain). The filamentous fungi were identified in-house by using mature cultures on suitable media (normally standard potato dextrose agar and/or oat meal agar) in order to ensure a good development of taxonomically relevant features, and following the identification keys provided by von Arx [20], Domsch et al. [21] and the more specific methodology of Singler and Carmichael [22] in the case of Ovadendron sulphureo-ochraceum. Preliminary screening of lipolytic filamentous fungi, yeasts and bacteria was carried out on BYPO [23] agar plates supplemented with a 10% (v/v) olive oil or tributyrin emulsion prepared in 10% (w/v) gum Arabic solution. Culture plates were incubated at 28°C and periodically examined for four days. Actinomycetes were screened on plates containing NE [24] medium with the same supplement, incubated at 28°C and examined for seven days. Colonies showing clear zones around them were picked out and screened for lipase production in liquid medium. Two different liquid media (H1 and H2) were employed for filamentous fungi. Medium H1 contained 0.5% yeast extract, 3% Bacto-peptone, 0.1% KH2PO4, 0.1% NaNO3, 0.05% MgSO4 and 1% (v/v) olive oil. Medium H2 contained 7.5%
soybean flour, 2% glucose, 0.1% KH2PO4, 0.1% NaNO3, 0.05% MgSO4 and 1% (v/v) olive oil. The final pH was 7.0 for both media. The liquid medium for actinomycetes (M9 medium) contained 1% soybean flour, 1.8% oatmeal, 0.5% yeast extract, 0.5% glucose and 1% (v/v) olive oil. The final pH was adjusted to 7.2. Yeasts were cultured in liquid YED medium and unicellular bacteria were cultured in liquid BYPO medium. All the cultures were grown in 250 ml flasks containing 50 ml of medium and in 2 litre flasks containing 250 ml of medium. Unicellular bacteria, filamentous fungi and yeasts were incubated at 28°C for 3 days and actinomycetes were cultured in the same conditions for 7 days. Flasks were incubated on a reciprocal shaker at 250 rpm. After this time, culture broths were collected, centrifuged and assayed for lipase activity. Positive supernatants were dialysed overnight against 25 mM Tris-HCl buffer, pH 7, and further lyophilised and stored at 4°C. 2.3. Assay of lipase activity Lipase activity present in culture broths was initially monitored using the chromogenic substrate 1,2-O-dilaurylrac-glycero-3-glutaric acid resorufinyl ester. The reaction was carried out at 37°C for 15 min in 96-well microtiter plates using the conditions described by the supplier. The appearance of a red colour due to free resorufinol indicated the presence of lipase activity in the culture broth. Supernatants that hydrolysed the chromogenic substrate were selected in order to quantitatively determine their lipase activity by using olive oil and tributyrin as substrates. This titrimetric assay was carried out essentially as described by Tietz and Fiereck [25]. 1 unit of lipase activity (U) is defined as the amount that releases 1 mol of fatty acid from triglyceride per minute of reaction at 37°C. 2.4. Heptyl oleate synthesis Heptyl oleate synthesis was carried out to verify the catalytic activity of the lipases in the synthesis of esters in organic medium. The reaction mixture was composed of 20 mM oleic acid, 20 mM 1-heptanol, 3 ml of isooctane, 0.3 ml of water and lyophilised crude enzyme. The amount of enzyme used per reaction was that corresponding to 50 ml of the centrifuged culture supernatant. The reaction was carried out at 30°C with vigorous magnetic stirring. Samples of 50 l were withdrawn at different reaction times and diluted in 500 l of isooctane. The amount of ester formed during the reaction was quantified by gas chromatography as described below. 2.5. Resolution of (R,S)-2(4-isobutylphenyl)propionic acid (Ibuprofen) The resolution of (R,S)-ibuprofen was performed by stereoselective esterification using 1-propanol as acyl acceptor. The reaction mixture was composed of 66 mM (R,S)-2-(4-
F. Cardenas et al. / Enzyme and Microbial Technology 28 (2001) 145–154
isobutylphenyl)propionic acid (ibuprofen), 66 mM 1-propanol, 2 ml of isooctane, 0.2 ml of water and lyophilised enzyme. The amount of enzyme used per reaction was that corresponding to 50 ml of the centrifuged culture supernatant. The reaction was carried out at 30°C with vigorous magnetic stirring. Samples of 30 l were withdrawn at different times and diluted in 500 l of isooctane. The amount of ester formed during the reaction and the enantiomeric excess of the remaining acid were determined by gas chromatography and by HPLC, respectively, as described below. 2.6. Resolution of (R,S)-1-phenylethanol The resolution of (R,S)-1-phenylethanol was performed by stereoselective transesterification using vinyl acetate as acyl donor. The reaction mixture was composed of 1 M (R,S)-1-phenylethanol, 1 M vinyl acetate, 5 ml of isooctane and lyophilised enzyme. The amount of enzyme used per reaction was that corresponding to 50 ml of the centrifuged culture supernatant. The reaction was carried out at 30°C with vigorous magnetic stirring. Samples of 100 l were withdrawn at different times and diluted in 900 l of isooctane. The amount of ester formed during the reaction and the enantiomeric excess of the remaining alcohol were determined by HPLC as described below. 2.7. Acylation of (R) and (S) glycidol The stereoselective acylation of (R) and (S) glycidol was carried out using vinyl acetate as acyl donor. The reaction mixture was composed of 100 mM (R) or (S) glycidol, 100 mM vinyl acetate, 4 ml di-isopropyl ether and 200 mg of lyophilised enzyme. The enzyme was obtained by precipitation of culture broths with (NH4)2SO4, to 90% saturation, dialysed and liophylised. Samples of 100 l were withdrawn and diluted in 900 l of a solution of 4 mM nheptanol (internal standard) in di-isopropyl ether. The amount of ester was determined by gas chromatography as described below. 2.8. Resolution of (R,S) 1-phenylethylamine The resolution of (R,S) 1-phenylethylamine was performed using the lipase-catalysed alcoxycarbonylation of the chiral amine. The reaction mixture was composed of 120 mM butyl-vinyl-carbamate, 120 mM (R,S) 1-phenylethylamine and 200 mg of lyophilised enzyme in 15 ml hexane. The enzyme was obtained by precipitation of culture broths as described above. The reaction products were analysed by HPLC as described below. 2.9. Gas chromatography analysis Gas chromatography analysis of the reaction products was performed using a Shimadzu GC-14 A gas chromato-
147
graph equipped with a FID detector and a SPB-1 column (15 m ⫻ 0.32 mm). For the heptyl oleate analysis the injector and FID temperature was 250°C. The column temperature was kept at 120°C for 5 min, and then programmed to rise 13°C per min to reach 250°C, and maintained at this temperature for 5 min. Nitrogen flux was kept constant at 60 ml/min. For the resolution of (R,S)-ibuprofen the injector temperature was 300°C and the FID temperature was 350°C. The column temperature was kept constant at 180°C and the nitrogen flux was 30 ml/min. For glycidyl acetate synthesis the injector temperature was 200°C and the FID temperature was 250°C. The column temperature was kept constant at 110°C and the nitrogen flux was 13 ml/min. The retention times for glycidyl acetate and heptanol were 2.7 and 4.3 min, respectively. 2.10. HPLC analysis In the resolution of (R,S)-ibuprofen the enantiomeric excess of the remaining acid was determined by HPLC using a Chiracel-OD column. The mobile phase was a mixture of n-hexane:isopropanol:trifluoroacetic acid (100: 1:0.1, v/v/v) at a flow rate of 0.8 ml/min. Detection was performed at 254 nm. The retention times for (R) and (S)-ibuprofen were 19 and 23 min, respectively. The 1-phenylethyl acetate formed during the transesterification reaction and the enantiomeric excess of the remaining alcohol were also determined using a Chiracel-OD column. The mobile phase was a mixture of n-hexane: isopropanol (97:3, v/v) at a flow rate of 0.7 ml/min. In all cases, quantification of the reaction products was performed by measuring their absorbance at 254 nm. The retention times for (R) and (S)1-phenylethanol, and for (R,S)1-phenylethylacetate were 17, 19 and 7 min, respectively. The yield of butyl-(R,S) 1-phenylethyl carbamate was determined by HPLC using a Nucleosil 120 C18 5 m column (25 ⫻ 0.46 cm) from Tracer Analytical. The mobile phase was H2O/MeOH (50:50, v/v). The flux was 0.3 ml/ min. Detection was performed at 254 nm. The retention times for the carbamate and the amine were 29.6 and 27.5 min, respectively. 2.11. Enantiomeric excess of carbamates The enantiomeric excess of butyl-(R,S)-1-phenylethylcarbamate produced was measured using europium (III) tris-[3-(heptafluoropropylhydroximethylen)-(⫹)-camphorate], using the 1H-NMR technique. The carbamate was isolated from the reaction mixture by column chromatography on neutral silica, using hexane-ethyl acetate (7:3, v/v) as eluent. The carbamate/Eu salt ratio used was 1/0.4 (mol/ mol) and the chemical shifts of CH were ␦ ⫽ 6.3 ppm for the (S)-isomer, and ␦ ⫽ 5.8 ppm for the (R) isomer. The enantiomeric excess was determined by comparison of the area of both peaks.
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Table 1 Number and distribution in microbial groups of the microorganisms selected in this study through the screening process Screening step
Soil isolation Solid medium screening Liquid medium screening (chromogenic assay) Liquid medium screening (titrimetric assay)
Number of selected microorganisms
Distribution in microbial groups Bacteria
Actinomycetes
Yeasts
Filamentous fungi
2,000 450 118
200 181 11
840 83 56
100 61 6
860 125 45
14
1
3
1
9
3. Results and discussion 3.1. Selection of lipolytic microorganisms The use of solid media supplemented with emulsified triglycerides is a standard methodology for the selection of lipase-producing microorganisms [26,27]. A total of 2,000 microorganisms isolated from soil samples, including unicellular bacteria (200 strains), actinomycetes (840 strains), yeasts (100 strains) and filamentous fungi (860 strains), were employed in this work (see Table 1). All the microbial strains were screened for lipase activity in solid medium plates containing olive oil or tributyrin as indicator. A total of 450 microorganisms produced a clear halo around them in plates containing tributyrin. Of these, only 92 microorganisms also showed hydrolysis on the olive oil plates, thus indicating a predominant presence of lipases acting on short chain triglycerides. The number of microorganisms that produced detectable levels of lipase in solid media represents 22% of the tested microorganisms, which we consider to be a very high percentage of lipase producing species. It should be noted that each microorganism has been only tested for lipase production in one solid culture medium, which probably would not be appropriate for the lipase induction from many microorganisms. The use of a set of different media for the screening could probably allow to the detection of a higher number of lipase producing microorganisms. The 450 microorganisms selected in the first step were cultured in liquid medium using olive oil as lipase inducer. After centrifugation of culture broths, supernatants were tested with 1,2-O-dilauryl-rac-glycero-3-glutaric acid resorufinyl ester, a chromogenic substrate specific for lipases. The appearance of a red colour due to the release of resorufinol indicated that the supernatant had lipase activity. 118 culture broths were positive in this qualitative assay and were used to further quantify their lipase activity by using tributyrin and olive oil as substrates. During this step, the relative distribution of the selected microorganisms suffered a new dramatic change with respect to the distribution found in the previous step. The lipase activity against the chromogenic substrate displayed by most of the selected bacteria
was very low and only a small percentage were selected for the next selection step. By contrast, a high percentage of actinomycetes were positive in this assay (Table 1). This difference between the bacterial and actinomycetes lipases against the chromogenic substrate could be due to differences in the level of lipase production in liquid media, or to different substrate specificity of the lipases from both groups. Hydrolytic activity of the lipases selected in the previous step was determined by using tributyrin and olive oil as substrates. From this analysis, 14 strains were selected showing the best lipase yields (lipase activity higher than 1 U/ml for one or for both triglycerides) and the best reproducibility in liquid medium cultures. The producing strains were taxonomically identified as follows: nine filamentous fungi (Acremonium murorum (Corda) W. Gams, Monascus mucoroides van Tiegh, Monascus sp., Arthroderma ciferrii Vars & Ajello, Fusarium poae (Pec.) Wollenw, Fusarium solani (Mart.) Sacc., Fusarium oxysporum, Penicillium chrysogenum Thom, Ovadendron sulphureo-ochraceum (van Beyma) Singler & Carmichael, one yeast (Rhodotorula araucariae), one unicellular bacterium (Brevibacterium linens) and three actinomycetes (Streptomyces fradiae, Streptomyces halstedii, Streptomyces sp.). As far as we know, A. murorum, M. mucoroides, A. ciferrii, F. poae, O. sulphureoochraceum, R. araucariae and S. halstedii are reported here for the first time as lipase producers. F. solani has been reported to produce a very active cutinase that also displays triacylglycerol lipase activity [28]. F. oxysporum, P. chrysogenum, B. linens and S. fradiae have also been previously found to have lipolytic activity in solid and in liquid media [1,29,30,31]. In addition, other strains belonging to Monascus [1,6], Fusarium [31], Streptomyces [32] and Rhodotorula [33] genus have been reported to show lipolytic activity. As shown in Table 2, all the selected lipases displayed better hydrolytic activity against tributyrin than against olive oil, a general characteristic previously observed during the screening in solid medium. In general, lipase yields obtained for microorganisms cultured in 250 ml flasks were similar to those obtained in 2 l flasks (data not shown). When stored at 4°C as culture broths, all the selected lipases maintained their activity for at least two weeks. When
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Table 2 Lipase activity (U/ml of supernatant) measured in culture broths from the fourteen microorganisms selected in this study
Table 3 Time course of heptyl oleate synthesis performed by the lipases selected in this study
Producing strain
Producing strain
A. murorum M. mucoroides Monascus sp. A. ciferrii F. poae F. solani F. oxysporum P. chrysogenum O. sulphureo-ochraceum R. araucariae B. linens S. fradiae S. halstedii Streptomyces sp.
Culture medium
Lipase activity (U/ml) Olive oil
Tributyrin
H1 H1 H1 H2 H2 H2 H2 H2 H2 YED BYPO M9 M9 M9
0.72 3.02 2.35 1.43 0.93 0.60 2.83 0.64 4.33 0.75 0.35 0.89 0.70 1.16
5.18 4.31 2.37 1.89 1.47 1.25 5.70 4.68 6.10 3.64 1.83 3.16 1.27 2.10
A. murorum M. mucoroides Monascus sp. A. ciferrii F. poae F. solani F. oxysporum P. chrysogenum O. sulphureo-ochraceum R. araucariae B. linens S. fradiae S. halstedii Streptomyces sp.
Yielda (%) 4h
8h
6 95 34 41 77 79 28 11 71 23 44 44 38 24
29 96 49 77 89 83 52 39 84 35 66 53 53 51
Culture were grown in 250 ml flasks in the conditions described above. Olive oil and tributyrin were used as substrates.
a Yield is given as the percentage of the initial oleic acid esterified at each time.
stored at 4°C for two months as lyophilised extracts, all the lipases showed a 90 –100% recovery of the enzyme activity previously measured in fresh culture broths. Thus, the stability of all the selected lipases can be regarded good in such storage conditions. A more detailed study on enzyme stability is in progress.
lytic activity, (Table 2) were very active in the synthesis of heptyl oleate (Table 3). On the contrary, lipase from F. oxysporum, which showed a very good activity against both tributyrin and olive oil substrates displayed a moderate activity in the heptyl oleate synthesis. The rest of the enzymes showed similar level of relative activity in both types of assay. From this, we can conclude the absence of a general correlation between hydrolytic and synthetic activities of the crude lipases, as it has been reported by other authors [36]. Therefore, all the following synthetic reactions were carried out with the same amount of catalyst, independently on the producer microorganism.
3.2. Heptyl oleate synthesis The main interest of lipases in organic synthesis is their ability to work in organic media and their broad substrate specificity, as it can be demonstrated with many unnatural substrates [34]. Esterification of oleic acid with 1-heptanol may be considered as a standard reaction to initially verify the activity of lipases in organic medium [35]. Addition of a small amount of water (0.1 ml H2O per ml of isooctane) was found to be necessary for enzyme activity. In the absence of added water, catalytic activity was undetectable. The importance of a small amount of water in this reaction— using lyophilised lipases— has been sufficiently described [35]. As it can be seen in Table 3, all the lipases used in this study gave an almost quantitative esterification of the oleic acid (85–100% yield) after 96 h of reaction (data not shown). The only exception to this was the lipase from A. murorum, with a 77% conversion. In four cases (M. mucoroides, F. poae, F. solani and O. sulphureo-ochraceum) the highest yields were attained after short reaction times (4 to 8 h). These results demonstrate that all the selected lipases are adequate to work in organic medium. Although all the lipases tested were very active in both hydrolytic and synthetic assays, the most active lipases in the hydrolytic assay were not necessarily the most active ones in the synthetic reaction (Table 3). Thus, lipases from F. poae and from F. solani, which displayed a low hydro-
3.3. Resolution of (R,S)-ibuprofen One of the most interesting processes catalysed by lipases in organic media is the resolution of racemic mixtures. Resolution of (R,S)-ibuprofen by stereoselective esterification using lipases as biocatalysts has been extensively studied and demonstrated to be implemented as an interesting process from the industrial point of view [37]. Taking this into account we have employed this reaction as a reference test to analyse the stereoselectivity of the selected lipases with respect to one carboxylic acid with the stereogenic centre in C␣ (Fig. 3A). As in the case of the heptyl oleate, all the lipases tested were active as biocatalysts in the esterification of the ibuprofen but the ester yield, the enantiomeric excess and the enantiopreference of the different enzymes were rather heterogeneous. As it can be seen in Table 4, the highest ester yields were obtained with the lipases from two filamentous fungi, F. oxysporum and O. sulphureo-ochraceum. (84.2 and 70.4%, respectively). The lowest yields were obtained with lipases from A. murorum, P. chrysogenum, Monascus
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F. Cardenas et al. / Enzyme and Microbial Technology 28 (2001) 145–154 Table 5 Resolution of (R, S)-1-phenylethanol by the selected lipases
Table 4 Resolution of (R,S)-ibuprofen by the selected lipases Producing strain
Reaction time (h)
Ester yielda (%)
eeb (%)
Stereopreferencec
Producing strain
Reaction time (h)
Ester yielda (%)
eeb (%)
Stereopreferencec
A. murorum M. mucoroides Monascus sp. A. ciferrii F. poae F. solani F. oxysporum P. chrysogenum O. sulphureo-ochraceum R. araucariae B. linens S. fradiae S. halstedii Streptomyces sp.
264 264 264 336 336 336 264 336 336 336 264 264 336 264
5.5 28.4 7.5 16.3 44.4 10.6 84.2 3.7 70.4 20.9 19.8 4.3 5.7 6
2.6 10.4 7 14.6 14.5 0.7 100 0.2 64.9 10.5 8.8 1.5 3.3 1.5
S S S S R R S R S S R S R R
F. oxysporum O. sulphureo-ochraceum R. araucariae B. linens S. fradiae S. halstedii Streptomyces sp.
240 240 264 264 432 432 432
41 16 2.6 2.4 6.6 4.6 3.6
62 19 2 2.3 5.7 5.2 3.5
R R R R R R R
a
Ester yield is given as the percentage of initial 1-phenylethanol esterified after the reaction time. b Enantiomeric excess of the remaining 1-phenylethanol. c Configuration of the enantiomer found in the ester.
sp. and those from the three actinomycetes. The best enantiomeric excess for the remaining acid was attained once again with the enzymes from F. oxysporum and O. sulphureo-ochraceum (100 and 64.9%, respectively) but the relative catalytic activity was different to that observed in hepthyl oleate synthesis (Table 3). These values are considerably higher than those previously found for the C. antarctica lipase [38] and similar to the values reported for other industrial lipases [39]. The evolution of the esterification of (R,S)-ibuprofen with 1-propanol catalysed by several of the selected lipases is pre-
sented in Fig. 1. Enzymes from O. sulphureo-ochraceum and from F. oxysporum showed a high esterification yield during the first 150 –200 hours of reaction, giving the highest optical purity of the remaining acid. By comparing the results obtained in the esterification of a natural substrate such as oleic acid (Table 3) and of the unnatural acid (R,S)-ibuprofen (Table 4) we can deduce that the active sites of the selected lipases can easily accept the n-alkanoic acid but the recognition of the unnatural chiral acid with an aromatic ring in the stereogenic centre is more problematic, probably due to the high steric hindrance of this molecule. In summary, the F. oxysporum and O. sulphureo-ochraceum lipases seem to be the best biocatalysts for the resolution of this drug, giving the active stereomer ((S)-acid) with high optical purity.
Fig. 1. Fig. 1. Time course of the esterification of (R,S)-ibuprofen with 1-propanol catalysed by several of the selected lipases. Yield is given as the percentage of the racemic ibuprofen esterified at each reaction time. Lipases from O. sulphureo-ochraceum (■), F. oxysporum (䊐), R. araucariae (M. mucoroides (E), F. poae (F) and B. linens (Œ) were used.
Fig. 2. Time course of the transesterification of (R,S)-1-phenylethanol with vinyl acetate catalysed by the lipases from F. oxysporum (E) and from O. sulphureo-ochraceum (F). Yield is given as the percentage of the racemic 1-phenylethanol esterified at each reaction time.
a Ester yield is given as the percentage of initial ibuprofen esterified after the reaction time. b Enantiomeric excess of the remaining ibuprofen. c Configuration of the enantiomer found in the ester.
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Fig. 3. Schematic representation of the reactions catalysed by the lipases from this study. A: Resolution of (R,S)-ibuprofen. B: Resolution of (R,S)-1phenylethanol. C: Acylation of (R) and (S)-glycidol. D: Synthesis of N-1-phenylethyl-butyl-carbamate.
3.4. Resolution of (R,S)-1-phenylethanol by transesterification The resolution of (R,S)-1-phenylethanol by stereoselective transesterification with vinyl acetate was performed as
a test reaction to analyse the steric restrictions of the active subsite of the lipases in the recognition of chiral secondary alcohols (Fig. 3B). The results obtained are shown in Table 5. Only lipases from the filamentous fungi, F. oxysporum and O. sulphureo-ochraceum, gave interesting results in this reaction.
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Table 6 Acylation of (R) and (S)-glycidol by the selected microbial lipases Lipase source O. sulphureo-ochraceum F. oxysporum F. solani F. poae Monascus sp. M. mucoroides P. chrysogenum A. ciferri R. araucariae P. cepacia B. linens S. fradiae S. halstedii
(R)-glycidyl acetate
(S)-glycidyl acetate
R/S ratio
7.8 34 21 6.5 8 2 28 14 9.5 29 15 14.5 7.5
9 41 15 16 7.6 1.5 28 14 9.0 19 12 15.1 11
0.8 0.8 1.4 0.4 1.1 1.1 0.95 1.0 1.1 1.5 1.3 0.95 0.7
Table 7 Acylation of (R) and (S)-glycidol synthesis of N-1-phenylethyl-butylcarbamate by the selected microbial lipases Lipase
Yield (%)a
e.e. (%)b
Preferred configuration
O. sulphureo-ochraceum F. oxysporum F. solani F. poae Monascus sp. M. mucoroides P. chrysogenum A. ciferrii R. araucariae P. cepacia B. linens S. halstedii
6 58 46 47 80 80 38 25 21 53 10 61
99 95 99 85 ⬎10 10 99 40 19 70 99 95
S⬎R R⬎S R R⬎S R⬎S R⬎S R R⬎S R⬎S S⬎R R⬎S R⬎S
a
In the cases where reaction proceeded, the (R)-1-phenylethanol was esterified much faster than the (S)-antipode. Therefore, these lipases showed the same enantiopreference that has been described for commercial enzymes [40,41]. As in the case of the resolution of the chiral acid ((R,S)ibuprofen) the lipases from F. oxysporum and O. sulphureoochraceum were found to be the best biocatalysts. Ester yields of 41 and 16%, respectively, were obtained after 240 h of reaction. It should be noted that these yields are equivalent to a very high ester concentration (410 and 160 mM, respectively) because the initial concentration of the alcohol was 1 M. Both lipases were active for a long period of time in the reaction conditions (Fig. 2). 3.5. Acylation of (R) and (S)-glycidol The acylation of (R) and (S)-glycidol is a process of interest for the obtention of chiral synthons useful in the synthesis of complex molecules [42]. Glycidol was selected as a small substrate model to explore the stereoselectivity of the microbial lipases when acting on a primary alcohol with the chiral carbon in the C␣ position (Fig. 3C). The results obtained are shown in Table 6. The lipases showed a heterogeneous behaviour in this reaction. Lipases from P. cepacia, F. solani and B. linens produced higher amounts of (R)-ester than of the (S)-counterpart. The same enantioselectivity has been previously described for the lipase from another P. cepacia strain [43]. Contrarily, the enzymes from F. oxysporum, O. sulphureoochraceum, S. halstedii and F. poae displayed preference for the (S)-stereomer. Of particular interest is the high enantiomeric excess obtained with the F. poae enzyme, because the lipases described in the literature generally present (R)-enantioselectivity for these kind of substrates [43,44]. 3.6. Resolution of (R,S)-1-phenylethylamine The resolution of (R,S)-1-phenylethylamine was carried out by the alcoxycarbonylation reaction using butyl-vinyl-
Determined by HPLC; b Determined by 1H-NMR using Eu(III) salt.
carbonate as the acylating agent (Fig. 3D). This methodology has provided good enantioselectivities in the resolution of the amines [45]. The data obtained in this reaction are shown in Table 7. From this data we can deduce that the lipases produced by P. chrysogenum, F. poae, F. oxysporum, F. solani and S. halstedii are very interesting as biocatalysts because the high enantiomeric excess obtained, with yields near 50%. The (R)-enantiopreference observed is in accordance with the enantiopreference observed in the acylation of (R,S)-1-phenylethanol. The (S)-enantiopreference observed in the case of the O. sulphureo-ochraceum lipase is not significant due to the low yield achieved. The enantioselectivity obtained with the lipase from P. cepacia is analogous to that observed with the commercial lipase of P. cepacia available from Amano [42]. On the other hand, this lipase was not active in the acylation of (R,S)-1-phenylethanol (Table 5).
4. Conclusions From the results reported in this paper we can deduce that the geometry of the active site of these new lipases is in accordance with the model proposed by Weissfloch and Kazlauskas [46]. The behaviour of the selected lipases in the stereoselective esterification of (R,S)-ibuprofen and of (R,S)-1-phenylethanol showed a considerable heterogeneity, as would be expected bearing in mind the high biodiversity of the lipase-producing microorganisms selected for this study. We conclude that the lipases from F. oxysporum and O. sulphureo-ochraceum showed characteristics which make them very appropriate to be used as biocatalysts in the resolution of these bulky organic molecules such as (R,S)ibuprofen, (R,S)-1-phenylethanol and (R,S) 1-phenylethylamine, with an aromatic ring in the stereogenic centre. Only the F. solani lipase displayed (R)-enantiopreference in the resolution of glycidol. The lipase from S. halstedii is, to our
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knowledge, the first active and stereoselective lipase from actinomycetes useful in the resolution of racemic compounds in organic media. Acknowledgment This paper has been supported by the grant BIO97-0514 from Ministerio de Educacio´n y Ciencia of Spain. References [1] Sztajer H, Maliszewska I, Wierczorek J. Production of exogenous lipases by bacteria, fungi, and actinomycetes. Enzyme Microb Technol 1988;10:492–7. [2] Aires-Barros MR, Taipa MA, Cabral J. Isolation and purification of lipases. In: Wooley P, Peterson SB, editors. Lipases, their structure, biochemistry and application. Cambridge University Press 1994:243–70. [3] Ionita A, Moscovici M, Popa C, Vamanu A, Popa A, Dinu L. Screening of yeast and fungal strains for lipolytic potential and determination of some biochemical properties of microbial lipases. J Mol Catal B: Enzymatic 1997;3:147–51. [4] Jaeger KE, Ransac S, Dijkstra BW, Colson C, van Heuvel M, Misset O. Bacterial lipases. FEMS Microbiol Rev 1994;15:29 – 63. [5] Sto¨cklein W, Sztajer H, Menge U, Schmid RD. Purification and properties of a lipase from Penicillium expansum. Biochim Biophys Acta 1993;1168:181–9. [6] Miura T, Yamane T. Screening for fungi that have lipolytic and acidolytic activities in biomass support particles. Biosci Biotechnol Biochem 1997;61:1252–7. [7] Tomizuka N, Ota Y, Yamada K. Studies on lipase from Candida cylindracea. Agric Biol Chem 1966;30:576 – 84. [8] Lotti M, Grandori R, Fusetti F, Longhi S, Brocca S, Tramontano A, Alberghina L. Cloning and analysis of Candida cylindracea lipase sequences. Gene 1993;124:45–55. [9] Jaeger KE, Liebeton K, Zonta A, Schimossek K, Reetz MT. Biotechnological applications of Pseudomonas aeruginosa lipase: efficient kinetic resolution of amines and alcohols. Appl Microbiol Biotechnol 1996;46:99 –105. [10] Cruz H, Pe´rez C, Wellington E, Castro C, Servı´n-Gonza´lez L. Sequence of the Streptomyces albus G lipase-encoding gene reveals the presence of a prokaryotic lipase family. Gene 1994;144:141–2. [11] Margolin AL. Enzymes in the synthesis of chiral drugs. Enzyme Microb Technol 1993;15:266 – 80. [12] Azerad R. Application of biocatalysts in organic synthesis. Bull Soc Chim Fr 1995;132:17–51. [13] Inada Y, Mahoto F, Sasaki H, Kodera Y, Hiroto M, Nishimura H, Matsushima A. Biomedical and biotechnological applications of PEG and PM-modified proteins. Trends Biotechnol 1995;13:86 –91. [14] Colowick SP, Kaplan NO. Novel techniques for and aspects of immobilised enzymes and cells. In: Mosbach K, editor. Methods in Enzymology. Vol. 137. Immobilised enzymes and cells (Part D). Academic Press Inc, 1988;583– 697. [15] Bell G, Halling PS, Moore BD, Partridge J, Rees DG. Biocatalyst behaviour in low water systems. Trends Biotechnol 1995;13:468 –73. [16] Zaks A, Klibanov AM. The effect of water on enzyme action in organic media. J Biol Chem 1988;263:8017–21. [17] Arroyo M, Sinisterra JV. Influence of the chiral carvones on selectivity of pure lipase B from Candida antarctica. Biotechnol Lett 1995;17:525–30. [18] Clausen IG. Aspects in lipase screening. J Mol Catal B: Enzymatic 1997;3:139 – 46. [19] Nakayama K. Sources of industrial microorganisms. In: Rehm H-J, Reed G, editors. Biotechnology, Vol. 1, Microbial Fundamentals. Verlag Chemie, Weinheim, 1981;355– 410.
153
[20] von Arx JA. Key to the orders of Fungi. In: The genera of fungi sporulating in pure culture. Cramer. Hirschberg, Germany, 1981. [21] Domsch KH, Gams W, Anderson TH. Key to the genera. In: Compendium of soil fungi. IHW, Eching, Germany, 1993. [22] Singler L, Carmichael JW. Taxonomy of Malbranchea and some other Hyphomycetes with arthroconidia. Mycotaxon 1976;4:349 – 88. [23] Frenken LGJ, Egmond MR, Batenburg AM, Wil Bos J, Visser C, Verrips CT. Cloning of the Pseudomonas glumae lipase gene and determination of the active site residues. Appl Environ Microbiol 1992;58:3787–91. [24] Holmes DJ, Cundliffe E. Analysis of a ribosomal RNA methylase gene from Streptomyces tenebrarius which confers resistance to gentamycin. Mol Gen Genet 1991;229:229 –37. [25] Tietz NW, Fiereck EA. A specific method for serum lipase determination. Clin Chim Acta 1966;13:352–7. [26] Sugihara A, Ueshima M, Shimada Y, Tsunasawa S, Tominaga Y. Purification and characterization of a novel thermostable lipase from Pseudomonas cepacia. J Biochem 1992;112:598 – 603. [27] Kojima Y, Yokoe M, Mase T. Purification and characterization of an alkaline lipase from Pseudomonas fluorescens AK 102. Biosci Biotechnol Biochem 1994;58:1564 – 8. [28] Mannesse ML, Cox RC, Koops BC, Verheij HM, de Haas GH, Egmond MR, van der Hijden HT, de Vlieg J. Cutinase from Fusarium solani pisi hydrolyzing triglyceride analogues. Effect of acyl chain length and position in the substrate molecule on activity and enantioselectivity. Biochemistry 1995;34:6400 –7. [29] Chander H, Sannabhadti SS, Elias J, Ranganathan B. Factors affecting lipase production by Penicillium chrysogenum. J Food Sci 1977; 42:1677– 82. [30] Sorhaug T, Ordal ZJ. Cell-bound lipase and esterase of Brevibacterium linens. Appl Microbiol 1974;27:607– 8. [31] Mase T, Matsumiya Y, Akiba T. Purification and characterization of a new lipase from Fusarium sp. YM-30. Biosci Biotechnol Biochem 1995;59:1771–2. [32] Brahimi-Horn MC, Luo SH, Wang SL, Gau SW, Mou DG. Synthesis of hydrolytic enzymes during production of tylosin by Streptomyces fradiae. J Ind Microbiol 1992;10:25–30. [33] Zviagintseva IS, Gorodnianskaia LI. Lipolytic activity of osmotolerant yeasts. Mikrobiologiia 1978;47:230 –3. [34] Zoete MC, van Rantwtjk F, Sheldon RA. Lipase-catalyzed transformation with unnatural acyl-acceptors. Catal Today 1994;22:563–90. [35] Sa´nchez-Montero JM, Hamon V, Thomas D, Legoy MD. Modulation of lipase hydrolysis and synthesis reactions using carbohydrates. Biochim Biophys Acta 1991;1078:345–50. [36] Furutani T, Su R, Ooshima H, Kato J. Simple screening method for lipase transesterification in organic solvent. Enzyme Microb Technol 1995;17:1067–72. [37] Mustranta A. Use of lipases in the resolution of racemic ibuprofen. Appl Microbiol Biotechnol 1992;38:61– 6. [38] Arroyo M, Sinisterra JV. High stereoselective esterification of 2-arylpropionic acids catalyzed by immobilized lipases from Candida antarctica: a mechanistic approach. J Org Chem 1994;59:4410 –7. [39] Goto M, Noda S, Kamiya N, Nakashio F. Enzymatic resolution of racemic ibuprofen by surfactant-coated lipases in organic media. Biotechnol Lett 1996;18:839 – 44. [40] Kazlauskas RJ, Weissfloch ANE. A structure-based rationalization of the enantiopreference of subtilisin toward secondary alcohols and isosteric prymary amines. J Mol Catal B: Enzymatic 1997;3:65–72. [41] Zueg J, Ho¨ning H, Schrag JD, Cygler M. Selectivity of lipases: Conformational analysis of suggested intermediates in ester hydrolysis of chiral primary and secondary alcohols. J Mol Catal B: Enzymatic 1997;3:83–98. [42] Goergens U, Schneider MP. Lipase resolution of 1-t-butylthio-2alkanols: Enzyme mediated routes to enantiomerically pure 1,2-epoxyalkanes and 2-alkanols. Tetrahedron Asymmetry 1992;3:1149 –52.
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[43] Okazaki S, Kamiya N, Goto M, Nakashio F. Enantioselective esterification of glycidol by surfactant-lipase complexes in organic media. Biotechnol Lett 1997;19:541–3. [44] Martins JF, Correa de Sampaio T, Borges de Carvalho I, Barreiros S. Lipase catalyzed esterification of glycidol in nonaqueous solvents: solvent effects on enzymatic activity. Biotechnol Bioeng 1994;44:119 –24.
[45] de Castro-Alvarez MS, Domı´nguez de Maria P, Sinisterra JV. Enzymatic amidation and alkoxycarbonylation of amides using native and immobilised lipases from different origin. A comparative study. Tetrahedron. In press. [46] Weissfloch ANE, Kazlauskas RJ. Enantioprefence of lipase from P. cepacia toward primary alcohols. J Org Chem 1995;60:6959 – 69.
www.elsevier.com/locate/enzmictec
Novel microbial lipases: catalytic activity in reactions in organic media F. Cardenasa,, M.S. de Castrob, J.M. Sanchez-Monterob, J.V. Sinisterrab, M. Valmasedaa, S.W. Elsona, E. Alvareza,* a
SmithKline Beecham, Centro de Investigacio´n Ba´sica, Santiago Grisolı´a, 4, Parque Tecnolo´gico de Madrid, 28760 Tres Cantos, Madrid, Spain b Departamento de Quı´mica Orga´nica y Farmace´utica, Facultad de Farmacia, Universidad Complutense, 28040 Madrid, Spain Received 30 May 2000; accepted 17 July 2000
Abstract 2,000 microbial strains were isolated from soil samples and tested to determine their lipolytic activity by employing screening techniques on solid and in liquid media. Culture broths were initially tested with 1,2-O-dilauryl-rac-glycero-3-glutaric acid-resorufinyl ester, a chromogenic substrate specific for lipases. Fourteen lipase-producing microorganisms were selected and their taxonomic identification was carried out. Hydrolysis of tributyrin or olive oil and the esterification of oleic acid with heptanol were selected to preliminary evaluate the catalytic activity of these lipases. All the selected lipases catalysed this esterification reaction with good yields. Resolution of (R,S)-2-(4isobutylphenyl) propionic acid, (R,S)-1-phenylethanol, (R,S) 1-phenylethylamine and of (R) or (S) glycidol were performed to evaluate the stereoselectivity of these novel enzymes as biocatalysts in reactions in organic media. Lipases from the fungi Fusarium oxysporum and Ovadendron sulphureo-ochraceum gave the best yields and enantioselectivities in the resolution of racemic ibuprofen and 1-phenylethanol. Several lipases displayed a high stereoselectivity in the resolution of chiral amines by an alcoxycarbonylation reaction. © 2001 Elsevier Science Inc. All rights reserved. Keywords: Screening; Microbial lipases; Lipase-catalysed reactions; Resolution of racemic acids; Alcohols and amines
1. Introduction Lipases (triacylglycerol hydrolases EC 3.1.1.3) are enzymes that catalyse the hydrolysis of triglycerides at the oil-water interface. They are produced by animals, plants and microorganisms [1–3]. Microbial lipases have a great potential for commercial applications due to their stability, selectivity and broad substrate specificity [4]. To date, a large number of lipases from filamentous fungi have been extensively studied, both from the biochemical and from the genetic point of view. The most productive species belong to the genera Geotrichum, Penicillium, Aspergillus and Rhizomucor [5,6]. There are also a certain number of lipases produced by yeasts, most of them belonging to the Candida genus, that have been used for biotechnological purposes [7,8]. Lipases from unicellular bacteria, mainly those produced by various species of the genus Pseudomonas, have also proved to be useful both in organic reactions and in the detergent industry. Many of them have been purified, char-
* Corresponding author. Tel.: ⫹34 1 807 4000. E-mail address: [email protected] (E. Alvarez).
acterised and their encoding genes cloned [9]. By contrast, and despite its high biotechnological interest for the production of secondary metabolites and enzymes, the actinomycetes have not been widely exploited for lipase studies, and there is little information available concerning lipaseproducing actinomycetes [1,10]. Among the high number of lipases described in the literature, only the enzymes belonging to a few species have been demonstrated to have adequate stability and biosynthetic capabilities to allow routine use in organic reactions, and hence their consideration as industrially relevant enzymes [11,12]. The application of different techniques of enzyme immobilisation and of chemical modification of enzymes has introduced considerable improvements in the efficiency of certain processes [13,14]. Similarly, a more detailed study on the nature of the reaction medium, involving parameters such as water activity and log P, and the use of optically active solvents has also made important contributions to both the increase in reaction yields, and the selectivity of the lipases [15–17]. However, despite the above mentioned advances in modification and improvement of enzyme activity, there still remain many important problems to be solved in the application of lipases to indus-
0141-0229/01/$ – see front matter © 2001 Elsevier Science Inc. All rights reserved. PII: S 0 1 4 1 - 0 2 2 9 ( 0 0 ) 0 0 2 7 8 - 7
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trial processes. Therefore, access to a wider range of lipase types would be beneficial [18]. In the present study we describe the selection by screening techniques of fourteen lipaseproducing microorganisms and the preliminary characterisation of the lipases as catalysts in reactions in organic media.
2. Materials and methods 2.1. Chemicals The lipase chromogenic substrate 1,2-O-dilauryl-racglycero-3-glutaric acid resorufinyl ester was purchased from Boehringer Mannheim Biochemicals. (R,S)-2-(4-isobutylphenyl)propionic acid (ibuprofen) was obtained from Boots Pharmaceuticals. Yeast extract and Bacto-peptone were purchased from Difco. The other components of the culture media and the chemical reagents were obtained from Merck and from Sigma in the highest purity available. All the reaction products were chemically synthesised and characterised by IR, 1H-NMR, 13C-NMR and elemental analysis. The data obtained agreed with those reported in the literature. These compounds were used as the reference in HPLC, GC and 1H-NMR-Eu analysis. 2.2. Isolation of microorganisms and screening of lipolytic activity All the microorganisms used in this study were isolated from soil samples collected from around the world. The isolation process was performed by serial dilution of the samples according to standard techniques [19]. Taxonomic identification of unicellular bacteria, actinomycetes and yeasts was performed by the Spanish Type Culture Collection (C.E.C.T., Valencia, Spain). The filamentous fungi were identified in-house by using mature cultures on suitable media (normally standard potato dextrose agar and/or oat meal agar) in order to ensure a good development of taxonomically relevant features, and following the identification keys provided by von Arx [20], Domsch et al. [21] and the more specific methodology of Singler and Carmichael [22] in the case of Ovadendron sulphureo-ochraceum. Preliminary screening of lipolytic filamentous fungi, yeasts and bacteria was carried out on BYPO [23] agar plates supplemented with a 10% (v/v) olive oil or tributyrin emulsion prepared in 10% (w/v) gum Arabic solution. Culture plates were incubated at 28°C and periodically examined for four days. Actinomycetes were screened on plates containing NE [24] medium with the same supplement, incubated at 28°C and examined for seven days. Colonies showing clear zones around them were picked out and screened for lipase production in liquid medium. Two different liquid media (H1 and H2) were employed for filamentous fungi. Medium H1 contained 0.5% yeast extract, 3% Bacto-peptone, 0.1% KH2PO4, 0.1% NaNO3, 0.05% MgSO4 and 1% (v/v) olive oil. Medium H2 contained 7.5%
soybean flour, 2% glucose, 0.1% KH2PO4, 0.1% NaNO3, 0.05% MgSO4 and 1% (v/v) olive oil. The final pH was 7.0 for both media. The liquid medium for actinomycetes (M9 medium) contained 1% soybean flour, 1.8% oatmeal, 0.5% yeast extract, 0.5% glucose and 1% (v/v) olive oil. The final pH was adjusted to 7.2. Yeasts were cultured in liquid YED medium and unicellular bacteria were cultured in liquid BYPO medium. All the cultures were grown in 250 ml flasks containing 50 ml of medium and in 2 litre flasks containing 250 ml of medium. Unicellular bacteria, filamentous fungi and yeasts were incubated at 28°C for 3 days and actinomycetes were cultured in the same conditions for 7 days. Flasks were incubated on a reciprocal shaker at 250 rpm. After this time, culture broths were collected, centrifuged and assayed for lipase activity. Positive supernatants were dialysed overnight against 25 mM Tris-HCl buffer, pH 7, and further lyophilised and stored at 4°C. 2.3. Assay of lipase activity Lipase activity present in culture broths was initially monitored using the chromogenic substrate 1,2-O-dilaurylrac-glycero-3-glutaric acid resorufinyl ester. The reaction was carried out at 37°C for 15 min in 96-well microtiter plates using the conditions described by the supplier. The appearance of a red colour due to free resorufinol indicated the presence of lipase activity in the culture broth. Supernatants that hydrolysed the chromogenic substrate were selected in order to quantitatively determine their lipase activity by using olive oil and tributyrin as substrates. This titrimetric assay was carried out essentially as described by Tietz and Fiereck [25]. 1 unit of lipase activity (U) is defined as the amount that releases 1 mol of fatty acid from triglyceride per minute of reaction at 37°C. 2.4. Heptyl oleate synthesis Heptyl oleate synthesis was carried out to verify the catalytic activity of the lipases in the synthesis of esters in organic medium. The reaction mixture was composed of 20 mM oleic acid, 20 mM 1-heptanol, 3 ml of isooctane, 0.3 ml of water and lyophilised crude enzyme. The amount of enzyme used per reaction was that corresponding to 50 ml of the centrifuged culture supernatant. The reaction was carried out at 30°C with vigorous magnetic stirring. Samples of 50 l were withdrawn at different reaction times and diluted in 500 l of isooctane. The amount of ester formed during the reaction was quantified by gas chromatography as described below. 2.5. Resolution of (R,S)-2(4-isobutylphenyl)propionic acid (Ibuprofen) The resolution of (R,S)-ibuprofen was performed by stereoselective esterification using 1-propanol as acyl acceptor. The reaction mixture was composed of 66 mM (R,S)-2-(4-
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isobutylphenyl)propionic acid (ibuprofen), 66 mM 1-propanol, 2 ml of isooctane, 0.2 ml of water and lyophilised enzyme. The amount of enzyme used per reaction was that corresponding to 50 ml of the centrifuged culture supernatant. The reaction was carried out at 30°C with vigorous magnetic stirring. Samples of 30 l were withdrawn at different times and diluted in 500 l of isooctane. The amount of ester formed during the reaction and the enantiomeric excess of the remaining acid were determined by gas chromatography and by HPLC, respectively, as described below. 2.6. Resolution of (R,S)-1-phenylethanol The resolution of (R,S)-1-phenylethanol was performed by stereoselective transesterification using vinyl acetate as acyl donor. The reaction mixture was composed of 1 M (R,S)-1-phenylethanol, 1 M vinyl acetate, 5 ml of isooctane and lyophilised enzyme. The amount of enzyme used per reaction was that corresponding to 50 ml of the centrifuged culture supernatant. The reaction was carried out at 30°C with vigorous magnetic stirring. Samples of 100 l were withdrawn at different times and diluted in 900 l of isooctane. The amount of ester formed during the reaction and the enantiomeric excess of the remaining alcohol were determined by HPLC as described below. 2.7. Acylation of (R) and (S) glycidol The stereoselective acylation of (R) and (S) glycidol was carried out using vinyl acetate as acyl donor. The reaction mixture was composed of 100 mM (R) or (S) glycidol, 100 mM vinyl acetate, 4 ml di-isopropyl ether and 200 mg of lyophilised enzyme. The enzyme was obtained by precipitation of culture broths with (NH4)2SO4, to 90% saturation, dialysed and liophylised. Samples of 100 l were withdrawn and diluted in 900 l of a solution of 4 mM nheptanol (internal standard) in di-isopropyl ether. The amount of ester was determined by gas chromatography as described below. 2.8. Resolution of (R,S) 1-phenylethylamine The resolution of (R,S) 1-phenylethylamine was performed using the lipase-catalysed alcoxycarbonylation of the chiral amine. The reaction mixture was composed of 120 mM butyl-vinyl-carbamate, 120 mM (R,S) 1-phenylethylamine and 200 mg of lyophilised enzyme in 15 ml hexane. The enzyme was obtained by precipitation of culture broths as described above. The reaction products were analysed by HPLC as described below. 2.9. Gas chromatography analysis Gas chromatography analysis of the reaction products was performed using a Shimadzu GC-14 A gas chromato-
147
graph equipped with a FID detector and a SPB-1 column (15 m ⫻ 0.32 mm). For the heptyl oleate analysis the injector and FID temperature was 250°C. The column temperature was kept at 120°C for 5 min, and then programmed to rise 13°C per min to reach 250°C, and maintained at this temperature for 5 min. Nitrogen flux was kept constant at 60 ml/min. For the resolution of (R,S)-ibuprofen the injector temperature was 300°C and the FID temperature was 350°C. The column temperature was kept constant at 180°C and the nitrogen flux was 30 ml/min. For glycidyl acetate synthesis the injector temperature was 200°C and the FID temperature was 250°C. The column temperature was kept constant at 110°C and the nitrogen flux was 13 ml/min. The retention times for glycidyl acetate and heptanol were 2.7 and 4.3 min, respectively. 2.10. HPLC analysis In the resolution of (R,S)-ibuprofen the enantiomeric excess of the remaining acid was determined by HPLC using a Chiracel-OD column. The mobile phase was a mixture of n-hexane:isopropanol:trifluoroacetic acid (100: 1:0.1, v/v/v) at a flow rate of 0.8 ml/min. Detection was performed at 254 nm. The retention times for (R) and (S)-ibuprofen were 19 and 23 min, respectively. The 1-phenylethyl acetate formed during the transesterification reaction and the enantiomeric excess of the remaining alcohol were also determined using a Chiracel-OD column. The mobile phase was a mixture of n-hexane: isopropanol (97:3, v/v) at a flow rate of 0.7 ml/min. In all cases, quantification of the reaction products was performed by measuring their absorbance at 254 nm. The retention times for (R) and (S)1-phenylethanol, and for (R,S)1-phenylethylacetate were 17, 19 and 7 min, respectively. The yield of butyl-(R,S) 1-phenylethyl carbamate was determined by HPLC using a Nucleosil 120 C18 5 m column (25 ⫻ 0.46 cm) from Tracer Analytical. The mobile phase was H2O/MeOH (50:50, v/v). The flux was 0.3 ml/ min. Detection was performed at 254 nm. The retention times for the carbamate and the amine were 29.6 and 27.5 min, respectively. 2.11. Enantiomeric excess of carbamates The enantiomeric excess of butyl-(R,S)-1-phenylethylcarbamate produced was measured using europium (III) tris-[3-(heptafluoropropylhydroximethylen)-(⫹)-camphorate], using the 1H-NMR technique. The carbamate was isolated from the reaction mixture by column chromatography on neutral silica, using hexane-ethyl acetate (7:3, v/v) as eluent. The carbamate/Eu salt ratio used was 1/0.4 (mol/ mol) and the chemical shifts of CH were ␦ ⫽ 6.3 ppm for the (S)-isomer, and ␦ ⫽ 5.8 ppm for the (R) isomer. The enantiomeric excess was determined by comparison of the area of both peaks.
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Table 1 Number and distribution in microbial groups of the microorganisms selected in this study through the screening process Screening step
Soil isolation Solid medium screening Liquid medium screening (chromogenic assay) Liquid medium screening (titrimetric assay)
Number of selected microorganisms
Distribution in microbial groups Bacteria
Actinomycetes
Yeasts
Filamentous fungi
2,000 450 118
200 181 11
840 83 56
100 61 6
860 125 45
14
1
3
1
9
3. Results and discussion 3.1. Selection of lipolytic microorganisms The use of solid media supplemented with emulsified triglycerides is a standard methodology for the selection of lipase-producing microorganisms [26,27]. A total of 2,000 microorganisms isolated from soil samples, including unicellular bacteria (200 strains), actinomycetes (840 strains), yeasts (100 strains) and filamentous fungi (860 strains), were employed in this work (see Table 1). All the microbial strains were screened for lipase activity in solid medium plates containing olive oil or tributyrin as indicator. A total of 450 microorganisms produced a clear halo around them in plates containing tributyrin. Of these, only 92 microorganisms also showed hydrolysis on the olive oil plates, thus indicating a predominant presence of lipases acting on short chain triglycerides. The number of microorganisms that produced detectable levels of lipase in solid media represents 22% of the tested microorganisms, which we consider to be a very high percentage of lipase producing species. It should be noted that each microorganism has been only tested for lipase production in one solid culture medium, which probably would not be appropriate for the lipase induction from many microorganisms. The use of a set of different media for the screening could probably allow to the detection of a higher number of lipase producing microorganisms. The 450 microorganisms selected in the first step were cultured in liquid medium using olive oil as lipase inducer. After centrifugation of culture broths, supernatants were tested with 1,2-O-dilauryl-rac-glycero-3-glutaric acid resorufinyl ester, a chromogenic substrate specific for lipases. The appearance of a red colour due to the release of resorufinol indicated that the supernatant had lipase activity. 118 culture broths were positive in this qualitative assay and were used to further quantify their lipase activity by using tributyrin and olive oil as substrates. During this step, the relative distribution of the selected microorganisms suffered a new dramatic change with respect to the distribution found in the previous step. The lipase activity against the chromogenic substrate displayed by most of the selected bacteria
was very low and only a small percentage were selected for the next selection step. By contrast, a high percentage of actinomycetes were positive in this assay (Table 1). This difference between the bacterial and actinomycetes lipases against the chromogenic substrate could be due to differences in the level of lipase production in liquid media, or to different substrate specificity of the lipases from both groups. Hydrolytic activity of the lipases selected in the previous step was determined by using tributyrin and olive oil as substrates. From this analysis, 14 strains were selected showing the best lipase yields (lipase activity higher than 1 U/ml for one or for both triglycerides) and the best reproducibility in liquid medium cultures. The producing strains were taxonomically identified as follows: nine filamentous fungi (Acremonium murorum (Corda) W. Gams, Monascus mucoroides van Tiegh, Monascus sp., Arthroderma ciferrii Vars & Ajello, Fusarium poae (Pec.) Wollenw, Fusarium solani (Mart.) Sacc., Fusarium oxysporum, Penicillium chrysogenum Thom, Ovadendron sulphureo-ochraceum (van Beyma) Singler & Carmichael, one yeast (Rhodotorula araucariae), one unicellular bacterium (Brevibacterium linens) and three actinomycetes (Streptomyces fradiae, Streptomyces halstedii, Streptomyces sp.). As far as we know, A. murorum, M. mucoroides, A. ciferrii, F. poae, O. sulphureoochraceum, R. araucariae and S. halstedii are reported here for the first time as lipase producers. F. solani has been reported to produce a very active cutinase that also displays triacylglycerol lipase activity [28]. F. oxysporum, P. chrysogenum, B. linens and S. fradiae have also been previously found to have lipolytic activity in solid and in liquid media [1,29,30,31]. In addition, other strains belonging to Monascus [1,6], Fusarium [31], Streptomyces [32] and Rhodotorula [33] genus have been reported to show lipolytic activity. As shown in Table 2, all the selected lipases displayed better hydrolytic activity against tributyrin than against olive oil, a general characteristic previously observed during the screening in solid medium. In general, lipase yields obtained for microorganisms cultured in 250 ml flasks were similar to those obtained in 2 l flasks (data not shown). When stored at 4°C as culture broths, all the selected lipases maintained their activity for at least two weeks. When
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Table 2 Lipase activity (U/ml of supernatant) measured in culture broths from the fourteen microorganisms selected in this study
Table 3 Time course of heptyl oleate synthesis performed by the lipases selected in this study
Producing strain
Producing strain
A. murorum M. mucoroides Monascus sp. A. ciferrii F. poae F. solani F. oxysporum P. chrysogenum O. sulphureo-ochraceum R. araucariae B. linens S. fradiae S. halstedii Streptomyces sp.
Culture medium
Lipase activity (U/ml) Olive oil
Tributyrin
H1 H1 H1 H2 H2 H2 H2 H2 H2 YED BYPO M9 M9 M9
0.72 3.02 2.35 1.43 0.93 0.60 2.83 0.64 4.33 0.75 0.35 0.89 0.70 1.16
5.18 4.31 2.37 1.89 1.47 1.25 5.70 4.68 6.10 3.64 1.83 3.16 1.27 2.10
A. murorum M. mucoroides Monascus sp. A. ciferrii F. poae F. solani F. oxysporum P. chrysogenum O. sulphureo-ochraceum R. araucariae B. linens S. fradiae S. halstedii Streptomyces sp.
Yielda (%) 4h
8h
6 95 34 41 77 79 28 11 71 23 44 44 38 24
29 96 49 77 89 83 52 39 84 35 66 53 53 51
Culture were grown in 250 ml flasks in the conditions described above. Olive oil and tributyrin were used as substrates.
a Yield is given as the percentage of the initial oleic acid esterified at each time.
stored at 4°C for two months as lyophilised extracts, all the lipases showed a 90 –100% recovery of the enzyme activity previously measured in fresh culture broths. Thus, the stability of all the selected lipases can be regarded good in such storage conditions. A more detailed study on enzyme stability is in progress.
lytic activity, (Table 2) were very active in the synthesis of heptyl oleate (Table 3). On the contrary, lipase from F. oxysporum, which showed a very good activity against both tributyrin and olive oil substrates displayed a moderate activity in the heptyl oleate synthesis. The rest of the enzymes showed similar level of relative activity in both types of assay. From this, we can conclude the absence of a general correlation between hydrolytic and synthetic activities of the crude lipases, as it has been reported by other authors [36]. Therefore, all the following synthetic reactions were carried out with the same amount of catalyst, independently on the producer microorganism.
3.2. Heptyl oleate synthesis The main interest of lipases in organic synthesis is their ability to work in organic media and their broad substrate specificity, as it can be demonstrated with many unnatural substrates [34]. Esterification of oleic acid with 1-heptanol may be considered as a standard reaction to initially verify the activity of lipases in organic medium [35]. Addition of a small amount of water (0.1 ml H2O per ml of isooctane) was found to be necessary for enzyme activity. In the absence of added water, catalytic activity was undetectable. The importance of a small amount of water in this reaction— using lyophilised lipases— has been sufficiently described [35]. As it can be seen in Table 3, all the lipases used in this study gave an almost quantitative esterification of the oleic acid (85–100% yield) after 96 h of reaction (data not shown). The only exception to this was the lipase from A. murorum, with a 77% conversion. In four cases (M. mucoroides, F. poae, F. solani and O. sulphureo-ochraceum) the highest yields were attained after short reaction times (4 to 8 h). These results demonstrate that all the selected lipases are adequate to work in organic medium. Although all the lipases tested were very active in both hydrolytic and synthetic assays, the most active lipases in the hydrolytic assay were not necessarily the most active ones in the synthetic reaction (Table 3). Thus, lipases from F. poae and from F. solani, which displayed a low hydro-
3.3. Resolution of (R,S)-ibuprofen One of the most interesting processes catalysed by lipases in organic media is the resolution of racemic mixtures. Resolution of (R,S)-ibuprofen by stereoselective esterification using lipases as biocatalysts has been extensively studied and demonstrated to be implemented as an interesting process from the industrial point of view [37]. Taking this into account we have employed this reaction as a reference test to analyse the stereoselectivity of the selected lipases with respect to one carboxylic acid with the stereogenic centre in C␣ (Fig. 3A). As in the case of the heptyl oleate, all the lipases tested were active as biocatalysts in the esterification of the ibuprofen but the ester yield, the enantiomeric excess and the enantiopreference of the different enzymes were rather heterogeneous. As it can be seen in Table 4, the highest ester yields were obtained with the lipases from two filamentous fungi, F. oxysporum and O. sulphureo-ochraceum. (84.2 and 70.4%, respectively). The lowest yields were obtained with lipases from A. murorum, P. chrysogenum, Monascus
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F. Cardenas et al. / Enzyme and Microbial Technology 28 (2001) 145–154 Table 5 Resolution of (R, S)-1-phenylethanol by the selected lipases
Table 4 Resolution of (R,S)-ibuprofen by the selected lipases Producing strain
Reaction time (h)
Ester yielda (%)
eeb (%)
Stereopreferencec
Producing strain
Reaction time (h)
Ester yielda (%)
eeb (%)
Stereopreferencec
A. murorum M. mucoroides Monascus sp. A. ciferrii F. poae F. solani F. oxysporum P. chrysogenum O. sulphureo-ochraceum R. araucariae B. linens S. fradiae S. halstedii Streptomyces sp.
264 264 264 336 336 336 264 336 336 336 264 264 336 264
5.5 28.4 7.5 16.3 44.4 10.6 84.2 3.7 70.4 20.9 19.8 4.3 5.7 6
2.6 10.4 7 14.6 14.5 0.7 100 0.2 64.9 10.5 8.8 1.5 3.3 1.5
S S S S R R S R S S R S R R
F. oxysporum O. sulphureo-ochraceum R. araucariae B. linens S. fradiae S. halstedii Streptomyces sp.
240 240 264 264 432 432 432
41 16 2.6 2.4 6.6 4.6 3.6
62 19 2 2.3 5.7 5.2 3.5
R R R R R R R
a
Ester yield is given as the percentage of initial 1-phenylethanol esterified after the reaction time. b Enantiomeric excess of the remaining 1-phenylethanol. c Configuration of the enantiomer found in the ester.
sp. and those from the three actinomycetes. The best enantiomeric excess for the remaining acid was attained once again with the enzymes from F. oxysporum and O. sulphureo-ochraceum (100 and 64.9%, respectively) but the relative catalytic activity was different to that observed in hepthyl oleate synthesis (Table 3). These values are considerably higher than those previously found for the C. antarctica lipase [38] and similar to the values reported for other industrial lipases [39]. The evolution of the esterification of (R,S)-ibuprofen with 1-propanol catalysed by several of the selected lipases is pre-
sented in Fig. 1. Enzymes from O. sulphureo-ochraceum and from F. oxysporum showed a high esterification yield during the first 150 –200 hours of reaction, giving the highest optical purity of the remaining acid. By comparing the results obtained in the esterification of a natural substrate such as oleic acid (Table 3) and of the unnatural acid (R,S)-ibuprofen (Table 4) we can deduce that the active sites of the selected lipases can easily accept the n-alkanoic acid but the recognition of the unnatural chiral acid with an aromatic ring in the stereogenic centre is more problematic, probably due to the high steric hindrance of this molecule. In summary, the F. oxysporum and O. sulphureo-ochraceum lipases seem to be the best biocatalysts for the resolution of this drug, giving the active stereomer ((S)-acid) with high optical purity.
Fig. 1. Fig. 1. Time course of the esterification of (R,S)-ibuprofen with 1-propanol catalysed by several of the selected lipases. Yield is given as the percentage of the racemic ibuprofen esterified at each reaction time. Lipases from O. sulphureo-ochraceum (■), F. oxysporum (䊐), R. araucariae (M. mucoroides (E), F. poae (F) and B. linens (Œ) were used.
Fig. 2. Time course of the transesterification of (R,S)-1-phenylethanol with vinyl acetate catalysed by the lipases from F. oxysporum (E) and from O. sulphureo-ochraceum (F). Yield is given as the percentage of the racemic 1-phenylethanol esterified at each reaction time.
a Ester yield is given as the percentage of initial ibuprofen esterified after the reaction time. b Enantiomeric excess of the remaining ibuprofen. c Configuration of the enantiomer found in the ester.
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151
Fig. 3. Schematic representation of the reactions catalysed by the lipases from this study. A: Resolution of (R,S)-ibuprofen. B: Resolution of (R,S)-1phenylethanol. C: Acylation of (R) and (S)-glycidol. D: Synthesis of N-1-phenylethyl-butyl-carbamate.
3.4. Resolution of (R,S)-1-phenylethanol by transesterification The resolution of (R,S)-1-phenylethanol by stereoselective transesterification with vinyl acetate was performed as
a test reaction to analyse the steric restrictions of the active subsite of the lipases in the recognition of chiral secondary alcohols (Fig. 3B). The results obtained are shown in Table 5. Only lipases from the filamentous fungi, F. oxysporum and O. sulphureo-ochraceum, gave interesting results in this reaction.
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Table 6 Acylation of (R) and (S)-glycidol by the selected microbial lipases Lipase source O. sulphureo-ochraceum F. oxysporum F. solani F. poae Monascus sp. M. mucoroides P. chrysogenum A. ciferri R. araucariae P. cepacia B. linens S. fradiae S. halstedii
(R)-glycidyl acetate
(S)-glycidyl acetate
R/S ratio
7.8 34 21 6.5 8 2 28 14 9.5 29 15 14.5 7.5
9 41 15 16 7.6 1.5 28 14 9.0 19 12 15.1 11
0.8 0.8 1.4 0.4 1.1 1.1 0.95 1.0 1.1 1.5 1.3 0.95 0.7
Table 7 Acylation of (R) and (S)-glycidol synthesis of N-1-phenylethyl-butylcarbamate by the selected microbial lipases Lipase
Yield (%)a
e.e. (%)b
Preferred configuration
O. sulphureo-ochraceum F. oxysporum F. solani F. poae Monascus sp. M. mucoroides P. chrysogenum A. ciferrii R. araucariae P. cepacia B. linens S. halstedii
6 58 46 47 80 80 38 25 21 53 10 61
99 95 99 85 ⬎10 10 99 40 19 70 99 95
S⬎R R⬎S R R⬎S R⬎S R⬎S R R⬎S R⬎S S⬎R R⬎S R⬎S
a
In the cases where reaction proceeded, the (R)-1-phenylethanol was esterified much faster than the (S)-antipode. Therefore, these lipases showed the same enantiopreference that has been described for commercial enzymes [40,41]. As in the case of the resolution of the chiral acid ((R,S)ibuprofen) the lipases from F. oxysporum and O. sulphureoochraceum were found to be the best biocatalysts. Ester yields of 41 and 16%, respectively, were obtained after 240 h of reaction. It should be noted that these yields are equivalent to a very high ester concentration (410 and 160 mM, respectively) because the initial concentration of the alcohol was 1 M. Both lipases were active for a long period of time in the reaction conditions (Fig. 2). 3.5. Acylation of (R) and (S)-glycidol The acylation of (R) and (S)-glycidol is a process of interest for the obtention of chiral synthons useful in the synthesis of complex molecules [42]. Glycidol was selected as a small substrate model to explore the stereoselectivity of the microbial lipases when acting on a primary alcohol with the chiral carbon in the C␣ position (Fig. 3C). The results obtained are shown in Table 6. The lipases showed a heterogeneous behaviour in this reaction. Lipases from P. cepacia, F. solani and B. linens produced higher amounts of (R)-ester than of the (S)-counterpart. The same enantioselectivity has been previously described for the lipase from another P. cepacia strain [43]. Contrarily, the enzymes from F. oxysporum, O. sulphureoochraceum, S. halstedii and F. poae displayed preference for the (S)-stereomer. Of particular interest is the high enantiomeric excess obtained with the F. poae enzyme, because the lipases described in the literature generally present (R)-enantioselectivity for these kind of substrates [43,44]. 3.6. Resolution of (R,S)-1-phenylethylamine The resolution of (R,S)-1-phenylethylamine was carried out by the alcoxycarbonylation reaction using butyl-vinyl-
Determined by HPLC; b Determined by 1H-NMR using Eu(III) salt.
carbonate as the acylating agent (Fig. 3D). This methodology has provided good enantioselectivities in the resolution of the amines [45]. The data obtained in this reaction are shown in Table 7. From this data we can deduce that the lipases produced by P. chrysogenum, F. poae, F. oxysporum, F. solani and S. halstedii are very interesting as biocatalysts because the high enantiomeric excess obtained, with yields near 50%. The (R)-enantiopreference observed is in accordance with the enantiopreference observed in the acylation of (R,S)-1-phenylethanol. The (S)-enantiopreference observed in the case of the O. sulphureo-ochraceum lipase is not significant due to the low yield achieved. The enantioselectivity obtained with the lipase from P. cepacia is analogous to that observed with the commercial lipase of P. cepacia available from Amano [42]. On the other hand, this lipase was not active in the acylation of (R,S)-1-phenylethanol (Table 5).
4. Conclusions From the results reported in this paper we can deduce that the geometry of the active site of these new lipases is in accordance with the model proposed by Weissfloch and Kazlauskas [46]. The behaviour of the selected lipases in the stereoselective esterification of (R,S)-ibuprofen and of (R,S)-1-phenylethanol showed a considerable heterogeneity, as would be expected bearing in mind the high biodiversity of the lipase-producing microorganisms selected for this study. We conclude that the lipases from F. oxysporum and O. sulphureo-ochraceum showed characteristics which make them very appropriate to be used as biocatalysts in the resolution of these bulky organic molecules such as (R,S)ibuprofen, (R,S)-1-phenylethanol and (R,S) 1-phenylethylamine, with an aromatic ring in the stereogenic centre. Only the F. solani lipase displayed (R)-enantiopreference in the resolution of glycidol. The lipase from S. halstedii is, to our
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knowledge, the first active and stereoselective lipase from actinomycetes useful in the resolution of racemic compounds in organic media. Acknowledgment This paper has been supported by the grant BIO97-0514 from Ministerio de Educacio´n y Ciencia of Spain. References [1] Sztajer H, Maliszewska I, Wierczorek J. Production of exogenous lipases by bacteria, fungi, and actinomycetes. Enzyme Microb Technol 1988;10:492–7. [2] Aires-Barros MR, Taipa MA, Cabral J. Isolation and purification of lipases. In: Wooley P, Peterson SB, editors. Lipases, their structure, biochemistry and application. Cambridge University Press 1994:243–70. [3] Ionita A, Moscovici M, Popa C, Vamanu A, Popa A, Dinu L. Screening of yeast and fungal strains for lipolytic potential and determination of some biochemical properties of microbial lipases. J Mol Catal B: Enzymatic 1997;3:147–51. [4] Jaeger KE, Ransac S, Dijkstra BW, Colson C, van Heuvel M, Misset O. Bacterial lipases. FEMS Microbiol Rev 1994;15:29 – 63. [5] Sto¨cklein W, Sztajer H, Menge U, Schmid RD. Purification and properties of a lipase from Penicillium expansum. Biochim Biophys Acta 1993;1168:181–9. [6] Miura T, Yamane T. Screening for fungi that have lipolytic and acidolytic activities in biomass support particles. Biosci Biotechnol Biochem 1997;61:1252–7. [7] Tomizuka N, Ota Y, Yamada K. Studies on lipase from Candida cylindracea. Agric Biol Chem 1966;30:576 – 84. [8] Lotti M, Grandori R, Fusetti F, Longhi S, Brocca S, Tramontano A, Alberghina L. Cloning and analysis of Candida cylindracea lipase sequences. Gene 1993;124:45–55. [9] Jaeger KE, Liebeton K, Zonta A, Schimossek K, Reetz MT. Biotechnological applications of Pseudomonas aeruginosa lipase: efficient kinetic resolution of amines and alcohols. Appl Microbiol Biotechnol 1996;46:99 –105. [10] Cruz H, Pe´rez C, Wellington E, Castro C, Servı´n-Gonza´lez L. Sequence of the Streptomyces albus G lipase-encoding gene reveals the presence of a prokaryotic lipase family. Gene 1994;144:141–2. [11] Margolin AL. Enzymes in the synthesis of chiral drugs. Enzyme Microb Technol 1993;15:266 – 80. [12] Azerad R. Application of biocatalysts in organic synthesis. Bull Soc Chim Fr 1995;132:17–51. [13] Inada Y, Mahoto F, Sasaki H, Kodera Y, Hiroto M, Nishimura H, Matsushima A. Biomedical and biotechnological applications of PEG and PM-modified proteins. Trends Biotechnol 1995;13:86 –91. [14] Colowick SP, Kaplan NO. Novel techniques for and aspects of immobilised enzymes and cells. In: Mosbach K, editor. Methods in Enzymology. Vol. 137. Immobilised enzymes and cells (Part D). Academic Press Inc, 1988;583– 697. [15] Bell G, Halling PS, Moore BD, Partridge J, Rees DG. Biocatalyst behaviour in low water systems. Trends Biotechnol 1995;13:468 –73. [16] Zaks A, Klibanov AM. The effect of water on enzyme action in organic media. J Biol Chem 1988;263:8017–21. [17] Arroyo M, Sinisterra JV. Influence of the chiral carvones on selectivity of pure lipase B from Candida antarctica. Biotechnol Lett 1995;17:525–30. [18] Clausen IG. Aspects in lipase screening. J Mol Catal B: Enzymatic 1997;3:139 – 46. [19] Nakayama K. Sources of industrial microorganisms. In: Rehm H-J, Reed G, editors. Biotechnology, Vol. 1, Microbial Fundamentals. Verlag Chemie, Weinheim, 1981;355– 410.
153
[20] von Arx JA. Key to the orders of Fungi. In: The genera of fungi sporulating in pure culture. Cramer. Hirschberg, Germany, 1981. [21] Domsch KH, Gams W, Anderson TH. Key to the genera. In: Compendium of soil fungi. IHW, Eching, Germany, 1993. [22] Singler L, Carmichael JW. Taxonomy of Malbranchea and some other Hyphomycetes with arthroconidia. Mycotaxon 1976;4:349 – 88. [23] Frenken LGJ, Egmond MR, Batenburg AM, Wil Bos J, Visser C, Verrips CT. Cloning of the Pseudomonas glumae lipase gene and determination of the active site residues. Appl Environ Microbiol 1992;58:3787–91. [24] Holmes DJ, Cundliffe E. Analysis of a ribosomal RNA methylase gene from Streptomyces tenebrarius which confers resistance to gentamycin. Mol Gen Genet 1991;229:229 –37. [25] Tietz NW, Fiereck EA. A specific method for serum lipase determination. Clin Chim Acta 1966;13:352–7. [26] Sugihara A, Ueshima M, Shimada Y, Tsunasawa S, Tominaga Y. Purification and characterization of a novel thermostable lipase from Pseudomonas cepacia. J Biochem 1992;112:598 – 603. [27] Kojima Y, Yokoe M, Mase T. Purification and characterization of an alkaline lipase from Pseudomonas fluorescens AK 102. Biosci Biotechnol Biochem 1994;58:1564 – 8. [28] Mannesse ML, Cox RC, Koops BC, Verheij HM, de Haas GH, Egmond MR, van der Hijden HT, de Vlieg J. Cutinase from Fusarium solani pisi hydrolyzing triglyceride analogues. Effect of acyl chain length and position in the substrate molecule on activity and enantioselectivity. Biochemistry 1995;34:6400 –7. [29] Chander H, Sannabhadti SS, Elias J, Ranganathan B. Factors affecting lipase production by Penicillium chrysogenum. J Food Sci 1977; 42:1677– 82. [30] Sorhaug T, Ordal ZJ. Cell-bound lipase and esterase of Brevibacterium linens. Appl Microbiol 1974;27:607– 8. [31] Mase T, Matsumiya Y, Akiba T. Purification and characterization of a new lipase from Fusarium sp. YM-30. Biosci Biotechnol Biochem 1995;59:1771–2. [32] Brahimi-Horn MC, Luo SH, Wang SL, Gau SW, Mou DG. Synthesis of hydrolytic enzymes during production of tylosin by Streptomyces fradiae. J Ind Microbiol 1992;10:25–30. [33] Zviagintseva IS, Gorodnianskaia LI. Lipolytic activity of osmotolerant yeasts. Mikrobiologiia 1978;47:230 –3. [34] Zoete MC, van Rantwtjk F, Sheldon RA. Lipase-catalyzed transformation with unnatural acyl-acceptors. Catal Today 1994;22:563–90. [35] Sa´nchez-Montero JM, Hamon V, Thomas D, Legoy MD. Modulation of lipase hydrolysis and synthesis reactions using carbohydrates. Biochim Biophys Acta 1991;1078:345–50. [36] Furutani T, Su R, Ooshima H, Kato J. Simple screening method for lipase transesterification in organic solvent. Enzyme Microb Technol 1995;17:1067–72. [37] Mustranta A. Use of lipases in the resolution of racemic ibuprofen. Appl Microbiol Biotechnol 1992;38:61– 6. [38] Arroyo M, Sinisterra JV. High stereoselective esterification of 2-arylpropionic acids catalyzed by immobilized lipases from Candida antarctica: a mechanistic approach. J Org Chem 1994;59:4410 –7. [39] Goto M, Noda S, Kamiya N, Nakashio F. Enzymatic resolution of racemic ibuprofen by surfactant-coated lipases in organic media. Biotechnol Lett 1996;18:839 – 44. [40] Kazlauskas RJ, Weissfloch ANE. A structure-based rationalization of the enantiopreference of subtilisin toward secondary alcohols and isosteric prymary amines. J Mol Catal B: Enzymatic 1997;3:65–72. [41] Zueg J, Ho¨ning H, Schrag JD, Cygler M. Selectivity of lipases: Conformational analysis of suggested intermediates in ester hydrolysis of chiral primary and secondary alcohols. J Mol Catal B: Enzymatic 1997;3:83–98. [42] Goergens U, Schneider MP. Lipase resolution of 1-t-butylthio-2alkanols: Enzyme mediated routes to enantiomerically pure 1,2-epoxyalkanes and 2-alkanols. Tetrahedron Asymmetry 1992;3:1149 –52.
154
F. Cardenas et al. / Enzyme and Microbial Technology 28 (2001) 145–154
[43] Okazaki S, Kamiya N, Goto M, Nakashio F. Enantioselective esterification of glycidol by surfactant-lipase complexes in organic media. Biotechnol Lett 1997;19:541–3. [44] Martins JF, Correa de Sampaio T, Borges de Carvalho I, Barreiros S. Lipase catalyzed esterification of glycidol in nonaqueous solvents: solvent effects on enzymatic activity. Biotechnol Bioeng 1994;44:119 –24.
[45] de Castro-Alvarez MS, Domı´nguez de Maria P, Sinisterra JV. Enzymatic amidation and alkoxycarbonylation of amides using native and immobilised lipases from different origin. A comparative study. Tetrahedron. In press. [46] Weissfloch ANE, Kazlauskas RJ. Enantioprefence of lipase from P. cepacia toward primary alcohols. J Org Chem 1995;60:6959 – 69.