INTRODUCTION

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Lipases catalyze the hydrolysis of acylglycerides and other fatty acid esters. They resemble esterases, but differ markedly from them in their ability to act on water-insoluble esters (5). Lipases and esterases have been recognized as very useful biocatalysts because of their wide-ranging versatility in industrial applications.

A variety of microbial lipases with different enzymological properties and substrate specificities have been found (22). The temperature stability of lipases has been regarded as the most important characteristic for use in industry (16). However, low stability is favorable for some purposes. For example, heat-labile enzymes can be easily inactivated by treatment for short periods of time at relatively low temperatures after being used for processing of food and other materials (32). One can therefore prevent the materials from damage during heat inactivation of the enzymes. Cold-adapted microorganisms, which are expected to produce cold-adapted enzymes, have been isolated. These microorganisms usually grow only slowly even under appropriate conditions (19). However, recent advances in genetic engineering have enabled efficient production of heterologous enzyme genes in an appropriate host strain such as Escherichia coli. Thus, cold-adapted enzymes from psychrotrophic microorganisms showing high catalytic activity at low temperatures can be highly expressed in such recombinant strains. The cold-adapted enzymes are expected to be applicable as additives to detergents used at low temperatures and biocatalysts for biotransformation of labile compounds at cold temperatures (32).

Recently, the genes of cold-adapted lipases from psychrotrophic bacteria Moraxella TA144 (10) and Psychrobacter immobilis B10 (1) isolated in Antarctica were cloned and sequenced. The enzymes showed high activities at temperatures as low as 3°C. None of these recombinant enzymes, however, have been purified to homogeneity due to strong interaction of the enzymes with lipopolysaccharides secreted by the bacterial cells (1).

Our group has also isolated cold-adapted microorganisms producing cold-adapted lipases from Alaskan and Siberian soils and cloned the lipase gene, lipP, from an Alaskan psychrotroph, Pseudomonas sp. strain B11-1. In this study, we report the cloning and sequencing of the lipP gene and the purification and characterization of the recombinant enzyme LipP.

	MATERIALS AND METHODS

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Bacterial strains and media. Psychrotrophic bacteria were isolated from tundra soils of Alaska and Siberia on agar plates of Luria-Bertani (LB) medium (pH 7.6) at 4°C. Lipase activity of the bacteria was detected through the formation of halos around the colonies in three kinds of agar plates as follows. The basal medium (1% tryptone, 0.5% yeast extract, 0.5% NaCl, pH 7.2) was supplemented with medium A, 0.01% CaCl₂ · 2H₂O and 1% Tween 80 (40); medium B, 0.01% CaCl₂ · 2H₂O and 1% tributyrin (26); or medium C, 2.5% olive oil and 0.001% rhodamine B (25). Lipase production on medium C plates was monitored by fluorescence with UV light at 350 nm.

Construction of a genomic DNA library and screening for a lipase gene from Pseudomonas sp. strain B11-1. DNA manipulation was carried out according to the methods described by Sambrook et al. (38). Restriction enzymes and DNA-modifying enzymes were purchased from Takara Shuzo, Kyoto, Japan. E. coli C600 was used as a host cell with pUC118 (Takara Shuzo, Kyoto, Japan) as a cloning vector. The chromosomal DNA was isolated from Pseudomonas sp. strain B11-1 cells by phenol treatment (37) and was partially digested with Sau3AI at 37°C. The resulting DNA fragments were electrophoresed in 0.8% agarose gel, and fragments of 1 to 10 kbp were electroeluted and then ligated with pUC118 which had been previously digested with BamHI and dephosphorylated with bacterial alkaline phosphatase. The resultant plasmids were introduced into E. coli C600 according to a previously described method (33), providing a gene library containing 35,000 recombinant E. coli clones. The clone cells producing a lipase were detected due to the formation of halos around the colonies on LB agar plates supplemented with ampicillin (0.1 mg/ml), 0.1 mM IPTG (isopropyl--D-thiogalactopyranoside), 1% tributyrin, and 1% gum arabic. After incubation at 28°C for 16 h, the plates were incubated further at 4°C for 4 days.

Assays. Lipase activity was determined with a lipase assay kit (Dainippon Pharmaceutical, Osaka, Japan) at 25°C. One unit of enzyme activity was defined as the amount of enzyme that catalyzes the formation of 1 µmol of 2,3-dimercaptopropan-1-ol from 2,3-dimercaptopropan-1-ol tributyl ester per min (27). The amount of protein was determined with a Bio-Rad protein assay kit with bovine serum albumin used as the standard (3). Esterase activity was determined by measuring p-nitrophenol formed from fatty acid p-nitrophenyl esters at 25°C. The following molar extinction coefficients of p-nitrophenol at 400 nm were used: 14,775 M¹ cm¹ in 0.1 M sodium phosphate buffer containing 0.1 M NaCl (pH 7.25) (36) and 15,100 M¹ cm¹ in Tris-HCl (pH 8.0) (24). The assay mixture (1 ml) contained p-nitrophenyl butyrate, 0.1 M sodium phosphate buffer containing 0.1 M NaCl (pH 7.25), and 1 µg of enzyme. Long-chain fatty acid esters of p-nitrophenol (e.g., p-nitrophenyl laurate and p-nitrophenyl palmitate) are not soluble in this assay mixture, and substrate specificity for various p-nitrophenyl esters was examined in a mixture containing 0.5 mM p-nitrophenyl esters, 0.1 M sodium phosphate buffer (pH 7.25), 0.1 M NaCl, 15% acetonitrile, and 0.038 mM Triton X-100. After preincubation for 10 min at the designated temperatures (from 4 to 70°C), the reaction was initiated by addition of the substrate.

Purification of LipP. All operations were performed at 4°C, and 20 mM Tris-HCl (pH 8.0) was used as the standard buffer unless otherwise stated.

DNA sequencing. All bases were sequenced at least once in each direction by the chain termination method with a Dye Primer or a Dye Terminator sequencing kit, by using an Applied Biosystem model 370A DNA sequencer. Sequence analysis was carried out with software from DNASTAR, Inc. (Madison, Wis.).

Nucleotide sequence accession number. The nucleotide sequence reported in this work has been assigned GenBank accession no. AF034088.

	RESULTS AND DISCUSSION

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Isolation of Pseudomonas sp. strain B11-1. We screened various psychrotrophic bacteria isolated from tundra soils of Alaska and Siberia (total, 88 strains) for bacterial strains showing high lipase activity at cold temperatures. The psychrotrophic bacteria were examined with a screening system suitable for detection of lipase producers as described in Materials and Methods. Strain B11-1 was selected as the best producer of lipase and was identified as a pseudomonad by the German Collection of Microorganisms, Braunschweig, Germany, on the basis of its taxonomic characteristics: motile by polar flagella; number of flagella, >1; rod shaped (0.5 to 0.8 by 1.5 to 3.0 mm); gram negative; anaerobic growth, negative; spore formation, negative; oxidase test, positive; catalase test, positive; O-F test, oxidative; urease test, negative; denitrification, positive; diffusible pigment formation, negative; gelatin hydrolysis, negative; carbon sources utilized: glucose, malate, adonitol, mannitol, sorbitol, -alanine, and L-valine; and nonutilized carbon sources: arabinose, L-rhamnose, adipate, citraconate, and erythritol. The 16S rRNA sequence of strain B11-1 showed a similarity of 99.3% to that of Pseudomonas syringae, but they were distinct from each other in a few physiological characteristics, in particular, those shown by the oxidase and denitrification tests (15). No strain identical to strain B11-1 could be found, and we named strain B11-1 Pseudomonas sp. strain B11-1.

Cloning and nucleotide sequencing of the lipase gene. A library of the chromosomal genes of Pseudomonas sp. strain B11-1 was constructed with vector plasmid pUC118 and host strain E. coli C600. About 35,000 recombinant colonies were isolated on the agar plates with medium B. Six clones showed clear halos around the colonies at 4°C due to hydrolysis of tributyrin in the medium. We selected the clone forming the clearest and largest halo and designated the cloned gene lipP. The cloned plasmid encoding lipP was named pPL2-1; it contained an insert DNA of approximately 2.5 kbp. Since the recombinant E. coli cells harboring pPL2-1 formed a clear halo in the presence IPTG, LipP is probably expressed under the control of its inherent promoter in E. coli. We determined the nucleotide sequence of the fragment and found a single open reading frame comprising 924 bp, which encodes a putative protein of 308 amino acids with a predicted molecular weight of 33,714. A putative Shine-Dalgarno sequence (5'-GAAGGA) was found seven bases upstream from the initiation codon ATG. The sequence GDSAG starting at residue 153 fits the GXSXG motif shared with various lipases, esterases, and other hydrolytic enzymes. Therefore, Ser155 probably acts as the nucleophile to form an acyl intermediate with the substrate as do many hydrolytic enzymes. The active-site serine is encoded by AGC in the same manner as various other lipases: AGY (Y, a pyrimidine) is known as the common code for the active-site serine residue of most lipases (34).

Comparison of amino acid sequences. The amino acid sequence of LipP was compared with those of other lipases. LipP shows a high sequence similarity to Lipase 2 from psychrotrophic Moraxella TA144 (12) and the mammalian hormone-sensitive lipase (28, 29), with homologies of 28.5 and 22.7%, respectively. The putative lipolytic proteins such as the E. coli lipase-like protein and Bacillus acidocaldarius ORF3 protein also show some sequence similarity to those of the three above-mentioned enzymes (28). However, the functions of these hypothetical lipases have not yet been clarified. LipP probably uses a catalytic triad (Ser-His-Asp/Glu) and contains an oxyanion hole to stabilize the tetrahedral intermediate during the acetylation and deacetylation steps (8). The consensus sequence of GDSAG occurs in LipP, and the central serine (Ser155) is probably a member of the triad. The enzyme has another conserved sequence in the region including Asp250, Asp254, and Glu255, and one of these acidic amino acid residues presumably participates in catalysis as a member of the triad. Two conserved histidine residues, His81 and His280, occur, and one of them is thought to form the triad. The histidine residue constituting the triad is quite often followed by a glycine. Both His81 and His280 precede a glycine residue, and either of them could be the catalytic His residue, although we have at present no information to predict which is more plausible.

Purification of LipP. LipP was purified with a 17% yield by 38-fold purification (Table 1). When the purity was judged by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, only a single major protein band, whose molecular mass (about 33 kDa) is consistent with the subunit molecular weight (33,714) of LipP deduced from the nucleotide sequence of lipP, was observed (Fig. 1).

View larger version (78K): [in this window] [in a new window]   FIG. 1.   Polyacrylamide gel electrophoresis of LipP. Lane 1, molecular weight markers; lane 2, the preparation after ammonium sulfate precipitation (amount of protein loaded, 25 µg); lanes 3 and 4, the preparation after DEAE-Cellulofine chromatography (amount of protein loaded, 15 µg); lanes 5 and 6, the preparation after Gigapite chromatography (amount of protein loaded, 10 µg).

Substrate specificity. Substrate specificity of LipP was examined with p-nitrophenyl esters of various fatty acids. The rates of hydrolysis for the following fatty acids were as indicated (in units/milligram): butyrate, 164; valerate, 64; caproate, 119; caprylate, 41; laurate, 12; and palmitate, 8. The enzyme showed high activity towards short- to medium-chain (C₄ and C₆) fatty acids, and the esters of longer-chain fatty acids were poor substrates. A similar specificity was found for the crude preparation of Lipase 2 from Moraxella sp. strain TA144 (10) and lipases of psychrotrophic pseudomonads isolated from refrigerated milk (23). The recombinant lipase from psychrotrophic Pseudomonas fluorescens SIK W1 also acted on the esters of medium-chain (C₆ and C₈) fatty acids (30). Positional specificity of LipP for triolein was examined by thin-layer chromatography (7). The products were analyzed after incubation for 10, 30, 60, and 120 min at 30°C: 1,2-diolein and monoolein were formed, but only a small amount of 1,3-diolein was accumulated. The results indicate that LipP has a 1,3-positional specificity toward the fatty acid triglyceride.

Effect of pH on lipase activity. Figure 2A shows the pH stability of LipP at 25°C with 0.5 mM p-nitrophenyl butyrate as a substrate. The enzyme was stable between pH 6 and 9 at the indicated pH range when incubated at 0°C for 24 h, but its activity decreased at acidic and alkaline pH values. The recombinant lipases from Pseudomonas pseudoalcaligenes and Pseudomonas mendocina were reported to be stable between pH 5 and 10, whereas the enzyme from Pseudomonas cepacia DSM 50181 was reported to be stable under acidic (pH < 2.0) and alkaline (pH > 12.0) conditions (44). Figure 2B shows the optimum pH for the LipP reaction with tributyrin as a substrate. The optimal pH was found to be around 8.0. The pH optima for the reactions catalyzed by Pseudomonas cepacia, Pseudomonas pseudoalcaligenes, and Pseudomonas mendocina enzymes were reported to be between 8 and 9 (44).

View larger version (11K): [in this window] [in a new window]   FIG. 2.   Effect of pH on the stability (A) and activity (B) of LipP. (A) The enzyme was incubated at various pH values at 4°C for 24 h, and the activity was measured. The enzyme activity after treatment with Tris-HCl (pH 9.0) was taken as 100%. Buffers used (final concentration, 20 mM) were glycine-HCl () (pH 2.2 to 3.6); sodium acetate () (pH 3.6 to 5.6); Tris-malate () (pH 5.6 to 8.6); Tris-HCl () (pH 7.4 to 9.0); glycine-NaOH () (pH 8.6 to 10.6). (B) The enzyme was incubated with 100 mM tributyrin as a substrate in NaH2PO4-NaOH buffer at various pH values at 25°C for 10 min, and free fatty acid formed was titrated with 0.05 M NaOH.

Effect of temperature on enzyme activity. The enzyme showed maximal activity at 45°C toward p-nitrophenyl butyrate (Fig. 3A) and at 37°C toward p-nitrophenyl laurate (data not shown). The activation energy of an enzyme reaction reflects the catalytic efficiency of the enzyme; low activation energy is due to the high catalytic efficiency of the enzyme. The reactions catalyzed by the enzymes derived from cold-adapted organisms are usually lower than those catalyzed by the corresponding enzymes from their mesophilic counterparts (11). Therefore, we determined the activation energy for the hydrolysis of p-nitrophenyl butyrate catalyzed by LipP. It was about 11.2 kcal/mol in the range 5 to 35°C (Fig. 3B) and constant over the assay temperatures below 35°C. Such a monophasic feature suggests that LipP does not undergo structural changes in this temperature range, although at higher temperatures LipP was inactivated irreversibly. Lower activation energy (7.7 kcal/mol) was observed toward p-nitrophenyl laurate. This value is much lower than those for the same substrate shown by enzymes from other sources: Antarctic bacteria, 12 to 17 kcal/mol, and mesophilic Pseudomonas aeruginosa, 25 kcal/mol (9). This indicates that the catalytic efficiency is much higher for LipP than for these enzymes. However, at temperatures above 40°C, enzyme activity fell drastically at an inactivation energy of 70.4 kcal/mol.

View larger version (10K): [in this window] [in a new window]   FIG. 3.   Effect of temperature on the activity of LipP. (A) The enzyme was incubated with a mixture containing 20 mM phosphate buffer (pH 7.25), 5% acetonitrile, and 0.5 mM p-nitrophenyl butyrate at various temperatures for 10 min, and p-nitrophenol formed was measured. The value obtained at 37°C was taken as 100%. (B) The logarithm of the specific activity (V) (in micromoles per milligram per minute) was plotted against the reciprocal of absolute temperature (T). The values shown are activation energy calculated from the linear part of the plot.

Thermostability. The enzyme was incubated at various temperatures (30, 40, 50, 60, and 70°C) for 30 min, and then the residual activity was measured at 25°C (100, 88, 66, 25, and 0%, respectively). Remaining activities after 60-min incubations at the same temperatures were 92, 85, 47, 0, and 0%, respectively. LipP is a little more stable than enzymes from other psychrotrophs, e.g., Moraxella TA144 (9) and Acinetobacter O₁₆ (4).

Effect of inhibitors. The enzyme was incubated with various compounds that may inhibit the enzyme, and the remaining activity was measured with p-nitrophenyl butyrate as the substrate at 25°C. The enzyme was not affected by phenylmethylsulfonyl fluoride, EDTA, and 2-mercaptoethanol but was strongly inhibited by Zn²⁺, Cu²⁺, Fe³⁺, and Hg²⁺ ions (Table 2). Hormone-sensitive lipase of rat adipose tissue is also inhibited by Hg²⁺ ions, indicating the occurrence of an essential thiol group of this family of lipases (13). Both Fe²⁺ and Fe³⁺ ions were found to inhibit the lipase from Aspergillus niger (20). On the other hand, the lipases from A. niger and Humicola lanuginosa were activated by Ca²⁺ ions, which facilitated the removal of free fatty acids formed in the reaction at the water-oil interface (21, 31). However, LipP was not activated by the addition of Ca²⁺ ions. Human liver arylacetamide deacetylase, which shares the sequences GDSAG and HGGG with LipP, was inhibited by bis-p-nitrophenyl phosphate (35), whereas LipP was not inhibited by this compound.

Effect of temperature on the Michaelis constant. Somero suggested that enzyme-ligand interactions are disrupted by both increases and decreases in temperature (41, 42). Various enzymes from poikilothermal organisms, including thermophilic and psychrophilic microorganisms, show the lowest K_m values for their substrates at the physiological temperatures of the source organisms. Trout produce the warm- and cold-adapted forms of acetylcholinesterase and citrate synthase according to their habitat temperature, and the K_m values of the enzymes are at a minimum at their acclimated temperatures (2, 17). Urocanase from a psychrotroph, Pseudomonas putida, shows a low K_m at low temperatures (18). On the other hand, the K_m of the thermostable enolase from the thermophile Thermus aquaticus YT-1 increases at low temperatures, and the lowest K_m is at high temperatures, near the optimal growth temperature for the strain (43). We examined the apparent K_m of LipP for p-nitrophenyl butyrate at various temperatures (Fig. 4). The K_m decreased with decreases in temperature, and the lowest K_m was observed in the range 5 to 15°C; this is consistent with the physiological temperature of the source organism.

View larger version (13K): [in this window] [in a new window]   FIG. 4.   Effect of temperature on the Michaelis constant of LipP. The enzyme was assayed with a mixture containing 0.1 M phosphate buffer (pH 7.25), 0.1 M NaCl, and various concentrations of p-nitrophenyl butyrate (0.005 to 0.5 mM) at 25°C. The Km for p-nitrophenyl butyrate was obtained from a plot of velocity versus substrate concentration with KaleidaGraph software (Synergy Software, Reading, Pa.). The values of four separate experiments were averaged and plotted against temperature.

Effect of organic solvents on enzyme activity. LipP was incubated with various water-miscible organic solvents at a concentration of either 15 or 30% (vol/vol) at 25°C for 1 h. The enzyme was not inhibited; rather, it was slightly activated by all the solvents examined except acetonitrile (Table 3). The enzyme was completely inactivated either by 30% acetonitrile at 25°C for 1 h or by 15% acetonitrile at 37°C for 5 min (data not shown). The lipase from Fusarium heterosporum is also a solvent-resistant enzyme but was completely inactivated in 50% acetonitrile (39). Other lipases from Pseudomonas and Bacillus were also activated in the presence of several water-miscible organic solvents (39). Although the three-dimensional structure of LipP has not yet been clarified, the solvents probably convert the closed form of the enzyme to the open form by affecting a lid or flap in LipP to activate the enzyme. The Candida rugosa lipase is also known to be activated by organic solvents, which keep the enzyme in the open conformation; the lid of the enzyme does not cover the active site-crevice, thus keeping a flexible conformation (6, 14). Whatever the mechanism of the activation of LipP by organic solvents is, stability to solvents is a useful characteristic of the enzyme.