Purification and characterization of two distinct thermostable lipases from the gram-positive thermophilic bacterium Bacillus thermoleovorans ID-1

Enzyme and Microbial Technology 29 (2001) 363–371

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Purification and characterization of two distinct thermostable lipases from the gram-positive thermophilic bacterium Bacillus thermoleovorans ID-1 Dong-Woo Leea, Hack-Woo Kima, Keun-Wook Leeb, Byoung-Chan Kima, Eun-Ah Choea, Han-Seung Leea, Doo-Sik Kimb, Yu-Ryang Pyuna,* a

Department of Biotechnology and Bioproducts Research Center, College of Engineering, Yonsei University, Seoul 120 –749, South Korea b Department of Biochemistry, College of Science, Yonsei University, Seoul 120 –749, South Korea Received 1 January 2001; received in revised form 21 May 2001; accepted 7 June 2001

Abstract The thermophilic bacterium Bacillus thermoleovorans ID-1 can hydrolyze a variety of oils such as olive oil, soybean oil, palm oil, and lard as a carbon source (1.5%, v/v) after 72 h of culture at 50°C. In this study, we purified to homogeneity two distinct thermostable lipases, designated BTID-A (B. thermoleovorans ID-1 lipase A) and BTID-B (B. thermoleovorans ID-1 lipase B). BTID-A was purified 300-fold from a cell-free culture supernatant of B. thermoleovorans ID-1 grown in modified TYEM medium in the absence of a lipid substrate as an inducer. Purification of BTID-A was carried out by ammonium sulfate precipitation, DEAE-Sepharose CL6B, Superdex 200, Resource PHE, and Mono Q column chromatography. Previously, the gene encoding BTID-B of B. thermoleovorans ID-1 has been cloned, sequenced, and expressed in Escherichia coli. Recombinant BTID-B was purified 108-fold from a cell extract of E. coli by heat precipitation, DEAE-Sepharose CL6B, and Sephacryl S200 column chromatography. Molecular mass of BTID-A was approximately 18 kDa and its activity was maximum at 60 to 65°C. The pH optimum for BTID-A was 9.0. On the other hand, BTID-B was a larger protein with a molecular mass of 43 kDa, but showed the similar optima for its activity as BTID-A. The activity of BTID-A was inhibited by organic solvents such as EtOH, DMSO, and ␤-mercaptoethanol, and divalent ions including Cu2⫹, Hg2⫹, and Co2⫹. In contrast, BTID-B was slightly activated by Ca2⫹, Co2⫹, and Mn2⫹ ions and strongly resistant to organic solvents. Although both of the enzymes showed different substrate specificities, their maximal activities were found with tricaprylin (C8) as a substrate. The Km values of BTID-A and BTID-B for the hydrolysis of tricaprylin were 1.82 mM (Vmax, 12.8 ␮mol min⫺1 mg⫺1) and 6.24 mM (Vmax, 63.3 ␮mol min⫺1 mg⫺1), respectively. © 2001 Elsevier Science Inc. All rights reserved. Keywords: Lipase; Purification; B. thermoleovorans ID-1

1. Introduction Many different bacterial species produce lipases (EC 3.1.1.3) which hydrolyze esters of glycerol with preferably long-chain fatty acids [1]. They act at the interface between a hydrophobic lipid substrate and a hydrophilic aqueous medium [2]. Interests in bacterial lipases have increased due to their biotechnological applications where the enzymes are used not only for the hydrolysis of lipids but also for the synthesis of a variety of industrially valuable products [1–5]. Thermostable enzymes are particularly attractive for Corresponding author. Tel.: ⫹82–2-2123–2883; fax: ⫹82–2-312– 6821. E-mail address: [email protected] (Y.-R. Pyun). *

industrial applications because of their high activities at the elevated temperatures and stabilities in organic solvents [6,7]. Recently several extracellular lipases have been reported from the genus Bacillus such as B. subtilis [8], B. liqueniformis [9], B. catenulatus [10,11], and B. stearothermophilus [12]. Among them, two extracellular thermostable lipases have been cloned, purified, and characterized from B. stearothermophilus L1 [12,13] and B. thermocatenulatus [10,11]. Amino acid sequence analysis of two enzymes showed no similarity to any other bacterial or fungal lipase [14 –21]. Furthermore, lipases from Bacillus species contain alanine as the first residue, instead of first glycine, in a highly conserved pentapeptide (Gly-X-Ser-X-Gly) forming a ␤-⑀Ser-␣ motif. To date, a number of lipases from meso-

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philes have been extensively characterized. However, little study has been reported for the lipases from thermophiles partly due to the low level of enzyme production and the strong hydrophobic interaction with their substrates [11,12, 22]. Previously, we reported that the thermophilic bacterium B. thermoleovorans ID-1 with its rapid specific growth rate of 2.50 h⫺1 at 65°C was isolated from geothermal hot springs in Indonesia [23]. An inducible extracellular lipase with a molecular mass of 34 kDa was produced by this strain. Growth of the organism in media containing a variety of lipid substrates led us to believe that it might produce heterogeneous lipases in addition to the inducible lipase. Thus, we tried to find different enzymes from its culture supernatant but failed to do. For this reason, we have cloned and expressed another lipase gene (BTID-B) of which the deduced sequence contains 416 amino acids corresponding to a molecular mass of 46 kDa [24]. Moreover, it was found that this organism could produce extracellular thermostable lipases (BTID-A) in the absence of a lipid substrate as an inducer. In this study, we have purified a noninducible extracellular lipase (BTID-A) from B. thermoleovorans ID-1 and a recombinant one (BTID-B) expressed in E. coli. Physicochemical properties of the purified enzymes have been also characterized. 2. Materials and methods 2.1. Bacterial strains, plasmids, and growth conditions Batch cultures of B. thermoleovorans ID-1 were carried out in modified TYEM medium (pH 6.0) in the absence of olive oil in a 5-liter bioreactor (Bioengineering AG, Wald, Switzerland) at 50°C as previously described [23]. E. coli DH5␣ expressing BTID-B was grown in Luria-Bertani (LB) medium containing 50 ␮g of ampicillin per ml at 37°C. pUC19 was a plasmid used for cloning and expression of BTID-B in E. coli [24,25]. 2.2. DNA techniques and recombination The genomic DNA of B. thermoleovorans ID-1 was isolated and partially digested with BamH ⌱ as previously described [24]. DNA fragments of 1.5– 6 kb were inserted into the vector pUC19 digested with BamH ⌱. E. coli DH5␣ was transformed with the vector pUC19 harboring BTID-B gene according to Sambrook et al. [26]. Extraction of the DNA molecules from an agarose gel was done with a Geneclean ⌸ kit (BIO 101 Inc., Vista, CA). The expression of true lipase was determined by rhodamine B agar plate assay according to Kouker and Jaeger [27]. 2.3. Enzyme assay Lipase activity was measured spectrophotometrically at 430 nm by monitoring copper soaps formed with free fatty

acids [23]. The enzyme reaction was carried out with 100 mM of tricaprylin (C8) as a substrate emulsified with 0.2 mM of gum arabic (4.7 mg/ml) in 50 mM potassium phosphate buffer (pH 7.5) at 60°C. One unit of lipase was defined as the amount of enzyme releasing 1 ␮mol of free fatty acids per min under the assay condition. Total protein concentration was determined using the bicinchoninic acid protein assay reagent kit (Sigma Chemical Co., St. Louis, MO) with bovine serum albumin as a standard. 2.4. Enzyme purification BTID-A and BTID-B were purified from the culture supernatant of B. thermoleovorans ID-1 and the cell extract of E. coli DH5␣, respectively. All procedures were performed at 4°C. Prior to use, all buffers were filtered through a 0.22-␮m-pore-size filter. All columns and column media were obtained from Pharmacia unless otherwise stated. 2.5. Purification of BTID-A A cell free supernatant obtained by centrifugation (10,000 ⫻ g for 20 min) was fractionated with ammonium sulfate (80% saturation). The precipitate was collected, resuspended in 20 mM Tris-HCl buffer (pH 7.5), and dialyzed overnight against the same buffer at 4°C. The dialyzed sample (50 ml) was applied onto a DEAE-Sepharose CL6B column (2.5 ⫻ 10 cm) equilibrated with 20 mM Tris-HCl buffer (pH 8.0) at 4°C. After washing with the same buffer, absorbed proteins were eluted with a linear gradient of 0 to 1.0 M NaCl at 1 ml/min. Fractions with lipase activity were pooled and concentrated approximately 10-fold with 10,000-molecular-weight-cutoff (MWCO) membranes (Amicon, Bedford, MA). The pooled and concentrated sample was applied to a HiLoad 16/60 Superdex 200 column (Pharmacia LKB) equilibrated with 20 mM Tris-HCl buffer (pH 7.5) containing 0.15 M NaCl at 0.5 ml/min. Ammonium sulfate was added to the pooled active fractions from the previous step to bring the sample at a final concentration of 1.5 M. The concentrated fractions were applied to a Resource PHE column (1 ml; Pharmacia LKB) equilibrated with 1.5 M ammonium sulfate in 20 mM Tris-HCl buffer (pH 7.5). Absorbed proteins were eluted with a descending salt gradient (1.5 to 0 M) and washed with 0 M ammonium sulfate in 20 mM Tris-HCl buffer. Proteins containing lipase activity were then eluted with an additional Triton X-100 gradient (0 to 0.2%) in the same buffer. The pooled fractions containing lipase activity were dialyzed overnight against 20 mM Tris-HCl buffer (pH 7.5) to eliminate detergent and concentrated as described above. The concentrated fractions were applied to a Mono Q HR 5/5 column (Pharmacia LKB) equilibrated with the same buffer at 0.5 ml/min. Elution was carried out using a linear NaCl gradient (0 to 0.5 M) and fractions containing lipase activity were analyzed with SDS-PAGE. Fractions showing

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2.8. N-terminal sequence After SDS-PAGE (12.5%), purified proteins were transferred to a Sequi-Blot PVDF membrane (Bio-Rad, Hercules, CA) in 10 mM CAPS buffer (pH 11), 10% methanol using a Mini Trans-Blot electrophoretic transfer apparatus (Bio-Rad, Hercules, CA). Protein sequencing was performed at Tufts University, Analytical Core Facility, Boston, MA. 2.9. Effects of temperature and pH

Fig. 1. Production of lipase by Bacillus thermoleovorans ID-1 in a batch fermentation on 1.5% (v/v) olive oil in modified TYEM medium at 50°C: profiles of cell density (f), soluble lipase activity measured in cell-free supernatant (䡬) and triolein concentration (Œ).

a single band on the gel were pooled and stored at 4°C until use. 2.6. Purification of BTID-B E. coli DH5␣ cells expressing BTID-B were harvested by centrifugation and resuspended in one-tenth volume of 50 mM potassium phosphate buffer (pH 8.0). The cells were disrupted by sonication and the cell debris was removed by centrifugation at 30,000 ⫻ g for 20 min. The cell extract was heated at 80°C for 10 min, and the denatured proteins were removed by centrifugation at 10,000 ⫻ g for 30 min. The lipase activity remained in the clear supernatant. The heat-treated cell extract was applied to a DEAE-Sepharose CL6B column (2.5 ⫻ 15 cm) equilibrated with 20 mM Tris-HCl buffer (pH 8.0) at 4°C at 1.0 ml/min. Proteins were eluted with a linear NaCl gradient (0 to 1 M) at 1.0 ml/min. The eluent from the DEAE column was concentrated using filters of 10 kDa-MWCO (Amicon, Bedford, MA) and then loaded to a Sephacryl S200 column (1.5 ⫻ 130 cm) equilibrated with 20 mM Tris-HCl buffer (pH 7.5, 0.15 M NaCl) at 0.25 ml/min. The blue dextran (2,000 kDa) was used to measure void volume (115 ml). 2.7. Gel electrophoresis Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (PAGE) (SDS-PAGE) was carried out to estimate the molecular mass according to the method of Laemmli [28]. Proteins were visualized by staining with Coomassie brilliant blue R-250.

The temperature optima were evaluated for both enzymes. In order to investigate thermostability, each enzyme was incubated for 30 min at 40, 50, 60, 70, or 80°C, and its residual activity was measured. The pH optima for both enzymes were determined using the following buffers: 50 mM sodium acetate buffer (pH 4 to 6), 50 mM potassium phosphate buffer (pH 6 to 8), 50 mM Tris-HCl buffer (pH 7 to 9), and 50 mM glycine-NaOH buffer (pH 8 to 11). Enzyme reactions in each buffer were carried out at 60°C and their residual activities were measured periodically. In order to determine the pH stability for both enzymes, preincubation was performed in each buffer at room temperature for 15 min. The pH of each buffer was adjusted at 60°C, the enzyme reaction temperature. 2.10. Substate specificity The substrate specificities of the purified lipases toward different triacylglycerols were determined by a standard assay. Substrate solutions of 100 mM tributyrin (C4), tricaproin (C6), tricaprylin (C8), tricaprin (C10), trilaurin (C12), trimyristin (C14), tripalmitin (C16), and tristearin (C18) (all triacylglycerols were obtained from Sigma) were emulsified with 0.2 mM of gum arabic (mol. wt. of approx. 250,000) prior to use. 2.11. Effects of metal ions and other reagents In order to determine the effects of various metal ions and other reagents on lipase activity, the activity was assayed at 60°C after pre-incubation of the purified enzymes with each compound for 15 min at room temperature. 2.12. Analysis of hydrolysis products and positional specificity The residual concentration of triolein (OOO), 1,3-dipalmitoyl-2-oleoyl-glycerol (POP), 1,2-dilinoleoyl-3-oleoyl-rac-glycerol (LLO), and 1,3-dioleoyl-2-palmitoyl-glycerol (OPO) in the culture broth was analyzed by gas-chromatography. Samples were extracted with n-hexane (Sigma, HPLC grade) and filtered with 0.2 ␮m pore size syringe filter (Adventec MFS, Inc., Pleasanton. CA). Gas chromatographic analysis was performed on a Hewlett Packard 6890 instrument using

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Table 1 Degree of hydrolysis of various substrates by B. thermoleovorans ID-1 Substratea

Degree of hydrolysis (%)b

Olive oil (OOO) Soybean oil (LLO) Palm oil (POP) Lard (OPO)

100 100 100 83.1

a B. thermoleovorans ID-1 was batch-cultured in modified TYEM medium with various substrates as the sole carbon source (1.5%, v/v) at 50°C. b Each aliquot of culture broth in 72 h was taken to be quantified by GC analysis. Degradation of each oil was determined by measuring the amount of its residual main triglyceride. Data are the averages of values at least three experiments.

a SGE HT5 capillary column (Supelco, 25 m ⫻ 0.32 mm i.d., 0.1 ␮m film thickness) and a flame ionization detector. The carrier gas was pure hydrogen using a head pressure of 16.1 psi. The injector was set for a split ratio of 1:50. Operation conditions were 360°C for both injector and detector. Positional specificity was analyzed by thin-layer chromatography (TLC) according to the method described by Sztajer et al. [29]. After enzyme reaction for 30 min at 60°C using 100 mM triolein as a substrate, 10 ␮l aliquots were taken and analyzed on TLC aluminium sheets silica gel 60 F254 plates (E. Merck, Darmstadt, Germany). The mobile phase consisted of petroleum ether, diethyl ether, acetic acid (70:30:1, v/v). Spots were visualized by spraying with 50% H2SO4 solution. 2.13. Kinetic parameters The Michaelis-Menten kinetic parameters, Vmax and Km values for both enzymes were calculated using tricaprylin (C8) as a substrate. Lineweaver-Burk plots were used to determine parameters, assuming that simple MichaelisMenten kinetics was followed.

3. Results and discussion 3.1. Lipase production and lipid hydrolysis The Gram-positive thermophilic bacterium B. thermoleovorans ID-1 produced extracellular lipases in modified

TYEM medium with olive oil (1.5%, v/v) as the sole carbon source at 50°C. Lipases were produced simultaneously to microbial growth and the maximal enzyme production (116 U/liter) took place during the late exponential phase (Fig. 1). The extracellular lipase activity decreased with prolonged cultivation possibly due to thermal inactivation and proteolysis. After 2 days of culture, olive oil in a culture broth was completely hydrolyzed. A variety of triglycerides such as soybean oil, palm oil, and lard as a carbon source were also done after 72 h culture at 50°C (Table 1). Since an oil contains heterogeneous triglycerides, it was difficult to directly measure the amount of substrates. Instead, degree of hydrolysis was quantitatively measured for olive oil using triolein, a major triglyceride in olive oil (40 – 45%). For other substrates, POP, LLO, and OPO were used for measuring the degree of hydrolysis. B. thermoleovorans ID-1 hydrolyzed olive oil, soybean oil, and palm oil completely while lard was hydrolyzed by 83.1%. These results suggest that B. thermoleovorans ID-1 may be useful not only as a proper bacterial strain for treatment of lipid-rich effluents but also as a lipase producer. 3.2. Purification of lipases Purification of the extracellular lipase, BTID-A, is summarized in Table 2. Salt precipitation with 80% ammonium sulfate resulted in 27% yield, relatively low recovery. To minimize the loss from the salt precipitation, the cell-free culture supernatant was directly applied onto DEAE-Sepharose column, but the resolution was poor (data not shown). So, the salt precipitate was loaded to a DEAE-Sepharose CL6B anion exchange column chromatography. In this anion-exchange step, two lipase activities were eluted at 200 mM and 700 mM NaCl (data not shown). The major active fractions eluted at 200 mM were pooled, concentrated, and applied to subsequent Superdex 200 gel column. The minor fraction could not be further purified because of the limited amount. On gel chromatography, the major peak was eluted in the void volume. This suggested that the enzyme might be hydrophobic in nature. It was consistent with the observation that in Resource PHE hydrophobic chromatography (FPLC), the lipases were eluted in the linear gradient of 0 to 0.2% triton X-100. Using Mono Q anion exchange chromatography, the major lipase (BTID-A) was purified 300-fold

Table 2 Purification of BTID-A from B. thermoleovorans ID-1 Purification step

Total protein (mg)

Total activity (U)

Sp act (U/mg)

Purification Fold

Yield (%)

Culture supernatant Ammonium sulfate precipitation DEAE CL6B Superdex 200 Resource PHE Mono Q

1,724 52.0 8.9 1.1 0.2 0.01

74.7 20.2 1.5 0.7 0.5 0.12

0.04 0.39 0.17 0.64 2.5 12

1 9.8 4.3 17.5 62.5 300

100 27 2 0.94 0.67 0.16

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Table 3 Purification of recombinant BTID-B from E. coli DH5 ␣ Purification step

Total protein (mg)

Total activity (U)

Sp act (U/mg)

Purification Fold

Yield (%)

Cell extract Heat treatment DEAE CL6B Sephacryl S200

837 65.4 2.9 0.25

2275 583.5 90.7 73.1

2.72 8.92 31.3 292.4

1 3.28 11.5 107.5

100 25.7 4 3.2

to homogeneity with a specific activity of 12 U/mg and a yield of 0.16%. Purification of BTID-B from a culture supernatant of B. thermoleovorans ID-1 was not successful because of the limited amount of the enzyme. In order to increase the amount of BTID-B, we cloned the gene encoding BTID-B from B. thermoleovorans ID-1 and expressed it with plasmid pUC19 in E. coli DH5␣ [24]. Recombinant BTID-B was purified from a cell extract of E. coli (Table 3). After disruption of cells by sonication, the cell extract was heattreated for 10 min at 80°C and centrifuged to remove the denatured E. coli proteins. The heat-treated cell extract was subjected to ion-exchange chromatography on DEAESepharose CL6B column. The fractions containing high enzyme activity were eluted at 100 mM NaCl. Overall 11.5-fold purification was achieved with 4% yield. Further purification with Sephacryl S200 gel chromatography resulted in 108-fold purification with a specific activity of 292.4 U/mg and a yield of 3.2%.

BTID-A and BTID-B was compared with the lipases from other thermophilic Bacillus strains. The half lives of the lipases from B. catenulatus and B. stearothermophilus L1 were 30 min at 62°C and 30 min at 60°C, respectively. In

3.3. Physicochemical properties of the purified lipases The molecular mass was determined for both purified lipases by SDS-PAGE (Fig. 2a and 2b). On SDS-PAGE, the molecular mass of BTID-A from B. thermoleovorans ID-1 was estimated to be 19 kDa, which was much smaller than those of other thermostable lipases previously characterized except one from B. thermocatenulatus (16 kDa) [11]. Interestingly, this enzyme was produced by B. thermoleovorans ID-1 without any lipid substrate in the culture media. This result showed that BTID-A is not an inducible enzyme. This is a distinct property from those of other lipases from thermophiles as well as from mesophiles previously reported. The lipases from Bacillus sp. H-257 (24 kDa) [30], B. thermocatenulatus (16 kDa) [11], B. licheniformis [9], Pseudomonas pseudoalcaligenes F-111 (32 kDa) [15], P. aeruginosa [31], Penicillium expansum (25 kDa) [20], Geotrichum candidum (56 kDa) [21], were reported to be inducible. Apparent molecular weight of BTID-B was 43 kDa when analyzed on SDS-PAGE, which was slightly smaller than that deduced from the DNA sequence (46 kDa). The optimal reaction temperature of BTID-A was observed at 60 – 65°C and that of BTID-B at 60°C (Fig. 3). When BTID-A was incubated for 30 min at 60°C, its residual activity was over 75% (Fig. 3a). Thermostability of

Fig. 2. SDS-PAGE of BTID-A (A) and BTID-B (B) from B. thermoleovorans ID-1. (A) Lane 1, molecular weight marker; lane 2, culture supernatant; lane 3, ammonium sulfate precipitate; lane 4, DEAE-Sepharose CL6B pool; lane 5, Superdex 200 pool; lane 6, Resource PHE pool; lane 7, purified BTID-A (B) Lane 1, molecular weight marker; lane 2, cell extract; lane 3, heat-treated sample; lane 4, DEAE-Sepharose CL6B pool; lane 5, purified BTID-B.

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Fig. 3. Effect of temperature on the activities (f ) of purified BTID-A (A) and BTID-B (B), and their thermostabilities (䡬). Enzyme activities at various temperatures were determined as described in the text.

contrast, that of BTID-B was 30 min at 70°C and very stable for 2 h at 60°C (Fig. 3b). These results indicated that lipases from B. thermoleovorans ID-1 are more stable than those from other Bacillus species at high temperatures. The pH optima of BTID-A and BTID-B for lipase activity were 9 and 8 –9, respectively (Fig. 4a and 4b). Both of them were stable at neutral pH range [6 – 8] for 24 h at room temperature (data not shown). Comparison of the deduced amino acid sequence and physicochemical characteristics of BTID-B with other lipases showed that it was similar to those of the lipases from B. catenulatus [10] and B. stearothermophilus L1 [12]. High activity of the enzymes at pH 9 –10 might result from the substitution of Ala for the first Gly-residue in the consensus sequence Gly-X-Ser-X-Gly [1]. The effect of various cations at a concentration of 1 mM on the activities of BTID-A and BTID-B were as-

Fig. 4. Effect of pH on purified BTID-A (A) and BTID-B (B). Enzyme activities at various pH values were determined as described in the text. Sodium acetate buffer was used for pH 4 to 6 (〫); potassium phosphate buffer for pH 6 to 8 (f); Tris-HCl buffer for pH 7 to 9 (‚); glycine-NaOH buffer for pH 9 to 11 (●).

sessed (Table 4). Ca2⫹, Co2⫹, Na⫹, and Mn2⫹ ions were found to enhance BTID-B activity. In contrast, none of the above ions could enhance BTID-A activity, and Cu2⫹ and Fe2⫹ inhibited BTID-A. As shown in Table 5, some organic solvents and inhibitors at a concentration of 1% (v/v) were inhibitory on BTID-A activity. BTID-A was strongly inhibited by PMSF, suggesting that BTID-A is a serine esterase. In contrast, BTID-B activity was enhanced by organic solvents such as DMSO, EtOH, and ␤-mercaptoethanol. This feature could be applicable for the synthesis of chiral compounds in nonaqueous solvents [5]. EDTA treatment strongly inhibited both enzymes, confirming that the enzymes require metal ions for their activity.

D.-W. Lee et al. / Enzyme and Microbial Technology 29 (2001) 363–371 Table 4 Effect of metal ions on the activities of purified BTID-A and BTID-B Metal ion

None CaCl2 CoCl2 MgCl2 HgCl2 KCl NaCl ZnSO4 MnSO4 CuSO4 FeSO4 AgNO3 *

Concn (mM)

1 1 1 1 1 1 1 1 1 1 1 1

Relative activity (%)* BTID-A

BTID-B

100 87 62 75 54 62 80 55 55 32 35 34

100 121 119 — 87 120 124 63 124 56 64 109

Table 6 Substrate specificities of purified BTID-A and BTID-B Substrate

Tributyrin (C4) Tricaproin (C6) Tricaprylin (C8) Tricaprin (C10) Trilaurin (C12) Trimyristin (C14) Tripalmitin (C16) Tristearin (C18) 1,2,3-Trioleoyl-glycerol (OOO) 1,3-Dipalmitoyl-2-oleoyl-glycerol (POP) 1,2-Dilinoleoyl-3-oleoyl-rac-glycerol (LLO) 1,3-Dioleoyl-2-palmitoyl-glycerol (OPO) *

Data are the averages of values at least three experiments.

3.4. Substrate specificity and positional analysis of lipases Both BTID-A and BTID-B showed broad specificities for triglycerides tested, although the substrate profiles differed for each enzyme (Table 6). Among the substrates tested, tricaprylin (C8) and tricaprin (C10) were almost completely hydrolyzed by both enzymes. In addition, BTID-A showed a high activity toward 1,2-dilinoleoyl-3oleoyl-rac-glycerol (LLO), a major triglyceride in soy bean oil. But it could not hydrolyse short chain triacylglycerides such as tributyrin (C4) and tricaproin (C6), and long chain unsaturated ones such as 1,3-dipalmitoyl-2-oleoyl-glycerol (POP) and 1,3-dioleoyl-2-palmitoyl-glycerol (OPO). On the other hand, BTID-B showed a high activity toward long chain saturated triglycerides, but slight activity toward the unsaturated triglycerides such as POP and OPO. In order to determine the positional specificities of both enzymes, thin layer chromatography (TLC) was performed (Fig. 5). The major hydrolysis products by BTID-B were 1,2- and 1,3-diolein, whereas 2-monoolein was also detected as a main product by BTID-A. These results indicated that BTID-A is likely to be an 1,3-specific lipase and

369

Relative activity (%)* BTID-A

BTID-B

0 0 100 94 39 42 31 15 4 3 97 0

7 67 99 100 95 58 99 55 20 19 46 12

Data are the averages of values at least three experiments.

BTID-B a positional nonspecific one. The difference of substrate specificities and positional specificities of both enzymes suggests that these distinct enzymes probably act synergistically to hydrolyze extracellular lipids to free fatty acids and glycerols. 3.5. Kinetic parameters Kinetic experiments were performed by using the standard activity assay with tricaprylin (C8) as a substrate. BTID-A and BTID-B exhibited simple Michaelis-Menten kinetics for tricaprylin. In a series of cases, the Km values for both lipases were determined to be 1.82 mM and 6.24 mM, respectively. From the Lineweaver-Burk plot, their Vmax values were calculated to be 12.8 ␮mol min⫺1 mg⫺1 and 63.3 ␮mol min⫺1 mg⫺1, respectively.

Table 5 Effect of organic solvents and inhibitors on the activities of purified BTID-A and BTID-B Reagents

None SDS EtOH ␤-mercaptoethanol Isopropyl alcohol DMSO PMSF DTT EDTA *

Concn (%, w/v)

1 1 1 1 1 1 1 1 1

Relative activity (%)* BTID-A

BTID-B

100 97 58 74 47 46 8 52 32

100 — 124 135 130 126 124 129 55

Data are the averages of values at least three experiments.

Fig. 5. TLC analysis of hydrolysis products after incubation of purified BTID-A and BTID-B on triolein as a substrate at 60°C for 30 min. Lane 1, standard materials (triolein, oleic acid, 1,2-diolein, 1,3-diolein, 2-monoolein); lane 2, control(without enzyme solution); lane 3, BTID-A; lane 4, BTID-B.

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3.6. N-terminal sequence analysis The N-terminal sequence of BTID-A was determined as ATLIELLAVIVILGI over the first fifteen amino acid residues. When compared with the sequences of other Bacillus lipase (B. stearothermophilus, B. thermocatenulatus, B. subtilis, and B. pulmilis) deposited in the EMBL 29.0 and Swissprot 20.0 databases, no significant homology was found. Although the fifteen N-terminal amino acid sequence of BTID-A showed high identity to that of the lipase (16 kDa) from B. thermocatenulatus, BTID-A had several distinct features. First, BTID-A may be a constitutive enzyme, whereas the lipase from B. thermocatenulatus was an inducible one [11]. In addition, BTID-A showed a high activity with tricaprylin (C8) and about two times more thermostable than the lipase from B. thermocatenulatus [10 – 12]. That BTID-A seems to be an 1,3-specific lipase distinguish it from other lipases.

4. Conclusion Described here is the purification and characterization of two distinct thermophilic lipases (BTID-A and –B) from B. thermoleovorans ID-1. B. thermoleovorans ID-1 has several advantageous features for industrial applications since the organism could produce multiple lipases, regardless of lipid substrates. The thermostable, lipid-hydrolyzing enzymes may be applied to treat lipid-rich industrial effluents treatments, to produce inter-esterification substances in food industry, or to synthesize useful chemical compounds. In addition, studies on B. thermoleovorans ID-1 as well as their lipases may lead to further understanding on the evolution of the thermophilic, Gram-positive Bacillus rRNA group 5.

5. Synopsis of the paper This paper presents a few fundamental experiments on lipid hydrolysis for the purpose on industrial applications. Purification and characterization for two lipases derived from Bacillus thermoleovorans ID-1 was also done. One purification was done from culture fluid, the other from an E.coli clone. Several comparative experiments were done with the two enzymes. The most significant differences between previously characterized lipases and those described here have to do with inducibility in the case of BTID-A, differences in substrate and inhibitor specificity, and thermostability.

Acknowledgments This work [985–1200-004 –2] was supported by the Korea Science and Engineering Foundation (KOSEF). Tech-

nical support for this work by Prof. G. Antranikian and H. Ma¨ rkl (Technical University of Hamburg-Harburg, Germany) is greatly appreciated. We express our gratitude to Dr. M.A. Yu for her help on preparing this paper.

References [1] Jaeger KE, Ransac S, Dijkstra BW, Colson C, Heuvel MV, Misset O. Bacterial lipases. FEMS Microbiol Rev 1994;15:29 – 63. [2] Verger R. Interfacial activation of lipases: facts and artifacts. Tibtech 1997;15:32– 8. [3] Jaeger KE, Reetz MT. Microbial lipases form versatile tools for biotechnology. Tibtech 1998;16:396 – 493. [4] Schmidt-Dannert, C. Recombinant microbial lipases for biotechnological applications. Bioorg Med Chem 1999;7:2123–30. [5] Zaks A, Klibanov AM. Enzymatic catalysis in nonaqueous solvents. J Biol Chem 1998;263:3194 –201. [6] Niehaus F, Bertoldo C, Ka¨ hler M, Antranikian G. Extremophiles as a source of novel enzymes for industrial application. Appl Microbiol Biotechnol 1999;51:711–29. [7] Pennisi E. In industry, extremophiles begin to make their mark. Science 1997;276:705– 6. [8] Dartois V, Baulard A, Schanck K, Colson C. Cloning, nucleotide sequence and expression of a lipase gene from Bacillus subtilis 168. Biochim Biophys Acta 1992;1131:253– 60. [9] Khyami-Horani H. Thermotelerant strain of Bacillus licheniformis producing lipase. World J Microbiol Biotechnol 1996;12:399 – 401. [10] Schmidt-Dannert C, Luisa Ru´ a M, Atomi H, Schmid RD. Thermoalkalo-philic lipase of Bacillus thermocatenulatus. ⌱. Molecular cloning, nucleotide sequence, purification and some properties. Biochim Biophys Acta 1996;1301:105–14. [11] Schmidt-Dannert C, Sztajer H, Sto¨ cklein W, Menge U, Schmid RD. Screening, purification and properties of a thermophilic lipase from Bacillus thermocatenulatus. Biochim Biophys Acta 1994;1214:43–53. [12] Kim, HK, Park SY, Lee JK, Oh TK. Gene cloning and characterization of thermostable lipase from Bacillus stearothermophilus L1. Biosci Biotechnol Biochem 1998;62(1):66 –71. [13] Kim MH, Kim HK, Lee JK, Park SY, Oh TK. Thermostable lipase of Bacillus stearothermophilus: high-level production, purification, and calcium-dependent thermostability. Biosci Biotechnol Biochem 2000;64:280 – 6 [14] Choo DW, Kurihara T, Suzuki T, Soda K, Esaki N. A cold-adapted lipase of an Alaskan psychrotroph, Pseudomonas sp. strain B11–1: Gene cloning and enzyme purification and characterization. Appl Environ Microbiol 1998;64:486 –91. [15] Lin SF, Chiou CM, Yeh CM, Tsai YC. Purification and partial characterization of an alkaline lipase from Pseudomonas pseudoalcaligenes F-111. Appl Environ Microbiol 1996;62:1093–5. [16] Stuer W, Jaeger KE, Winkler UK. Purification of extracellular lipase from Pseudomonas aeruginosa. J. Bacteriol. 1986;168:1070 – 4. [17] Drouault S, Corthier G, Ehrlich D, Renault P. Expression of the Staphylococcus hyicus lipase in Lactococcus lactis. Appl Environ Microbiol 2000;66:588 –98. [18] Li X, Tetling S, Winkler UK, Jaeger KE, Benedik MJ. Gene cloning, sequence analysis, purification and secretion by Escherichia coli of an extracellular lipase from Serratia marcescens. Appl Environ Microbiol 1995;61:2674 – 80. [19] Sommer P, Bormann C, Go¨ tz F. Genetic and biochemical characterization of a new extracellular lipase from Streptomyces cinnamomeus. Appl Environ Microbiol 1997;63:3553– 60 [20] 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.

D.-W. Lee et al. / Enzyme and Microbial Technology 29 (2001) 363–371 [21] Veeraragavan K, Colpitts T, Gibbs B. Purification and characterization of two distinct lipases from Geotrichum candidum. Biochim Biophys Acta 1990;1044:26 –33 [22] Becker P, Abu-Reesh I, Markossian S, Antranikian G, Ma¨ rkl H. Determination of the kinetic parameters during continuous cultivation of the lipase-producing thermophile Bacillus sp. IHI-91 on olive oil. Appl Microbiol Biotechnol 1997;48:184 –90. [23] Lee DW, Koh YS, Kim KJ, Kim BC, Choi HJ, Kim DS, Suhartono MT, Pyun YR. Isolation and characterization of a thermophilic lipase from Bacillus thermoleovorans ID-1. FEMS Microbiol Lett 1999; 179:393– 400. [24] Cho AR, Yoo SK, Kim EJ. Cloning, sequencing and expression in Escherichia coli of a thermophilic lipase from Bacillus thermoleovorans ID-1. FEMS Microbiol Lett 2000;186:235– 8. [25] Yanish-Perron C, Vieira J, Messing J. Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13 mp18 and pUC19 vectors. Gene 1985;33:102–19.

371

[26] Sambrook J, Fritsch EF, Maniatis T. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor NY: Cold Spring Harbor Laboratory, 1989. [27] Kouker G, Jaeger KE. Specific and sensitive plate assay for bacterial lipases. Appl Environ Microbiol 1987;53:211–3. [28] Laemmli UK. Cleavage of structural proteins dufing the assembly of the head of bacteriophage T4. Nature 1970;227:680 –5. [29] Sztajer H, Lu¨ nsdorf H, Erdmann H, Menge U, Schmid RD. Purification and properties of lipase from Penicillium simplicissimum. Biochim Biophys Acta 1992;1124:253– 61. [30] Imamura S, Kitaura S. Purification and characterization of a monoacylglycerol lipase from the moderately thermophilic Bacillus sp. H-257. J Biochem 2000;127:419 –25. [31] Jane Gilbert E, Drozd Jan W, Jones CW. Physiological regulation and optimization of lipase activity in Pseudomonas aeruginosa EF2. J Gen Microbiol 1991;137:2215–21.