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Screening, purification and characterization of the thermoalkalophilic lipase produced by Bacillus thermoleovorans CCR11
Enzyme and Microbial Technology 37 (2005) 648–654
Screening, purification and characterization of the thermoalkalophilic lipase produced by Bacillus thermoleovorans CCR11 Lelie D. Castro-Ochoa, Citlali Rodr´ıguez-G´omez, Gerardo Valerio-Alfaro, Rosamar´ıa Oliart Ros ∗ Unidad de Investigaci´on y Desarrollo en Alimentos, Instituto Tecnol´ogico de Veracruz. Av. Miguel A. de Quevedo 2779, Veracruz, Ver, 91897 Veracruz, M´exico Accepted 20 June 2005
Abstract Among eleven aerobic thermophilic Bacillus strains, isolated from “El Carrizal” hot springs in Veracruz, Mexico, Bacillus thermoleovorans CCR11 (EMBL # AJ536599) was selected for lipase characterization because of its high lipase specific activity. Lipase was purified by diafiltration (polyethersulfone ultrafiltration membrane co500,000), and preparative isoelectrofocusing. The lipase had a relative molecular mass of 11 kDa (the lowest Mr reported), although it formed higher molecular weight aggregates in native form. The optimum catalytic conditions for Bacillus thermoleovorans CCR11 lipase were 60 ◦ C and pH 9–10. Hg2+ , PMSF, SDS, Tween 80 and Tween 20 had an inhibitory effect on lipase activity, whereas Ca2+ salts and Triton X-100 increased it. Lipase activity was compatible with the presence of organic solvents, except for butanol. Lipase showed a notable preference for C6–C10 p-nitrophenyl esters, with the highest activity toward p-nitrophenyl caproate (C10). Lipase stability in the presence of organic solvents, as well as in acidic and alkaline pHs and at high temperatures makes it a good candidate for its application in non-aqueous biocatalysis. © 2005 Elsevier Inc. All rights reserved. Keywords: Lipase; Bacillus thermoleovorans; Thermophile; Enzyme characterization
1. Introduction Lipases (triacylglycerol acylhydrolases; EC 3.1.1.3) constitute a group of enzymes defined as carboxylesterases that catalyze the hydrolysis (and synthesis) of long chain acylglycerols at the lipid-water interface. Microbial lipases have been widely used for biotechnological applications in detergents, dairy and textile industries, production of surfactants, and oil processing. In recent years, lipases have received considerable attention with regard to the preparation of enantiomerically pure pharmaceuticals, since they have a number of unique characteristics: substrate specificity, regio-specificity, and chiral selectivity [1]. Given that such reactions are sometimes performed most
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efficiently at elevated temperatures and in organic solvents, converging attempts have been made to find thermostable lipases which would have advantages over labile enzymes in such applications [2]. In recent years, a number of thermophilic microorganisms producing thermoactive lipases and esterases have been purified and characterized [3–9]. Thermophilic lipases show higher thermostability, higher activity at elevated temperatures, and often show more resistance to chemical denaturation, making them ideal tools in industrial and chemical procesess where relatively high reaction temperatures and/or organic solvents are used. As each industrial application may require specific properties of the biocatalysts, there is still an interest in finding new lipases that could create novel applications. In this study, we describe the purification and characterization of a thermoalkaliphilic lipase produced by the thermophilic strain Bacillus thermoleovorans CCR11, isolated from a hot spring in Veracruz, Mexico.
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2. Materials and methods 2.1. Bacterial strains Eleven aerobic thermophilic Bacillus strains, isolated from “El Carrizal” hot springs in Veracruz, Mexico, were analyzed for lipolytic activity. Bacillus thermoleovorans CCR11 (EMBL # AJ536599) was selected for lipase production and characterization because it showed the highest lipase specific activity. 2.2. Screening All eleven thermophilic Bacillus strains were screened for lipase activity on agar plates containing Rhodamine B 0.001% (w/v), nutrient broth 0.8% (w/v), NaCl 0.4% (w/v), agar 1% (w/v), and olive oil 3%, in distilled water, pH 6.5 [10]. Plates were incubated at 55 ◦ C for 18 h, and lipase production was identified as an orange halo around colonies under UV light at 350 nm. In order to select the best lipase producer for enzyme purification and characterization, strains with lipolytic activity on the plates were cultured in liquid medium (nutrient broth 0.325% (w/v), CaCl2 0.1% (w/v), olive oil 2.5%, and gum arabic 1%, pH 6.5, 55 ◦ C, 150 rpm [3]), and lipase activity was determined spectrophotometrically with p-nitrophenyl-laurate as substrate (see below). 2.3. Assay for lipolytic activity Lipolytic activity was determined by a spectrophotometric assay using p-nitrophenil-laurate (pNPL) as substrate [5]. The reaction mixture consisted of: 0.1 ml enzyme extract, 0.8 ml 0.05 M phosphate buffer (pH 6.5), and 0.1 ml 0.01 M pNPL in ethanol. The hydrolytic reaction was carried out at 60 ◦ C for 30 min, after which 0.25 ml of 0.1 M Na2 CO3 was added. The mixture was centrifuged (16,000 × g, 15 min, 25 ◦ C) and the absorbance at 410 nm was determined. One unit of lipase activity was defined as the amount of enzyme that caused the release of 1 mol of p-nitrophenol (molar absorption coefficient 4.6 mM−1 cm−1 ) from pNP-laurate in 30 min under test conditions. 2.4. Lipase production Bacillus thermoleovorans CCR11 was grown in a liquid medium containing nutrient broth 0.325% (w/v), CaCl2 0.1% (w/v), olive oil 2.5%, and gum arabic 1%, pH 6.5 [3]. Olive oil was emulsified in a blender in distilled water containing 1% gum arabic at maximum speed for 2 min. Culture conditions were: 55 ◦ C, 150 rpm in a rotary shaker, in 1 L Erlenmeyer flasks containing 300 ml of medium. An aliquot of 3 ml of a 4 h preculture in LB medium (peptone 1% (w/v), yeast extract 0.5% (w/v), NaCl 0.5% (w/v), pH 6.5, 55 ◦ C), was used as inoculum.
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Cell growth was measured by total viable counts by the standard pour plate dilution method, and colonies on LB agar plates were counted after incubation at 55 ◦ C for 12 h. 2.5. Lipase purification Bacillus thermoleovorans CCR11 was grown until the stationary phase was reached (44 h) on liquid medium as described above. Cells from the cultures were removed by centrifugation (10,000 × g, 15 min, 25 ◦ C). The cell-free supernatant was then passed through a 0.45-m pore size membrane filter (Millipore) at 25 ◦ C, and the filtrate was diafiltered (with distilled water) and concentrated through a polyethersulfone ultrafiltration membrane, cut off 500,000 (Millipore) using a 200 ml stirred cell (Amicon). After addition of 0.16% Triton X-100 and 3 ml of Bio-Lyte ampholites (pH range 3–10, Bio Rad Labs.), 50 ml of the enzyme solution was loaded into a Rotofor® preparative electro focusing cell (Bio Rad Labs.). Proteins were focused at 15 W constant power for 4 h at 4 ◦ C. Twenty fractions of 2.5 ml each were collected and analyzed for pH, lipase activity, and by SDS-PAGE. Fractions showing lipase activity were pooled, concentrated by ultrafiltration through a polyethersulfone ultrafiltration membrane, cut off 10,000 (Millipore), and then subjected to non-denaturing polyacrylamide gel electrophoresis (16 cm × 16 cm × 3 mm). The band corresponding to the lipase, identified by zymography (see below), was cut off and electroeluted using the Mini Whole Gel Eluter unit (Bio Rad Labs.), at 140 V for 20 min. Electroeluted lipase was used for polypeptide relative molecular mass (Mr ) determination on SDS-PAGE. 2.6. Electrophoresis Polyacrylamide gel electrophoresis (PAGE) under nondenaturing conditions and SDS-PAGE were carried out as described by Laemmli (1970) [11], in a Protean II xi Electrophoresis Cell (Bio Rad Labs.). For non-denaturing PAGE, a 10% separating gel and a 4% stacking gel were used. For SDS-PAGE, a 15% separating gel and a 4% stacking gel were used. Proteins were stained by the silver stain method (Bio Rad Labs.), and Mr was estimated by comparison with broad range molecular weight standards (10–250 kDa) (Bio Rad Labs.). 2.7. Zymography In situ lipase activity was detected by zymography following non-denaturing PAGE. After electrophoresis, the gel was placed onto a 2% agar plate containing 3% olive oil and 0.001% Rhodamine B. After incubation for 12 h at 55 ◦ C, lipase activity was visualized as a fluorescent band under 350 nm UV light.
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2.8. Lipase characterization Lipase characterization was performed using the enzyme obtained after preparative electrofocusing, since recovery after electroelution was low. 2.8.1. Effect of pH and temperature on lipase activity and stability The optimal temperature for activity was determined by the spectrophotometric assay using pNP-laurate as substrate at different temperatures (30–70 ◦ C), at pH 6.5. For determination of temperature stability, 0.05 ml of the enzyme solution were incubated for 1 h in 50 mM phosphate buffer pH 6.5, at 30, 40, 50, 60 and 70 ◦ C. Residual activity was measured by the spectrophotometric assay described above. Optimal pH was determined by the spectrophotometric assay using pNP-laurate as substrate, at 60 ◦ C in buffer solutions of pH values ranging from 6 to 11 (0.05 M sodium acetate pH 5 and 6; 0.05 M potasium phosphate pH 6.5 and 7; 0.05 M Tris–HCl pH 8; 0.05 M CHES pH 9, 9.5 and 10; 0.05 M Borax–NaOH pH 11). The effect of pH on enzyme stability was analyzed by the spectrophotometric assay after pre-incubation of 0.2 ml of enzyme solution for 26 h at 30 ◦ C, in 0.5 ml of the above mentioned buffer solutions (pH 5–11). 2.8.2. Effect of detergents on lipase The effect of various detergents on enzyme activity was analyzed by incubating an enzyme aliquot (0.05 ml) for 1 h at 30 ◦ C in 0.05 M phosphate buffer (pH 6.5), containing 1% (v/v) of the detergent (Triton X-100, Tween 80, Tween 20 and SDS (Sigma)). Activity was measured by the spectrophotometric assay after incubation time. 2.8.3. Effect of organic solvents and diverse chemicals on lipase The effect of organic solvents on enzyme activity was determined by the spectrophotometric assay after preincubation of enzyme (0.2 ml) for 1 and 2 h at 30 ◦ C, in 0.5 ml of acetone, methanol, ethanol, 2-propanol, or butanol. Residual activity was measured by the spectrophotometric assay using 0.05 ml of the incubation mixture. The effect of diverse chemicals on activity was determined by the spectrophotometric assay, after pre-incubation with 1 mM of BaCl2 , CaCl2 , MgCl2 , KCl, LiCl, HgCl2 , EDTA, -mercaptoetanol, and phenylmethyl-sulfonyl fluoride (PMSF) in 0.05 M phosphate buffer (pH 6.5), at 30 ◦ C for 1 h.
instructions. Bovine serum albumin (Sigma) at a concentration of 10–100 g was used as standard.
3. Results and discussion 3.1. Strain selection Four of the eleven thermophilic Bacillus strains that were screened showed an orange fluorescent halo when observed under 350 UV light, indicating the production of an extracellular lipase. Bacillus thermoleovorans CCR11 was chosen for lipase characterization since it produced the highest lipase activity (5500 U/mg protein). Bacillus thermoleovorans CCR11 is an aerobic, non motile and spore-forming rod, able to grow at high temperatures (40–70 ◦ C), with an optimum at 55 ◦ C, and an optimum pH of 6.5. The full 16S rDNA sequencing of Bacillus thermoleovorans CCR11 showed the highest homology (99%) with Bacillus thermoleovorans and Bacillus kaustophilus. 3.2. Lipase production Typical growth and lipase production curves of Bacillus thermoleovorans CCR11 are shown in Fig. 1. The highest lipase production was found at the begining of the stationary phase, after 44 h of growth in a medium containing nutrient broth 0.325% (w/v), CaCl2 0.1% (w/v), olive oil 2.5%, and gum arabic 1%, pH 6.5 [3]. A decrease in lipase activity was observed during the late stationary phase, probably due to the presence of proteases in the culture medium. 3.3. Enzyme purification After 44 h of culture, cell-free supernatant was prepared by centrifugation (10,000 × g, 15 min, 4 ◦ C) of culture broth and filtration through a 0.45 m membrane. Attempts to pre-
2.8.4. Substrate specificity Lipase subtrate specificity was analyzed by the spectrophotometric assay, using 0.01 M p-nitrophenyl esters (C2, C3, C4, C6, C10, C12, C16, C18) dissolved in ethanol as substrates. 2.9. Protein measurement The protein concentration was determined with a Bio-Rad Protein Assay Kit (Lowry) according to the manufacturer’s
Fig. 1. Growth curve () and lipolytic activity () of B. thermoleovorans CCR11. Culture conditions are described in the text. Lipolytic activity was measured in the growth medium at intervals using p-nitrophenyl laurate as substrate.
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Fig. 2. Silver stained SDS-PAGE gel of Rotofor® fractions. Lane numbers correspond to Rotofor® fraction numbers. Std: molecular weight standards 10–150 kDa (Bio Rad).
cipitate protein by ammonium sulfate failed, since lipase activity was associated with protein aggregates that floated in the salt solutions, as has been previously reported [12]. Aggregation of lipase was confirmed by Sephadex G-75 gel filtration chromatography, since the enzyme eluted in the void volume, indicating the presence of protein aggregates with a molecular mass greater than 80 kDa (data not shown). Aggregation of microbial extracellular lipases has been previously reported [2,3,7,13–15]. Taking advantage of the protein aggregates formation, crude extract was diafiltered and concentrated two-fold through a polyethersulfone ultrafiltration membrane of 500,000 Da cut off. This resulted in a six-fold increase in specific activity in comparison to crude enzyme preparation (34,000 U/mg protein), with a 100% recovery. In order to avoid lipase aggregation in the following purification steps, 0.16% Triton X-100 was added to the enzyme preparation. Lipase was isoelectrofocused on a preparative isoelectrofocusing cell, resulting in eight fractions with lipolytic activity corresponding to pH values from 5 to 6.7. Fractions 8–12 showed the highest lipolytic activity and purity since on SDS-PAGE (Fig. 2) only two stained bands were observed, one corresponding to the lipase. This evidence suggests that preparative isoelectrofocusing is an effective method for the isolation and near purification of Bacillus thermoleovorans CCR11 extracellular lipase. Active fractions were pooled, concentrated by ultrafiltration with a 10,000 Da cut off polyethersulfone membrane, and used for enzyme characterization. The final recovery of the procedure was 27%. 3.4. Lipase characterization 3.4.1. Relative molecular mass (Mr ) Electrofocusing fractions showing lipase activity were pooled, concentrated and subjected to non-denaturing gel electrophoresis. Lipase location was determined by zymography (Fig. 3A). The band corresponding to the lipase was cut out and electro-eluted. The lipase was confirmed to be homogeneous by a single band on SDS-PAGE with a rela-
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Fig. 3. (A) Zymography of purified lipase from B. thermoleovorans CCR11 after non-denaturing PAGE, under 350 nm UV light. (B) SDS-PAGE of purified lipase. Std: molecular weight standards 10–150 kDa (Bio Rad).
tive molecular mass of 11 kDa (Fig. 3B). Reported molecular weights of microbial lipases are variable, ranging from 12 kDa [16] to 76 kDa [17]. In particular, molecular sizes of lipases from thermophilic bacteria range from 16 kDa in Bacillus thermocatenulatus [3], to 69 kDa in Bacillus sp THL027 [7]. The lipase produced by Bacillus thermoleovorans CCR11 in this study is the smallest published lipase, with a Mr close to that reported for the lipase produced by Bacillus thermoleovorans ID-1 (18 kDa) [18]. The lipase isoelectric point was around 6, as determined by preparative isoelectrofocusing. Isoelectric points of lipases produced by Bacillus strains are variable to some extent. Lipase from Bacillus subtilis 168, Bacillus sp A30-1, Bacillus stearothermophilus, and Bacillus sp H-257 have isoelectric points of 9.9, 5.15, 7.4 and 4.66, respectively [2,9,15,19]. 3.4.2. Effect of pH and temperature on lipase activity and stability Bacillus thermoleovorans CCR11 lipase was found to be most active at a pH between 9 and 10 (Fig. 4), similar to that reported for lipases from B. thermoleovorans ID1 (pH 9) [18] and Bacillus A30-1 (pH 9.5) [19], and higher to other reported lipases from thermophilic Bacillus which lie in the range of pH 7.2–8.5 [3–7,9,12]. Our lipase was stable in a broad range of pH values (5–11), retaining more than 80% of activity after 26 h at 30 ◦ C (Fig. 4). The optimum catalytic temperature for Bacillus thermoleovorans CCR11 lipase was 60 ◦ C (Fig. 5), similar to that reported for lipases produced by Bacillus sp J33 (60 ◦ C) [5], Bacillus sp A30-1 (60 ◦ C) [19], and Bacillus thermoleovorans ID-1 (60–65 ◦ C) [18]. Stability was tested after incubation at different temperatures for 1 h at pH 6.5. The enzyme was relatively thermostable, since it retained almost 100% of its activity at 30–40 ◦ C, 75% at 50–60 ◦ C, while at 70 ◦ C the enzyme lost its activity completely (Fig. 5). The stability of the enzyme in acidic and alkaline pHs and at high temperatures suggests its usefulness in industrial applications.
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Table 2 Effect of detergents and enzyme inhibitors on B. thermoleovorans CCR11 lipase activity
Fig. 4. Effect of pH on enzyme activity and stability (dashed line). Optimal pH was determined at 60 ◦ C and various pH buffers (6–11). Stability was analyzed after preincubating the enzyme preparation for 26 h at 30 ◦ C in buffer solutions at various pHs (5–11). Results are presented as percentage of the initial activity. 0.05 M sodium acetate (pH 5, 6); 0.05 M potasium phosphate (pH 6.5, 7); 0.05 M Tris–HCl (pH 8); 0.05 M CHES (pH 9, 9.5 and 10); 0.05 M Borax–NaOH (pH 11).
Fig. 5. Effect of temperature on enzyme activity and stability (dashed line). For temperature stability the enzyme preparation was preincubated at different temperatures for 1 h at pH 6.5 and the remaining activity was measured under standard conditions.
3.4.3. Effect of metal ions and inhibitors on lipase The effect of different metal ions on the activity of the lipase is shown in Table 1. Lipase activity was not affected by the presence of Na+ and Ba+ salts, whereas Mg2+ , K+ , and Li+ salts decreased activity by 25, 24, and 23%, respectively, Table 1 Effect of metal ions on B. thermoleovorans CCR11 lipase activity Metal ions (1 mM)
Relative activity (0 h)
Control BaCl2 CaCl2 MgCl2 KCl LiCl NaCl HgCl
100.0 94.9 159.2 93.6 100.4 94.0 98.3 24.9
± ± ± ± ± ± ± ±
4.57 2.85 2.82a 0.65 1.21 1.13 8.21 1.07a
Relative activity (1 h) 100.0 100.1 134.9 75.3 76.8 77.1 96.4 11.3
± ± ± ± ± ± ± ±
4.19 4.19 5.75a 2.44a 1.94a 0.82a 3.21 2.83a
Lipase preparation was incubated in the presence of each metal ion at 30 ◦ C for 1 h. Values represent the mean of three replicates. a p < 0.001 with respect to control.
Compound (1 mM)
Relative activity%
Control EDTA PMSF Triton X-100 Tween 20 Tween 80 SDS -Mercaptoethanol
100.0 ± 8.8 61.7 ± 1.4a 17.3 ± 0.5a 108.7 ± 1.96 0 0 0 95.3 ± 8.8
Lipase preparation was incubated in the presence of each compound at 30 ◦ C for 1 h. Values represent the mean of three replicates. a p < 0.001 with respect to control.
after 1 h of incubation at 30 ◦ C. It has been suggested that the effect of metal ions could be attributed to a change in the solubility and the behavior of the ionized fatty acids at interfaces, and from a change in the catalytic properties of the enzyme itself [15]. Hg2+ had a strong inhibitory effect on lipase activity suggesting it is able to alter enzyme conformation. As has been reported for other thermophilic lipases [6,12,18], Ca2+ salts increased activity immediately (59%) and after 1 h of incubation at 30 ◦ C (35%). The calcium-induced increase on lipase activity could be attributed to the complex action of calcium ions on the released fatty acids, and on enzyme structure stabilization due to the binding of calcium ions to the lipase, bridging the active region to a second subdomain of the protein and hence stabilizing enzyme tertiary structure [2]. Lipase activity was strongly diminished (83%) when the extract was incubated in the presence of 1 mM phenylmethylsulfonyl fluoride (PMSF) at 30 ◦ C for 1 h, which suggests the presence of a serine residue at the catalytic triad of the active site (Table 2). 3.4.4. Effect of detergents and organic solvents on lipase activity The effects of detergents and organic solvents on lipase activity are depicted in Tables 2 and 3. Addition of 1% Triton X-100 to the lipase mixture increased slightly the enzyme activity (108% with respect to the control), after 30 min at 60 ◦ C, pH 6.5; but it was completely inhibited in the presence of 1% SDS, Tween 80 or Tween 20. The response to the presence of detergents by reported thermophilic lipases is variable to some extent. In accordance to our results, SchmidtDannert et al. [3] reported a total loss of lipolytic activity in the presence of Tween 20 and Tween 80, but no effect was observed when incubated with Triton X-100. Nawani et al. [5] also found a total loss of activity in the presence of SDS but in contrast, activity was enhanced in the presence of Triton X-100, Tween 20 and Tween 80. Lipases are diverse in their sensitivity to solvents, but there is general agreement that polar water miscible solvents are more destabilizing than water immiscible solvents [5]. In this study, the lipase from B. thermoleovorans CCR11 showed
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Table 3 Stability of B. thermoleovorans CCR11 lipase in organic solvents Organic solvent
Relative activity% (0 h)
Control Acetone Methanol Ethanol 2-Propanol Butanol
100 89.2 121.1 102.0 89.2 0
± ± ± ± ±
3.7 1.3a 1.2a 3.2 4.7a
Relative activity% (1 h) 100 97.7 92.4 98.6 101.3 0
± ± ± ± ±
6.1 13.2 9.7 1.9 11.4
Relative activity% (2 h) 100 39.6 65.7 71.8 82.8 0
± ± ± ± ±
1.6 2.1a 2.6a 4.3a 2.5a
Lipase preparation was incubated in each organic solvent (70%) at 30 ◦ C for 1 and 2 h. Values represent the mean of three replicates. a p < 0.001 with respect to control.
high stability in the presence of water miscible organic solvents, since it retained almost 100% activity after exposure by 1 h at 30 ◦ C in 70% methanol, 70% ethanol, 70% 2-propanol and 70% acetone (Table 3). Addition of 70% methanol to the lipase mixture caused a 21% immediate increase in the lipolytic activity in comparison to the control. Similar results have been obtained for BTL-1 and BTL-2 lipases [3,20], which increased activity in the presence of 30% methanol, 30% ethanol and 30% acetone (BTL-1) and lost only 20% activity after 1 h of incubation at 30 ◦ C (BTL-2). It has been proposed that a thin layer of water molecules remains tightly bound to the enzyme acting as a protective sheath along the enzyme’s hydrophilic surfaces and allowing retention of the native conformation [5]. On the contrary, and as reported for B. stearothermophilus (Tok19A) esterase [21], 70% butanol caused an immediate and complete loss of lipolytic activity, probably because it provoked a rapid protein denaturation. Stability of this lipase in organic solvents suggests possible applications in non-aqueous biocatalysis. 3.4.5. Substrate specificity The enzyme specificity was studied with p-nitrophenyl esters of different alkyl chain lenghts (Fig. 6). The highest hydrolytic activity was obtained with C6-C10 p-nitrophenyl esters, with the highest activity toward p-nitrophenyl caproate (C10), indicating a clear preference of the enzyme for
medium acyl chain lenghts. A similar specificity has been reported for BTL-2 lipase [20], although other reported thermophilic lipases have shown preference for esters with shorter (C4 and C6) fatty acids when assayed against pnitrophenyl derivatives [4,6,9,12]. The lipase produced by B. thermoleovorans CCR11 showed similarities with lipases produced by B. thermoleovorans ID-1 [18] and Bacillus thermocatenulatus [3]. B. thermoleovorans species are characterized by its phenotypic heterogeneity and genotypic homogeneity [22]. Both belong to Group 5 of Bacillus species which comprises thermophilic strains (B. stearothermophilus, B. thermocatenulatus, B. thermoleovorans, B. kaustophilus, B. thermoglucosidasius, B. thermodenitrificans, B. caldolyticus, B. caldotenax, B. caldovelox, B. thermoantarticus, Saccharococcus thermophilus), displaying very high homology among their 16S rRNA sequences (98.5–99.2%) [23]. In particular, it has been suggested that B. thermoleovorans, B. kaustophilus and B. thermocatenulatus should be combined into one species (B. thermoleovorans) [22]. The similarities and differences found in the characteristics of the lipases produced by these species remarks the phenotypic heterogeneity found in this taxonomic group. In conclusion, we purified a lipase produced by the thermophilic strain Bacillus thermoleovorans CCR11 isolated from a hot spring in Mexico. The results obtained after characterization of the enzyme indicated that the lipase produced by this strain is an alkaline and thermophilic lipase, with one of the lowest molecular weight reported in the literature. Its stability in the presence of organic solvents, as well as in acidic and alkaline pHs and its tollerance to high temperatures makes it a good candidate for its application in non-aqueous biocatalysis. Acknowledgements
Fig. 6. Relative activities of the lipase on various p-NP esters. Lipase activities are expressed as the percentage of that of p-nitrophenyl caprate (C10), which was taken as 100%.
Lelie D. Castro (M.Sc.) and Citlali Rodr´ıguez (M.Sc.) acknowledge their scholarships from the mexican National Council for Science and Technology (Conacyt). This work was supported by Grant 431.01-P from the National Council on Technological Education (Cosnet). The advise and discussion of Dr. Hugo S. Garc´ıa Galindo and Ing. Oscar Calahorra Fuertes is gratefully acknowledged.
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[14] Ruiz B, Farres A, Langley E, Masso F, Sanchez S. Purification and characterization of an extracellular lipase from Penicillium candidum. Lipids 2001;36:283–9. [15] Lesuisse E, Schanck K, Colson C. Purification and preliminary characterization of the extracellular lipase of Bacillus subtilis 168, an extremely basic pH-tolerant enzyme. Eur J Biochem 1993;216:155–60. [16] Mase T, Matsumiya Y, Akiba T. Purification and characterization of a new lipase from Fusarium sp YM-30. Biosci Biotech Biochem 1995;59:1771–2. [17] Jurgens D, Huser H, Brunner H, Fehrenbach FJ. Purification and characterization of Staphilococcus aureus lipase. FEMS Microbiol Lett 1981;12:195–9. [18] Lee DW, Kim HK, Lee KW, Kim BC, Choe AC, Lee HS, et al. Purification and characterization of two distinct thermostable lipases from the gram-positive thermophilic bacterium Bacillus thermoleovorans ID-1. Enz Microb Technol 2001;29:363–71. [19] Wang Y, Srivastava KC, Shen GJ, Wang HY. Thermostable alkaline lipase from a newly isolated thermophilic Bacillus, strain A30-1 (ATCC 53841). J Ferm Bioeng 1995;79:433–8. [20] Schmidt-Dannert C, Rua ML, Atomi H, Schmid RD. Thermoalkalophilic lipase of Bacillus thermocatenulatus. I. Molecular cloning, nucleotide sequence, purification and some properties. Biochim Biophys Acta 1996;1301:105–14. [21] Wood ANP, Fernandez-Lafuente R, Cowan DA. Characterization of a novel thermostable esterase as an industrial catalyst. Biotechnol Appl Biochem 1995;21:313–22. [22] Sunna A, Tokajian S, Burghsrdt J, Rainey F, Antranikian G, Hashwa F. Identification of Bacillus kaustophilus, Bacillus thermocatenulatus and Bacillus strain HSR as members of Bacillus thermolerovorans. Syst Appl Microbiol 1997;20:232–7. [23] Nazina TN, Tourova TP, Poltaraus AB, Novikova EV, Grigoryam AA, Ivanova AE, et al. Taxonomic study of aerobic thermophilic bacilli: descriptions of Geobacillus subterraneus gen. nov., sp. nov., and Geobacillus uzenensis sp. nov. from petroleum reservoirs and transfer of Bacillus stearothermophilus, Bacillus thermocatenulatus, Bacillus thermoleovorans, Bacillus kaustophilus, Bacillus thermoglucosidasius and Bacillus thermodenitrificans to Geobacillus as the new combionations G. stearothermophilus, G. thermocatenulatus, G. thermoleovorans, G. kaustophilus, G. thermoglucosidasius and G. termodenitrificans. Int J Syst Evol Microbiol 2001;51:433–46.
Screening, purification and characterization of the thermoalkalophilic lipase produced by Bacillus thermoleovorans CCR11 Lelie D. Castro-Ochoa, Citlali Rodr´ıguez-G´omez, Gerardo Valerio-Alfaro, Rosamar´ıa Oliart Ros ∗ Unidad de Investigaci´on y Desarrollo en Alimentos, Instituto Tecnol´ogico de Veracruz. Av. Miguel A. de Quevedo 2779, Veracruz, Ver, 91897 Veracruz, M´exico Accepted 20 June 2005
Abstract Among eleven aerobic thermophilic Bacillus strains, isolated from “El Carrizal” hot springs in Veracruz, Mexico, Bacillus thermoleovorans CCR11 (EMBL # AJ536599) was selected for lipase characterization because of its high lipase specific activity. Lipase was purified by diafiltration (polyethersulfone ultrafiltration membrane co500,000), and preparative isoelectrofocusing. The lipase had a relative molecular mass of 11 kDa (the lowest Mr reported), although it formed higher molecular weight aggregates in native form. The optimum catalytic conditions for Bacillus thermoleovorans CCR11 lipase were 60 ◦ C and pH 9–10. Hg2+ , PMSF, SDS, Tween 80 and Tween 20 had an inhibitory effect on lipase activity, whereas Ca2+ salts and Triton X-100 increased it. Lipase activity was compatible with the presence of organic solvents, except for butanol. Lipase showed a notable preference for C6–C10 p-nitrophenyl esters, with the highest activity toward p-nitrophenyl caproate (C10). Lipase stability in the presence of organic solvents, as well as in acidic and alkaline pHs and at high temperatures makes it a good candidate for its application in non-aqueous biocatalysis. © 2005 Elsevier Inc. All rights reserved. Keywords: Lipase; Bacillus thermoleovorans; Thermophile; Enzyme characterization
1. Introduction Lipases (triacylglycerol acylhydrolases; EC 3.1.1.3) constitute a group of enzymes defined as carboxylesterases that catalyze the hydrolysis (and synthesis) of long chain acylglycerols at the lipid-water interface. Microbial lipases have been widely used for biotechnological applications in detergents, dairy and textile industries, production of surfactants, and oil processing. In recent years, lipases have received considerable attention with regard to the preparation of enantiomerically pure pharmaceuticals, since they have a number of unique characteristics: substrate specificity, regio-specificity, and chiral selectivity [1]. Given that such reactions are sometimes performed most
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0141-0229/$ – see front matter © 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.enzmictec.2005.06.003
efficiently at elevated temperatures and in organic solvents, converging attempts have been made to find thermostable lipases which would have advantages over labile enzymes in such applications [2]. In recent years, a number of thermophilic microorganisms producing thermoactive lipases and esterases have been purified and characterized [3–9]. Thermophilic lipases show higher thermostability, higher activity at elevated temperatures, and often show more resistance to chemical denaturation, making them ideal tools in industrial and chemical procesess where relatively high reaction temperatures and/or organic solvents are used. As each industrial application may require specific properties of the biocatalysts, there is still an interest in finding new lipases that could create novel applications. In this study, we describe the purification and characterization of a thermoalkaliphilic lipase produced by the thermophilic strain Bacillus thermoleovorans CCR11, isolated from a hot spring in Veracruz, Mexico.
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2. Materials and methods 2.1. Bacterial strains Eleven aerobic thermophilic Bacillus strains, isolated from “El Carrizal” hot springs in Veracruz, Mexico, were analyzed for lipolytic activity. Bacillus thermoleovorans CCR11 (EMBL # AJ536599) was selected for lipase production and characterization because it showed the highest lipase specific activity. 2.2. Screening All eleven thermophilic Bacillus strains were screened for lipase activity on agar plates containing Rhodamine B 0.001% (w/v), nutrient broth 0.8% (w/v), NaCl 0.4% (w/v), agar 1% (w/v), and olive oil 3%, in distilled water, pH 6.5 [10]. Plates were incubated at 55 ◦ C for 18 h, and lipase production was identified as an orange halo around colonies under UV light at 350 nm. In order to select the best lipase producer for enzyme purification and characterization, strains with lipolytic activity on the plates were cultured in liquid medium (nutrient broth 0.325% (w/v), CaCl2 0.1% (w/v), olive oil 2.5%, and gum arabic 1%, pH 6.5, 55 ◦ C, 150 rpm [3]), and lipase activity was determined spectrophotometrically with p-nitrophenyl-laurate as substrate (see below). 2.3. Assay for lipolytic activity Lipolytic activity was determined by a spectrophotometric assay using p-nitrophenil-laurate (pNPL) as substrate [5]. The reaction mixture consisted of: 0.1 ml enzyme extract, 0.8 ml 0.05 M phosphate buffer (pH 6.5), and 0.1 ml 0.01 M pNPL in ethanol. The hydrolytic reaction was carried out at 60 ◦ C for 30 min, after which 0.25 ml of 0.1 M Na2 CO3 was added. The mixture was centrifuged (16,000 × g, 15 min, 25 ◦ C) and the absorbance at 410 nm was determined. One unit of lipase activity was defined as the amount of enzyme that caused the release of 1 mol of p-nitrophenol (molar absorption coefficient 4.6 mM−1 cm−1 ) from pNP-laurate in 30 min under test conditions. 2.4. Lipase production Bacillus thermoleovorans CCR11 was grown in a liquid medium containing nutrient broth 0.325% (w/v), CaCl2 0.1% (w/v), olive oil 2.5%, and gum arabic 1%, pH 6.5 [3]. Olive oil was emulsified in a blender in distilled water containing 1% gum arabic at maximum speed for 2 min. Culture conditions were: 55 ◦ C, 150 rpm in a rotary shaker, in 1 L Erlenmeyer flasks containing 300 ml of medium. An aliquot of 3 ml of a 4 h preculture in LB medium (peptone 1% (w/v), yeast extract 0.5% (w/v), NaCl 0.5% (w/v), pH 6.5, 55 ◦ C), was used as inoculum.
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Cell growth was measured by total viable counts by the standard pour plate dilution method, and colonies on LB agar plates were counted after incubation at 55 ◦ C for 12 h. 2.5. Lipase purification Bacillus thermoleovorans CCR11 was grown until the stationary phase was reached (44 h) on liquid medium as described above. Cells from the cultures were removed by centrifugation (10,000 × g, 15 min, 25 ◦ C). The cell-free supernatant was then passed through a 0.45-m pore size membrane filter (Millipore) at 25 ◦ C, and the filtrate was diafiltered (with distilled water) and concentrated through a polyethersulfone ultrafiltration membrane, cut off 500,000 (Millipore) using a 200 ml stirred cell (Amicon). After addition of 0.16% Triton X-100 and 3 ml of Bio-Lyte ampholites (pH range 3–10, Bio Rad Labs.), 50 ml of the enzyme solution was loaded into a Rotofor® preparative electro focusing cell (Bio Rad Labs.). Proteins were focused at 15 W constant power for 4 h at 4 ◦ C. Twenty fractions of 2.5 ml each were collected and analyzed for pH, lipase activity, and by SDS-PAGE. Fractions showing lipase activity were pooled, concentrated by ultrafiltration through a polyethersulfone ultrafiltration membrane, cut off 10,000 (Millipore), and then subjected to non-denaturing polyacrylamide gel electrophoresis (16 cm × 16 cm × 3 mm). The band corresponding to the lipase, identified by zymography (see below), was cut off and electroeluted using the Mini Whole Gel Eluter unit (Bio Rad Labs.), at 140 V for 20 min. Electroeluted lipase was used for polypeptide relative molecular mass (Mr ) determination on SDS-PAGE. 2.6. Electrophoresis Polyacrylamide gel electrophoresis (PAGE) under nondenaturing conditions and SDS-PAGE were carried out as described by Laemmli (1970) [11], in a Protean II xi Electrophoresis Cell (Bio Rad Labs.). For non-denaturing PAGE, a 10% separating gel and a 4% stacking gel were used. For SDS-PAGE, a 15% separating gel and a 4% stacking gel were used. Proteins were stained by the silver stain method (Bio Rad Labs.), and Mr was estimated by comparison with broad range molecular weight standards (10–250 kDa) (Bio Rad Labs.). 2.7. Zymography In situ lipase activity was detected by zymography following non-denaturing PAGE. After electrophoresis, the gel was placed onto a 2% agar plate containing 3% olive oil and 0.001% Rhodamine B. After incubation for 12 h at 55 ◦ C, lipase activity was visualized as a fluorescent band under 350 nm UV light.
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2.8. Lipase characterization Lipase characterization was performed using the enzyme obtained after preparative electrofocusing, since recovery after electroelution was low. 2.8.1. Effect of pH and temperature on lipase activity and stability The optimal temperature for activity was determined by the spectrophotometric assay using pNP-laurate as substrate at different temperatures (30–70 ◦ C), at pH 6.5. For determination of temperature stability, 0.05 ml of the enzyme solution were incubated for 1 h in 50 mM phosphate buffer pH 6.5, at 30, 40, 50, 60 and 70 ◦ C. Residual activity was measured by the spectrophotometric assay described above. Optimal pH was determined by the spectrophotometric assay using pNP-laurate as substrate, at 60 ◦ C in buffer solutions of pH values ranging from 6 to 11 (0.05 M sodium acetate pH 5 and 6; 0.05 M potasium phosphate pH 6.5 and 7; 0.05 M Tris–HCl pH 8; 0.05 M CHES pH 9, 9.5 and 10; 0.05 M Borax–NaOH pH 11). The effect of pH on enzyme stability was analyzed by the spectrophotometric assay after pre-incubation of 0.2 ml of enzyme solution for 26 h at 30 ◦ C, in 0.5 ml of the above mentioned buffer solutions (pH 5–11). 2.8.2. Effect of detergents on lipase The effect of various detergents on enzyme activity was analyzed by incubating an enzyme aliquot (0.05 ml) for 1 h at 30 ◦ C in 0.05 M phosphate buffer (pH 6.5), containing 1% (v/v) of the detergent (Triton X-100, Tween 80, Tween 20 and SDS (Sigma)). Activity was measured by the spectrophotometric assay after incubation time. 2.8.3. Effect of organic solvents and diverse chemicals on lipase The effect of organic solvents on enzyme activity was determined by the spectrophotometric assay after preincubation of enzyme (0.2 ml) for 1 and 2 h at 30 ◦ C, in 0.5 ml of acetone, methanol, ethanol, 2-propanol, or butanol. Residual activity was measured by the spectrophotometric assay using 0.05 ml of the incubation mixture. The effect of diverse chemicals on activity was determined by the spectrophotometric assay, after pre-incubation with 1 mM of BaCl2 , CaCl2 , MgCl2 , KCl, LiCl, HgCl2 , EDTA, -mercaptoetanol, and phenylmethyl-sulfonyl fluoride (PMSF) in 0.05 M phosphate buffer (pH 6.5), at 30 ◦ C for 1 h.
instructions. Bovine serum albumin (Sigma) at a concentration of 10–100 g was used as standard.
3. Results and discussion 3.1. Strain selection Four of the eleven thermophilic Bacillus strains that were screened showed an orange fluorescent halo when observed under 350 UV light, indicating the production of an extracellular lipase. Bacillus thermoleovorans CCR11 was chosen for lipase characterization since it produced the highest lipase activity (5500 U/mg protein). Bacillus thermoleovorans CCR11 is an aerobic, non motile and spore-forming rod, able to grow at high temperatures (40–70 ◦ C), with an optimum at 55 ◦ C, and an optimum pH of 6.5. The full 16S rDNA sequencing of Bacillus thermoleovorans CCR11 showed the highest homology (99%) with Bacillus thermoleovorans and Bacillus kaustophilus. 3.2. Lipase production Typical growth and lipase production curves of Bacillus thermoleovorans CCR11 are shown in Fig. 1. The highest lipase production was found at the begining of the stationary phase, after 44 h of growth in a medium containing nutrient broth 0.325% (w/v), CaCl2 0.1% (w/v), olive oil 2.5%, and gum arabic 1%, pH 6.5 [3]. A decrease in lipase activity was observed during the late stationary phase, probably due to the presence of proteases in the culture medium. 3.3. Enzyme purification After 44 h of culture, cell-free supernatant was prepared by centrifugation (10,000 × g, 15 min, 4 ◦ C) of culture broth and filtration through a 0.45 m membrane. Attempts to pre-
2.8.4. Substrate specificity Lipase subtrate specificity was analyzed by the spectrophotometric assay, using 0.01 M p-nitrophenyl esters (C2, C3, C4, C6, C10, C12, C16, C18) dissolved in ethanol as substrates. 2.9. Protein measurement The protein concentration was determined with a Bio-Rad Protein Assay Kit (Lowry) according to the manufacturer’s
Fig. 1. Growth curve () and lipolytic activity () of B. thermoleovorans CCR11. Culture conditions are described in the text. Lipolytic activity was measured in the growth medium at intervals using p-nitrophenyl laurate as substrate.
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Fig. 2. Silver stained SDS-PAGE gel of Rotofor® fractions. Lane numbers correspond to Rotofor® fraction numbers. Std: molecular weight standards 10–150 kDa (Bio Rad).
cipitate protein by ammonium sulfate failed, since lipase activity was associated with protein aggregates that floated in the salt solutions, as has been previously reported [12]. Aggregation of lipase was confirmed by Sephadex G-75 gel filtration chromatography, since the enzyme eluted in the void volume, indicating the presence of protein aggregates with a molecular mass greater than 80 kDa (data not shown). Aggregation of microbial extracellular lipases has been previously reported [2,3,7,13–15]. Taking advantage of the protein aggregates formation, crude extract was diafiltered and concentrated two-fold through a polyethersulfone ultrafiltration membrane of 500,000 Da cut off. This resulted in a six-fold increase in specific activity in comparison to crude enzyme preparation (34,000 U/mg protein), with a 100% recovery. In order to avoid lipase aggregation in the following purification steps, 0.16% Triton X-100 was added to the enzyme preparation. Lipase was isoelectrofocused on a preparative isoelectrofocusing cell, resulting in eight fractions with lipolytic activity corresponding to pH values from 5 to 6.7. Fractions 8–12 showed the highest lipolytic activity and purity since on SDS-PAGE (Fig. 2) only two stained bands were observed, one corresponding to the lipase. This evidence suggests that preparative isoelectrofocusing is an effective method for the isolation and near purification of Bacillus thermoleovorans CCR11 extracellular lipase. Active fractions were pooled, concentrated by ultrafiltration with a 10,000 Da cut off polyethersulfone membrane, and used for enzyme characterization. The final recovery of the procedure was 27%. 3.4. Lipase characterization 3.4.1. Relative molecular mass (Mr ) Electrofocusing fractions showing lipase activity were pooled, concentrated and subjected to non-denaturing gel electrophoresis. Lipase location was determined by zymography (Fig. 3A). The band corresponding to the lipase was cut out and electro-eluted. The lipase was confirmed to be homogeneous by a single band on SDS-PAGE with a rela-
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Fig. 3. (A) Zymography of purified lipase from B. thermoleovorans CCR11 after non-denaturing PAGE, under 350 nm UV light. (B) SDS-PAGE of purified lipase. Std: molecular weight standards 10–150 kDa (Bio Rad).
tive molecular mass of 11 kDa (Fig. 3B). Reported molecular weights of microbial lipases are variable, ranging from 12 kDa [16] to 76 kDa [17]. In particular, molecular sizes of lipases from thermophilic bacteria range from 16 kDa in Bacillus thermocatenulatus [3], to 69 kDa in Bacillus sp THL027 [7]. The lipase produced by Bacillus thermoleovorans CCR11 in this study is the smallest published lipase, with a Mr close to that reported for the lipase produced by Bacillus thermoleovorans ID-1 (18 kDa) [18]. The lipase isoelectric point was around 6, as determined by preparative isoelectrofocusing. Isoelectric points of lipases produced by Bacillus strains are variable to some extent. Lipase from Bacillus subtilis 168, Bacillus sp A30-1, Bacillus stearothermophilus, and Bacillus sp H-257 have isoelectric points of 9.9, 5.15, 7.4 and 4.66, respectively [2,9,15,19]. 3.4.2. Effect of pH and temperature on lipase activity and stability Bacillus thermoleovorans CCR11 lipase was found to be most active at a pH between 9 and 10 (Fig. 4), similar to that reported for lipases from B. thermoleovorans ID1 (pH 9) [18] and Bacillus A30-1 (pH 9.5) [19], and higher to other reported lipases from thermophilic Bacillus which lie in the range of pH 7.2–8.5 [3–7,9,12]. Our lipase was stable in a broad range of pH values (5–11), retaining more than 80% of activity after 26 h at 30 ◦ C (Fig. 4). The optimum catalytic temperature for Bacillus thermoleovorans CCR11 lipase was 60 ◦ C (Fig. 5), similar to that reported for lipases produced by Bacillus sp J33 (60 ◦ C) [5], Bacillus sp A30-1 (60 ◦ C) [19], and Bacillus thermoleovorans ID-1 (60–65 ◦ C) [18]. Stability was tested after incubation at different temperatures for 1 h at pH 6.5. The enzyme was relatively thermostable, since it retained almost 100% of its activity at 30–40 ◦ C, 75% at 50–60 ◦ C, while at 70 ◦ C the enzyme lost its activity completely (Fig. 5). The stability of the enzyme in acidic and alkaline pHs and at high temperatures suggests its usefulness in industrial applications.
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Table 2 Effect of detergents and enzyme inhibitors on B. thermoleovorans CCR11 lipase activity
Fig. 4. Effect of pH on enzyme activity and stability (dashed line). Optimal pH was determined at 60 ◦ C and various pH buffers (6–11). Stability was analyzed after preincubating the enzyme preparation for 26 h at 30 ◦ C in buffer solutions at various pHs (5–11). Results are presented as percentage of the initial activity. 0.05 M sodium acetate (pH 5, 6); 0.05 M potasium phosphate (pH 6.5, 7); 0.05 M Tris–HCl (pH 8); 0.05 M CHES (pH 9, 9.5 and 10); 0.05 M Borax–NaOH (pH 11).
Fig. 5. Effect of temperature on enzyme activity and stability (dashed line). For temperature stability the enzyme preparation was preincubated at different temperatures for 1 h at pH 6.5 and the remaining activity was measured under standard conditions.
3.4.3. Effect of metal ions and inhibitors on lipase The effect of different metal ions on the activity of the lipase is shown in Table 1. Lipase activity was not affected by the presence of Na+ and Ba+ salts, whereas Mg2+ , K+ , and Li+ salts decreased activity by 25, 24, and 23%, respectively, Table 1 Effect of metal ions on B. thermoleovorans CCR11 lipase activity Metal ions (1 mM)
Relative activity (0 h)
Control BaCl2 CaCl2 MgCl2 KCl LiCl NaCl HgCl
100.0 94.9 159.2 93.6 100.4 94.0 98.3 24.9
± ± ± ± ± ± ± ±
4.57 2.85 2.82a 0.65 1.21 1.13 8.21 1.07a
Relative activity (1 h) 100.0 100.1 134.9 75.3 76.8 77.1 96.4 11.3
± ± ± ± ± ± ± ±
4.19 4.19 5.75a 2.44a 1.94a 0.82a 3.21 2.83a
Lipase preparation was incubated in the presence of each metal ion at 30 ◦ C for 1 h. Values represent the mean of three replicates. a p < 0.001 with respect to control.
Compound (1 mM)
Relative activity%
Control EDTA PMSF Triton X-100 Tween 20 Tween 80 SDS -Mercaptoethanol
100.0 ± 8.8 61.7 ± 1.4a 17.3 ± 0.5a 108.7 ± 1.96 0 0 0 95.3 ± 8.8
Lipase preparation was incubated in the presence of each compound at 30 ◦ C for 1 h. Values represent the mean of three replicates. a p < 0.001 with respect to control.
after 1 h of incubation at 30 ◦ C. It has been suggested that the effect of metal ions could be attributed to a change in the solubility and the behavior of the ionized fatty acids at interfaces, and from a change in the catalytic properties of the enzyme itself [15]. Hg2+ had a strong inhibitory effect on lipase activity suggesting it is able to alter enzyme conformation. As has been reported for other thermophilic lipases [6,12,18], Ca2+ salts increased activity immediately (59%) and after 1 h of incubation at 30 ◦ C (35%). The calcium-induced increase on lipase activity could be attributed to the complex action of calcium ions on the released fatty acids, and on enzyme structure stabilization due to the binding of calcium ions to the lipase, bridging the active region to a second subdomain of the protein and hence stabilizing enzyme tertiary structure [2]. Lipase activity was strongly diminished (83%) when the extract was incubated in the presence of 1 mM phenylmethylsulfonyl fluoride (PMSF) at 30 ◦ C for 1 h, which suggests the presence of a serine residue at the catalytic triad of the active site (Table 2). 3.4.4. Effect of detergents and organic solvents on lipase activity The effects of detergents and organic solvents on lipase activity are depicted in Tables 2 and 3. Addition of 1% Triton X-100 to the lipase mixture increased slightly the enzyme activity (108% with respect to the control), after 30 min at 60 ◦ C, pH 6.5; but it was completely inhibited in the presence of 1% SDS, Tween 80 or Tween 20. The response to the presence of detergents by reported thermophilic lipases is variable to some extent. In accordance to our results, SchmidtDannert et al. [3] reported a total loss of lipolytic activity in the presence of Tween 20 and Tween 80, but no effect was observed when incubated with Triton X-100. Nawani et al. [5] also found a total loss of activity in the presence of SDS but in contrast, activity was enhanced in the presence of Triton X-100, Tween 20 and Tween 80. Lipases are diverse in their sensitivity to solvents, but there is general agreement that polar water miscible solvents are more destabilizing than water immiscible solvents [5]. In this study, the lipase from B. thermoleovorans CCR11 showed
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Table 3 Stability of B. thermoleovorans CCR11 lipase in organic solvents Organic solvent
Relative activity% (0 h)
Control Acetone Methanol Ethanol 2-Propanol Butanol
100 89.2 121.1 102.0 89.2 0
± ± ± ± ±
3.7 1.3a 1.2a 3.2 4.7a
Relative activity% (1 h) 100 97.7 92.4 98.6 101.3 0
± ± ± ± ±
6.1 13.2 9.7 1.9 11.4
Relative activity% (2 h) 100 39.6 65.7 71.8 82.8 0
± ± ± ± ±
1.6 2.1a 2.6a 4.3a 2.5a
Lipase preparation was incubated in each organic solvent (70%) at 30 ◦ C for 1 and 2 h. Values represent the mean of three replicates. a p < 0.001 with respect to control.
high stability in the presence of water miscible organic solvents, since it retained almost 100% activity after exposure by 1 h at 30 ◦ C in 70% methanol, 70% ethanol, 70% 2-propanol and 70% acetone (Table 3). Addition of 70% methanol to the lipase mixture caused a 21% immediate increase in the lipolytic activity in comparison to the control. Similar results have been obtained for BTL-1 and BTL-2 lipases [3,20], which increased activity in the presence of 30% methanol, 30% ethanol and 30% acetone (BTL-1) and lost only 20% activity after 1 h of incubation at 30 ◦ C (BTL-2). It has been proposed that a thin layer of water molecules remains tightly bound to the enzyme acting as a protective sheath along the enzyme’s hydrophilic surfaces and allowing retention of the native conformation [5]. On the contrary, and as reported for B. stearothermophilus (Tok19A) esterase [21], 70% butanol caused an immediate and complete loss of lipolytic activity, probably because it provoked a rapid protein denaturation. Stability of this lipase in organic solvents suggests possible applications in non-aqueous biocatalysis. 3.4.5. Substrate specificity The enzyme specificity was studied with p-nitrophenyl esters of different alkyl chain lenghts (Fig. 6). The highest hydrolytic activity was obtained with C6-C10 p-nitrophenyl esters, with the highest activity toward p-nitrophenyl caproate (C10), indicating a clear preference of the enzyme for
medium acyl chain lenghts. A similar specificity has been reported for BTL-2 lipase [20], although other reported thermophilic lipases have shown preference for esters with shorter (C4 and C6) fatty acids when assayed against pnitrophenyl derivatives [4,6,9,12]. The lipase produced by B. thermoleovorans CCR11 showed similarities with lipases produced by B. thermoleovorans ID-1 [18] and Bacillus thermocatenulatus [3]. B. thermoleovorans species are characterized by its phenotypic heterogeneity and genotypic homogeneity [22]. Both belong to Group 5 of Bacillus species which comprises thermophilic strains (B. stearothermophilus, B. thermocatenulatus, B. thermoleovorans, B. kaustophilus, B. thermoglucosidasius, B. thermodenitrificans, B. caldolyticus, B. caldotenax, B. caldovelox, B. thermoantarticus, Saccharococcus thermophilus), displaying very high homology among their 16S rRNA sequences (98.5–99.2%) [23]. In particular, it has been suggested that B. thermoleovorans, B. kaustophilus and B. thermocatenulatus should be combined into one species (B. thermoleovorans) [22]. The similarities and differences found in the characteristics of the lipases produced by these species remarks the phenotypic heterogeneity found in this taxonomic group. In conclusion, we purified a lipase produced by the thermophilic strain Bacillus thermoleovorans CCR11 isolated from a hot spring in Mexico. The results obtained after characterization of the enzyme indicated that the lipase produced by this strain is an alkaline and thermophilic lipase, with one of the lowest molecular weight reported in the literature. Its stability in the presence of organic solvents, as well as in acidic and alkaline pHs and its tollerance to high temperatures makes it a good candidate for its application in non-aqueous biocatalysis. Acknowledgements
Fig. 6. Relative activities of the lipase on various p-NP esters. Lipase activities are expressed as the percentage of that of p-nitrophenyl caprate (C10), which was taken as 100%.
Lelie D. Castro (M.Sc.) and Citlali Rodr´ıguez (M.Sc.) acknowledge their scholarships from the mexican National Council for Science and Technology (Conacyt). This work was supported by Grant 431.01-P from the National Council on Technological Education (Cosnet). The advise and discussion of Dr. Hugo S. Garc´ıa Galindo and Ing. Oscar Calahorra Fuertes is gratefully acknowledged.
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