Directed evolution of Pseudomonas aeruginosa lipase for improved amide-hydrolyzing activity

Ryota Fujii1,2, Yuichi Nakagawa1, Jun Hiratake1,3, Atsushi Sogabe4 and Kanzo Sakata1

1Institute for Chemical Research, Kyoto University, Gokasho, Uji, Kyoto 611-0011, Japan and 4Tsuruga Institute of Biotechnology, Toyobo Co., Ltd, 10–24 Toyo-Cho, Tsuruga, Fukui 914-0047, Japan

3 To whom correspondence should be addressed. E-mail: hiratake{at}scl.kyoto-u.ac.jp


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
A lipase from Pseudomonas aeruginosa was subjected to directed molecular evolution for increased amide-hydrolyzing (amidase) activity. A single round of random mutagenesis followed by screening for hydrolytic activity for oleoyl 2-naphthylamide as compared with that for oleoyl 2-naphthyl ester identified five mutants with 1.7–2.0-fold increased relative amidase activities. Three mutational sites (F207S, A213D and F265L) were found to affect the amidase/esterase activity ratios. The combination of these mutations further improved the amidase activity. Active-site titration using a fluorescent phosphonic acid ester allowed the molecular activities for the amide and the ester to be determined for each mutant without purification of the lipase. A double mutant F207S/A213D gave the highest molecular activity of 1.1 min–1 for the amide, corresponding to a 2-fold increase compared with that of the wild-type lipase. A structural model of the lipase indicated that the mutations occurred at the sites near the surface and remote from the catalytic triad, but close to the calcium binding site. This study is a first step towards understanding why lipases do not hydrolyze amides despite the similarities to serine proteases in the active site structure and the reaction mechanism and towards the preparation of a general acyl transfer catalyst for the biotransformation of amides.

Keywords: amidase activity/directed evolution/esterase activity/Pseudomonas aeruginosa lipase/random mutagenesis


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Lipases (triacylglycerol hydrolases, EC 3.1.1.3 [EC] ) are serine hydrolases that catalyze the hydrolysis of fatty acid esters (triglycerides). In contrast to most other enzymes, lipases have extraordinarily broad substrate specificity and accommodate a wide range of structurally diverse esters, alcohols and carboxylic acids as substrates (Schmid and Verger, 1998Go). Furthermore, lipases are stable and active in organic solvents and show a high degree of enantio- and regioselectivity for the conversion of various unnatural substrates in both aqueous and non-aqueous media (Kazlauskas and Bornscheuer, 1998Go; Klibanov, 2001Go). Hence lipases find wide applications as versatile acyl transfer catalysts for the interconversion of carboxylic esters, acids and alcohols for the preparation of optically active pharmaceuticals and fine chemicals (Kazlauskas and Bornscheuer, 1998Go; Schmid and Verger, 1998Go; Sharma et al., 2001Go).

The active site of lipases consists of a Ser–His–Asp/Glu catalytic triad that resembles the active site of serine proteases. According to the X-ray crystal structures of lipases, the architecture and the relative spatial arrangement of the catalytic residues are very similar to those found in serine proteases (Brady et al., 1990Go; Winkler et al., 1990Go; Schrag et al., 1991Go), although the triad of lipases is approximately a mirror image of those seen in the chymotrypsin, trypsin and subtilisin families (Schrag et al., 1991Go; Ollis et al., 1992Go). Accordingly, lipases catalyze the hydrolysis of esters by the same double displacement mechanism via an acyl enzyme intermediate as observed with serine proteases (Jaeger et al., 1999Go). Of particular interest, however, is that lipases do not hydrolyze amides, whereas serine proteases such as chymotrypsin and subtilisin hydrolyze both amides and esters (Zerner et al., 1964Go; Abrahmsén et al., 1991Go; Bonneau et al., 1991Go). Actually, lipases are very poor catalysts for the conversion of amides (Smidt et al., 1996Go; Wagegg et al., 1998Go; Adam et al., 2000Go; Duarte et al., 2000Go; Forró and Fülöp, 2003Go), which has hampered the use of lipases as catalysts for the biotransformation of amides (Kazlauskas and Bornscheuer, 1998Go; Henke and Bornscheuer, 2003Go). This is due in part to the resonance-stabilized nature of amides compared with esters: the OH-catalyzed hydrolysis of acetamide is more than 103 times slower than that of the corresponding esters (Bender et al., 1962Go; Zerner et al., 1964Go; Bender and Kézdy, 1965Go). The catalytic properties of serine proteases also conform to the general reactivity of amides and esters. Thus, most serine proteases hydrolyze specific ester substrates with acylation rates that exceed those of amides by at least 2–3 orders of magnitude (Bender et al., 1962Go; Zerner et al., 1964Go; Bender and Kézdy, 1965Go; Walsh, 1979Go; Bennet and Brown, 1998Go). Nevertheless, serine proteases hydrolyze amides with significantly higher rates than lipases. The amidase activity of serine proteases, however, is often damaged when structural perturbation is introduced into the enzymes by site-directed mutagenesis (Abrahmsén et al., 1991Go), by chemical modification (Nakatsuka et al., 1987Go; West et al., 1988Go, 1990Go; Zhong et al., 1991Go; Plettner et al., 1999Go; Lloyd et al., 2000Go) and by using hydrophilic organic solvents (Barbas et al., 1988Go; Kidd et al., 1999Go). On the other hand, the esterase activity of serine proteases is much less damaged and can be increased in some cases by these modifications (Lloyd et al., 2000Go). It seems, therefore, that the amidase activity of serine proteases is more ‘sophisticated’ than the activity for esters and that serine hydrolases require this higher degree of catalytic activity for the hydrolysis of energetically more demanding amides. In this regard, serine proteases are considered to be more evolved enzymes than lipases in terms of their ability to hydrolyze both amides and esters. If this is the case, then two questions may arise: first, why do lipases not hydrolyze amides despite the similarities to serine proteases in the active site structure and the reaction mechanism, and second, can lipases be evolved to hydrolyze amides? To address these questions, we started a program for directed evolution of a lipase for improved amide-hydrolyzing (amidase) activity. A combinatorial protein engineering approach (Cherry and Fidantsef, 2003Go; Hult and Berglund, 2003Go; Jaeger and Eggert, 2004Go; Reetz, 2004Go) is advantageous, because the catalytic property of a lipase can be altered without knowledge of the detailed reaction mechanism and the three-dimensional structure and the analysis of mutants may find a clue to the structural and mechanistic basis for the differences between the esterase and the amidase activities. Furthermore, mutant lipases with high amidase activities should expand the scope of applications of lipases to the hydrolysis of amides for the preparation of optically active amines (Balkenhohl et al., 1997Go) and for enzymatic cleavage of structurally diverse amide protective groups under mild conditions (Kadereit and Waldmann, 2001Go).

In this paper, we report the directed evolution of a lipase from Pseudomonas aeruginosa TE3285 for improved amidase activity. The application of random mutagenesis combined with screening for improved activity for oleoyl 2-naphthylamide (2) relative to that for the naphthyl ester (1) identified three mutations responsible for increasing the amidase activity. The combination of the mutations further increased the activity for the amide. The sites of mutation are discussed on the basis of a three-dimensional structural model of the lipase.



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    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Materials

Escherichia coli JM109 and DH5{alpha} were used as a host for manipulating plasmids. Pseudomonas aeruginosa PAO1162 was used for the expression of the lipase. These bacterial strains and pSET2LPL, a plasmid containing the gene of the lipase (lipA) and the lipase activator protein (lipB) (Chihara-Siomi et al., 1992Go; Sogabe et al., 2002Go), were kindly provided by Toyobo (Fukui, Japan). The plasmid was reconstructed for the expression vector pLPLsmA by replacing the tetracycline resistance marker with the streptomycin (Sm) resistance one derived from pHRP310 (Parales and Harwood, 1993Go). The plasmid pHRP310 was a kind gift from the National Institute of Genetics (Shizuoka, Japan). The commercial lipase from P.aeruginosa TE3285 (LPL-312) (Chihara-Siomi et al., 1992Go) and DNA polymerases (rTaq DNA polymerase and KOD-plus) were provided by Toyobo. The PCRx Enhancer System was purchased from Invitrogen. DNA ligation was performed with DNA Ligation Kit Version 2 (Takara). Plasmid DNA was isolated using a QIAprep Miniprep Kit (Qiagen). DNA was extracted from agarose gels by using a MinElute Gel Extraction Kit (Qiagen). Nitrocellulose membrane was purchased from Bio-Rad. The substrates 1 and 2 were prepared from oleoyl chloride and 2-naphthol and 2-naphthylamine, respectively, by a standard procedure (CH2Cl2, pyridine, room temperature, 12 h) and were purified by flash column chromatography on silica gel using ethyl acetate–hexane (1:1) as eluent. Fast Garnet GBC sulfate salt (F8761, dye content >90%) was purchased from Sigma. Micro-scale assay was carried out on a Molecular Devices SPECTRAmax190 96-well plate reader. Fluorescence was measured on a Hitachi F-2000 spectrofluorimeter. A GeneAmp 9700 polymerase chain reaction (PCR) system was purchased from Applied Biosystems. DNA sequencing was carried out on an ABI PRISM 377 DNA sequencer (Applied Biosystems) by using an ABI PRISM BigDye Terminator Cycle Sequencing Ready Reaction Kit.

Error-prone PCR

Random mutations were introduced into the lipase gene by error-prone PCR (Leung et al., 1989Go). The plasmid pLPLsmA coding the wild-type lipase gene was used as a template. Two primers (5'-CCGGCTCGTATAATGTGTGG-3' and 5'-GGAGGATTTTCTTCACGCGA-3') for the amplification of the gene were synthesized to complement 39 bp upstream and 45 bp downstream, respectively, of the coding region (933 bp) for the lipase. The PCR reaction was performed in a total volume of 50 µl containing 0.2 ng/µl pLPLsmA, 0.2 µM of each primer, 0.05 U/µl rTaq polymerase, 0.2 mM dNTP, 1% DMSO, 10% PCR enhancer, 0.2 or 0.3 mM MnCl2 in a standard buffer for PCR [50 mM KCl, 10 mM Tris–HCl (pH 8.3), 1.5 mM MgCl2]. The reactions were run in a GeneAmp 9700 PCR system according to the following cycle program: 96°C for 5 min, then 25 cycles of 96°C for 30 s, 53°C for 30 s and 72°C for 30 s, followed by a final step of 72°C for 10 min. The amplified DNA fragment was purified by phenol–chloroform extraction and ethanol precipitation.

Construction of plasmid library

The gene fragments of mutated lipases and pLPLsmA were digested with NcoI/XhoI and run on a 1% agarose gel. The lipase gene (972 bp) and the vector (8662 bp) were excised and extracted from the gel using a MinElute Gel Extraction Kit (Qiagen). The digested lipase gene (50 fmol) was ligated with the digested vector (20 fmol) for 3 h at 16°C using the DNA Ligation Kit Version 2 (Takara). Ligation mixtures were transformed into E.coli JM109 or DH5{alpha} according to the literature method (Inoue et al., 1990Go). The cells were plated on LB plates containing 20 µg/ml Sm and were incubated until colonies on the plate grew 1–2 mm in diameter. The cells were soaked and collected in 5 ml of LB medium. The plasmid DNA was extracted from the cells using a QIAprep Miniprep Kit (Qiagen) to give a plasmid library of mutated lipases.

Expression of lipase for activity staining

Pseudomonas aeruginosa PAO1162 was grown in LB medium (100 ml) with vigorous shaking at 37°C until log phase (OD660 = 0.4–0.6). The cells were harvested by centrifugation at 10 000 g for 10 min at 4°C, washed twice with 5 mM potassium phosphate buffer (pH 7.0) containing 300 mM sucrose and finally resuspended in 0.01 volume of the same buffer to give electrocompetent cells. An aliquot of the electrocompetent cells (100 µl) was transformed with 10 ng of the plasmid library using a Bio-Rad MicroPulser (0.2 cm electrode, EC2 condition). The cells were incubated in LB medium (900 µl) at 37°C for 1 h with shaking at 160 r.p.m. The mixture was plated on a nitrocellulose membrane placed on rectangular LB agarose plates containing 200 µg/ml Sm by using 5 mm diameter glass beads to disperse the cells uniformly. The cells were incubated overnight at 30°C until colonies on the membrane grew 1 mm in diameter. The colonies were replicated on to a nitrocellulose membrane. The expression of the lipases was induced by putting and incubating the replica membrane at 30°C for 24 h on a 1.5% agarose plate containing 10% LB, 100 µg/ml Sm and 1 mM IPTG.

For the preparation of plates for activity staining, agarose (0.3 g) was added to 20 ml of 100 mM Tris–HCl (pH 8.0) containing 1% Nonidet P-40 (NP-40) and was autoclaved. After cooling the solution to 50°C, the substrate (10 mM of the ester 1 or the amide 2 in DMSO, 200 µl) and Fast Garnet GBC (10 mg/ml in water, 200 µl) were added to the solution. The solution was poured into a rectangular plate and fixed at room temperature. The nitrocellulose membrane with the bacterial colonies expressing the lipase was placed face up on the gel to start the activity staining. After 5 min (for the ester 1) or 1 h (for the amide 2) of incubation at room temperature, the membrane was removed and air-dried for 3 h. The bottom side of the membrane was imaged by a color image scanner (24-bit color, 400 dpi). The image was edited using Adobe Photoshop to distinguish the stained colonies on the membrane as follows. The stained spot was contrasted sharply by setting ‘highlight’ and ‘shadow’ values using the Levels command. A dark spot (an unstained colony) and a red spot (a stained colony) on the image were assigned to ‘highlight’ and ‘shadow’, respectively, by using white and black eyedropper tools to give a clear image of the stained colonies. A threshold of the image was then adjusted using the Brightness/Contrast command to exclude less-stained colonies. By increasing the brightness and contrast level, well-stained colonies were highlighted for selection.

Assay for amide- and ester-hydrolyzing activity in solution

Each well-stained colony was inoculated into LB medium (200 µl) containing 100 µg/ml Sm on each well of a 96-well plate and was cultivated overnight at 30°C. An aliquot (5 µl) of each culture medium was transferred into another LB medium (195 µl) containing 100 µg/ml Sm, 100 µM CaCl2, 0.1% bovine serum albumin and 1 mM IPTG on a 96-well plate. The mixture was incubated at 30°C for 10 h with shaking at 160 r.p.m. to express the lipase. Cells were removed by centrifuging the plate at 900 g for 30 min, followed by transferring the supernatant to another plate. The supernatant was centrifuged again and an aliquot (75 µl) of the supernatant was placed on a deep-well plate (each well of 1100 µl volume) and was mixed with methanol (675 µl) to precipitate the lipase. After incubation at 0°C for 1.5 h, the plate was centrifuged at 900 g for 30 min and the supernatant was discarded. The precipitate was dried in vacuo for 15 min and then dissolved in 100 µl of 100 mM Tris–HCl (pH 7.0) containing 1% NP-40 to give a lipase solution.

The amidase activity of lipase was measured by adding 150 µl of a solution of the amide 2 (10 µM, final concentration) in 100 mM Tris–HCl (pH 7.0) containing 10 µg/ml Fast Garnet GBC (final concentration) and 1% NP-40 to an aliquot (50 µl) of the lipase solution placed in each well of a 96-well assay plate. The absorbance at 530 nm was measured on a 96-well plate reader at 30 min intervals at 25°C for 3 h. The initial velocity was calculated from the slope of each progress curve. One unit of amidase activity was defined as the amount of the lipase that liberated 1 µmol of 2-naphthylamine per minute. The ester-hydrolyzing activity was measured in a similar way by using a diluted lipase solution (100- or 1600-fold) and the ester substrate 1 (50 or 100 µM) in 100 mM Tris–HCl (pH 7.0) containing 10 µg/ml Fast Garnet GBC and 1% NP-40. The absorbance at 530 nm was monitored continuously for 2 min at 25°C on a 96-well plate reader and the initial velocity was calculated from the slope of each progress curve. One unit of esterase activity was defined as the amount of the enzyme that liberated 1 µmol of 2-naphthol per minute. Values were determined in three parallel assays.

Saturation mutagenesis

Saturation mutagenesis at the amino acid positions 207, 213 and 265 was performed by the overlap extension technique (Urban et al., 1997Go). Two DNA fragments having the overlapping ends were generated by PCR using two pairs of the primers (P1/P2 and P3/P4, respectively). The fragments were mixed and amplified by PCR using the primers P1' and P4 to give he lipase gene fragment. The primers P1, P1' and P4 (TCCCCCTCGGCCTGGCCATCGGCC, AAGGTCAACCTGATCGGCCACA and GGAGGATTTTCTTCACGCGAGGGG, respectively) were synthesized to complement 494 bp upstream of the EcoRI site, 215 bp upstream of the EcoRI site and 6 bp downstream of the XhoI site on pLPLsmA, respectively. The overlapping primers (P2 and P3) were synthesized to introduce a mutation at the amino acid positions 207 (GAAGGCGTCGCTCGGATCGAGMNNGTTGGTCAGC and CTCGATCCGAGCGACGCCTTC), 213 (AGCGACGAGGCGCCGAGGAAMNNGTCGCTCGGAT and TTCCTCGGCGCCTCGTCGCT) and 265 (ACGCTGACCGGGCTGGTCTCMNNCAGGCTGGTG and GAGACCAGCCCGGTCAGCGT), respectively, where N is A, T, G or C and M is A or C. The first PCR reactions (P1/P2 and P3/P4) were performed in a total volume of 50 µl containing 0.2 µM primers, 0.2 ng/µl pLPLsmA, 0.05 U/µl KOD-plus, 0.2 mM dNTP, 1% DMSO, 10% PCR enhancer, 1 mM MgCl2 and the buffer. The following PCR program was used: 96°C for 5 min, then 25 cycles of 96°C for 30 s, 60°C for P1/P2 (or 55°C for P3/P4) for 30 s and 68°C for 30 s, followed by a final step of 68°C for 10 min. The second PCR reaction was performed by overlapping the two pairs of the first PCR products, in a total volume of 50 µl containing 1 µl of the first PCR products, 0.2 µM primers (P1' and P4), 0.05 U/µl KOD-plus, 0.2 mM dNTP, 1% DMSO, 10% GIBCO PCR enhancer, 1 mM MgCl2 and the buffer. The PCR program was as follows: 96°C for 5 min, then 25 cycles of 96°C for 30 s, 51°C for 30 s and 68°C for 30 s, followed by a final step of 68°C for 10 min.

Site-directed mutagenesis

The mutations (F207S, A213D and F265L) were combined by site-directed mutagenesis using the overlap extension technique. The P1 and P4 primers were the same as those used in the saturation mutagenesis. The P3 primer (TTCCTCGGCGCCTCGTCGCT) was common to all the experiments. The P2' and P2' primers were AGCGACGAGGCGCCGAGGAAGTCGTCGCTCGGAT and AGCGACGAGGCGCCGAGGAAGGCGTCGCTCGGAT, respectively. The double mutant (F207S/A213D) was prepared by using P1/P2' as primers and the gene from the clone 7B5 (F207S mutant) as template for the upstream region of the gene, and P3/P4 primers and the wild-type gene as template for the downstream region of the gene. The double mutant (A213D/F265L) was prepared by using P1/P2' as primers and the wild-type gene as template for the upstream region of the gene and P3/P4 primers and the gene from the clone 2E7 (F265L mutant) as template for the downstream region of the gene. The double mutant (F207S/F265L) was prepared by using P1/P2' as primers and the gene from the clone 7B5 (F207S mutant) as template for the upstream region of the gene and P3/P4 primers and the gene from the clone 2E7 (F265L mutant) as template for the downstream region of the gene. The triple mutant (F207S/A213D/F265L) was prepared by using P1/P2' primers and the gene from the clone 7B5 (F207S mutant) as template for the upstream region of the gene and P3/P4 primers and the gene from the clone 2E7 (F265L mutant) as template for the downstream region. Each of the upstream and downstream fragments were mixed and amplified by PCR using P1'/P4 primers to give the complete gene fragments for the mutant lipases.

Determination of molecular activity

A lipase solution for active-site titration was prepared from a medium-scale culture (2 ml) according to the same procedure as described in Assay for amide- and ester-hydrolyzing activity in solution. The active-site of the lipase was titrated by using ethyl 4-methylumbelliferyl heptylphosphonate (3) (Fujii et al., 2003Go). An aliquot of a lipase solution was diluted with 100 mM Tris–HCl buffer (pH 7.0) containing 1% NP-40 (990 µl) in a cuvette. A stock solution of 3 (1 mM in DMSO, 10 µl) was added to the solution and the fluorescence was measured (excitation at 363 nm, emission at 445 nm) after the fluorescence increase had leveled off (~40 min). The amount of the active site of lipase was calculated from the amount of released 4-methylumbelliferone (4MU) by calibration with a calibration curve for 4MU after subtracting the background fluorescence before adding 3. Values were determined in three parallel assays. The esterase and amidase activities determined at 100 µM substrate concentration were divided by the titrated concentration of the active lipase to give the molecular activities for the ester 1 and the amide 2, respectively.


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 Abstract
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 Materials and methods
 Results
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 References
 
Random mutagenesis

The gene encoding the lipase was mutated randomly by error-prone PCR by using 0.2–0.3 mM MnCl2. The amplified lipase gene was ligated upstream of the refolding protein lip B (Shibata et al., 1998Go) in the expressing vector of pLPLsmA and was transferred into E.coli to construct a library of mutated lipases. A total of 6 x 104 E.coli transformants were obtained. DNA sequencing of randomly selected several transformants revealed that the mutations were distributed uniformly over the 855 bp lipase gene with a mutational frequency of three to four nucleotides, which corresponds to about one amino acid substitution in the whole lipase gene (Reetz and Jaeger, 1999Go). Under these mutational conditions, 50–70% of the total clones were found to have lost the esterase activity as determined by the activity staining using the ester substrate 1 (see Screening).

Screening

The E.coli transformants were collected and the amplified plasmids were recovered. To express the mutated lipase genes, P.aeruginosa PAO1162 was transformed with the plasmid library by an electroporation method (Sambrook and Russell, 2001Go). A total of 20 000 colonies expressing and secreting the lipase were grown on a nitrocellulose membrane placed on an agar plate containing IPTG. The clones were replicated and subjected to activity staining using the ester 1 or the amide 2 as substrate in the presence of a diazo dye (Fast Garnet GBC) as a coloring agent (Miller and Mackinnon, 1974Go). By this method, the lipase secreted from the cells stayed near the colony and cleaved the ester (or amide) substrate to release 2-naphthol (or 2-naphthylamine), which reacted with Fast Garnet GBC to form a red dye precipitate at the colony. Under these conditions, ~30–50% of the mutants were stained immediately with the ester 1, but the rest of the clones tested negatively, indicating that 50–70% of the clones had lost the esterase activity by introducing one amino acid substitution on average into the whole lipase gene (285 amino acids) by the error-prone PCR.

The library of lipases was screened for the mutants with amidase activity by activity staining using the amide 2 as substrate. Notably, the clones that tested positively with the ester 1 were also colored with the amide 2 by incubating the colonies for a long period (~1 h), indicating that the wild-type lipase had a weak but a distinct activity for the amide (see Discussion). Colonies deeply stained by amide (total 1068 colonies) were picked from the total of 20 000 colonies. Each colony was inoculated into LB medium containing IPTG to express the lipase. The lipase secreted into the culture medium was partially purified by precipitation with methanol. A total of >90% lipase activity in the culture medium was recovered by precipitation as tested by using the wild-type lipase (data not shown). The lipase recovered from the precipitate was assayed for amidase activity by measuring the initial rates (v0, amide) for the hydrolysis of 2. To normalize the amount of lipase, each mutant was also assayed with 1 for esterase activity (v0, ester), which was taken as a measure of the expression level of the lipase. Figure 1A shows the amidase activity (v0, amide) plotted against the esterase activity (v0, ester) of the 1068 clones. Since the proteins recovered from the culture medium of the host cells without the lipase gene hydrolyzed neither ester nor amide (squares), the effect of possible contaminant hydrolases such as esterases and proteases produced by the host was negligible. As reference, the activities of varying concentrations of the purified lipase LPL-312 (triangles) and the wild-type lipase expressed by the host (open circles) were also measured and plotted. The activities of the wild-type lipase were scattered significantly along the abscissa (esterase activity), but were distributed along a slightly downward curved line, indicating that the wild-type lipase exhibited the same relative amidase activity, irrespective of differences in the purity and the expression level. The majority of the mutants (black dots) were located near the line, suggesting that these clones were identical or similar to the wild-type with respect to the relative amidase activity. It is worth noting, however, that six clones (2E7, 2H1, 5B5, 7B5, 8D11 and 10C11) appeared well above the line. These clones were expected to have higher amidase activity than that of the wild-type lipase. Figure 1B shows the ratio of the amidase and esterase activities (v0, amide/v0, ester, A/E ratio) of each clone relative to that for the wild-type, plotted in order of decreasing value. Among a number of clones exhibiting lower A/E ratios than the wild-type, the six clones scored the highest values (1.5–1.7) indicating higher relative amidase activities than that of the wild-type. Further re-examination identified five clones (2E7, 5B5, 7B5, 8D11 and 10C11) that showed 1.5–1.8-fold higher A/E ratios than the wild-type lipase.



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Fig. 1. Amide- and ester-hydrolyzing activities of randomly mutated lipases. (A) Plot of the initial rates for the hydrolysis of the ester 1 (v0, ester) and the amide 2 (v0, amide). The rates were measured photometrically at 530 nm in 100 mM Tris–HCl (pH 7.0) containing 10 µg/ml Fast Garnet GBC, 1% NP-40 and the substrate (100 µM ester 1 or 10 µM amide 2). Black dots, mutated lipase; open circles, wild-type lipase; triangles, purified wild-type lipase (LPL-312); squares, negative control (no lipase gene). (B) Cumulative frequency of v0, amide/v0, ester ratio relative to that for the wild-type lipase (relative A/E ratio). The solid and dashed lines indicate the average value and the standard deviation, respectively, for the wild-type lipase.

 
DNA sequencing of these mutants identified the sites of the mutations (Table I). Each mutant had one or two amino acid substitutions. The clones 8D11 and 10C11 were found to be the same mutant (A213D). The clones 5B5 and 7B5 exhibited almost the same relative A/E ratio (1.7) and contained the same mutation of F207S. Hence the mutation of F207S was also found to contribute to the increase in the A/E ratio, whereas the P96H mutation in the clone 5B5 was considered to be neutral. The sequence analysis also identified the third mutation (F265L) in the clone 2E7.


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Table I. The mutations identified in the selected mutants

 
Saturation mutagenesis

The first screening identified three independent mutations (F207S, A213D and F265L) to improve the A/E ratio. These mutational sites might be substituted by other amino acids for further improvement of the A/E ratio, because a single base mutagenesis by error-prone PCR is accessible to a limited number of amino acid substitutions (5.7 substitutions on average) (Miyazaki and Arnold, 1999Go). In order to examine the substitution by possible 19 amino acids, the positions 207, 213 and 265 were subjected to saturation mutagenesis by introducing a random codon (NNK) in the gene at these sites. Since the possible maximum number of mutation is 32 at the nucleotide level for each site, a library size of 200 clones was enough to cover all the possible mutants for each site. The P.aeruginosa host cells were transformed with a library of expression plasmid and a total 200 colonies were produced for each mutation on a nitrocellulose membrane for the activity staining followed by the solution assay. In contrast to the random mutagenesis by error-prone PCR, almost all the clones were active as tested by the activity staining using 1. From a plot of v0, amide against v0, ester of in total 600 clones (data not shown), we selected 90 clones showing apparently higher amidase activity (Figure 2). Scrutinizing these clones, however, did not yield mutants that exhibited higher amidase activity than that of the parent mutants. We therefore concluded that the amino acid substitutions identified in the original random mutagenesis (F207S, A213D and F265L) were the best mutations at each site.



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Fig. 2. Cumulative frequency of relative A/E ratio (v0, amide/v0, ester) of mutated lipases prepared by saturation mutagenesis. The solid and dotted lines indicate the average value (taken as reference) and the standard deviation, respectively, for the wild-type lipase.

 
Combination of mutations

Since the effect of mutations is often additive, the combination of the mutations of F207S, A213D and F265L was examined for further improvement of the relative A/E ratio. Three double mutants (F207S/A213D, F207S/F265L and A213D/F265L) and the triple mutant (F207S/A213D/F265L) were prepared by site-directed mutagenesis and were assayed for the amidase and esterase activities (Table II). The combination of F207S and A213D was adaptive and increased the A/E ratio by 1.2- and 1.4-fold compared with that for each single mutation F207S and A213D, respectively. However, the addition of the third mutation (F265L) to this double mutant to give the triple mutant resulted in decrease in the A/E ratio by 20%. The effect of F265L on the single mutants (F207S and A213D) seemed to be neutral, because the A/E ratios of the double mutants (F207S/F265L and A213D/F265L) were almost the same as those of the corresponding parental mutants.


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Table II. Activities of mutants and wild-type lipasea

 
Determination of molecular activity

The amino acid substitutions of F207S, A213D and F265L were found to improve the A/E ratio compared with the wild-type, but it is not clear whether the increase in the A/E ratios was due either to the increase in the amidase activity or to the decrease in the activity for the ester or to the combination of both. In addition, one of our goals was to increase the specific activity for the amide 2. It was therefore highly desirable to know the ‘absolute’ activity of each mutant lipase for the ester 1 and the amide 2. For this purpose, we carried out an active-site titration of the lipase by using the phosphonate 3 to determine the molar concentration of the active lipase (Fujii et al., 2003Go) and calculated the molecular activities for the hydrolysis of the ester and the amide. The results are summarized in Table II. Importantly, the molecular activities shown here are approximately equal to kcat/Km[S], where [S] was 10–100 µM in the standard assay (see Materials and methods), because a plot of v0 against the concentration of each substrate exhibited no saturation, giving a straight line up to 100 µM (data not shown). The molecular activity for the amide 2 increased uniformly for all the single mutants. However, the molecular activities for the ester 1 decreased by ~28% for the mutants F207S and F265L, whereas the A213D mutant had the same esterase activity as the wild-type, indicating that the mutation A213D increased the amidase activity without decreasing the esterase activity. Among three single mutants, the A213D mutant exhibited the highest molecular activity for the amide, whereas F207S was the best mutant with respect to the relative A/E ratio owing to some decrease in the esterase activity.

It is worth noting that the combination of F207S and A213D was additive with respect to the molecular activities for both the amide and the ester. Thus, the molecular activity of the F207S/A213D mutant for the amide 2 (1.1 min–1) was higher than that of either the parental single mutants and was twice that of the wild-type, whereas the F207S/A213D mutant stood between the parents with respect to the activity for the ester 1. The effect of F265L on the other mutations was variable. The addition of a F265L mutation to the other single mutants resulted in a slight increase (F207S/F265L) or a slight decrease (A213D/F265L) in the molecular activities for both the amide and ester. However, the addition of this mutation into the double mutant (F207S/A213D) decreased the amidase activity by 20%, whereas the esterase activity was constant, indicating that the effect of F265L was negative for the double mutant with respect to the amidase activity. As a result, the double mutant F207S/A213D was the best mutant in terms of both the molecular activity for the amide and the relative A/E ratio.

Structural analysis

This lipase belongs to the Pseudomonas group I lipases (Arpigny and Jaeger, 1999Go) and the amino acid sequence is 99% homologous with that of a lipase from P.aeruginosa PAO1 (Jaeger et al., 1994Go), for which a crystal structure has been determined (Nardini et al., 2000Go). Hence the structure of the lipase PAO1 was used as a model to examine the mutational sites found in our lipase from a structural point of view. Figure 3 shows the amino acid sequence of the wild-type lipase. All the mutations were found near the residues responsible for the coordination to a calcium ion (D209, D253, Q257 and L261) or the residues interacting with the calcium ion via water molecules (T205, S211 and D212) (Nardini et al., 2000Go).



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Fig. 3. Amino acid sequence of the lipase from P.aeruginosa TE3285. The reversal letters indicate the mutated positions (F207, A213 and F265). The closed triangle, closed circle and open triangle symbols indicate the residues related to the catalytic triad, the oxyanion hole and the calcium binding site, respectively (Nardini et al., 2000Go).

 
Figure 4 shows the location of F207, A213 and F265 in the crystal structure of the lipase from P.aeruginosa PAO1 (Nardini et al., 2000Go). The residues F207 and F265 are located on the surface loop, whereas the residue A213 is on helix {alpha}8 and is also near the surface. These residues are not interacting directly, but are not far from each other: the C{alpha} distance between F207 and A213 is 8 Å and that between F207 and F265 is 13 Å. As expected from the amino acid sequence, these residues are located near the calcium binding site, but are far from the active site of the lipase. Since the calcium binding site is located ~15 Å away from the catalytic nucleophile S82, the mutations F207S, A213D and F265L are not likely to affect the amidase/esterase activities by interacting directly with the catalytic triad or the oxyanion hole (M16 and H83).



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Fig. 4. Crystal structure of the lipase from P.aeruginosa PAO1 (Nardini et al., 2000Go). The side chains of the mutated positions (F207, A213 and F265), the catalytic triad (S82, D229, H251) and the oxyanion hole (M16, H83) are shown in stick representation. The sphere represents the calcium ion.

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Lipases are extremely useful chiral acyl transfer catalysts for the biotransformation of esters, alcohols and carboxylic acids. Optically active amines are also accessible by lipase catalysis (Kazlauskas and Bornscheuer, 1998Go), but the reaction is practically limited to the acylation of amines (Kazlauskas and Bornscheuer, 1998Go; Henke and Bornscheuer, 2003Go), a process identical with the hydrolysis of esters from a mechanistic point of view. Owing to resonance stabilization, the hydrolysis of amides is chemically more demanding than that of esters (Bender et al., 1962Go; Zerner et al., 1964Go; Bender and Kézdy, 1965Go) and is usually not amenable to the catalysis by lipases. A lipase from Candida antarctica (CAL-B) has an exceptionally high amidase activity (Wagegg et al., 1998Go; Duarte et al., 2000Go). However, the primary sequence of this lipase has no significant sequence homology with that of any other known lipases and is more like an esterase than a lipase, since the CAL-B lipase has a small lid covering the active site and shows little interfacial activation (Uppenberg et al., 1994Go). Hence few reports evaluating the amide-hydrolyzing activity of the ‘true’ lipases have appeared (Henke and Bornscheuer, 2003Go).

Directed evolution is a powerful technique in altering the properties of enzymes such as thermal stability, substrate specificity and enantioselectivity (Cherry and Fidantsef, 2003Go; Hult and Berglund, 2003Go; Reetz, 2004Go), but altering the reaction specificity is more challenging, because a large evolutionary gap usually lies between the enzymes that catalyze different reactions. The evolutionary gap, however, might be overcome if the reactions are mechanistically related, as nature has used common mechanistic features of reactions to diversify enzymes to catalyze different reactions (O'Brien and Herschlag, 1999Go). The hydrolysis of amides by lipases is such a reaction and is amenable to directed evolution, because the hydrolysis of amides is mechanistically similar to that of esters (Bennet and Brown, 1998Go). In addition, serine proteases, which are analogous to lipases with respect to the active site structure and the reaction mechanism, catalyze the hydrolysis of both amides and esters. In this paper, we have demonstrated that a lipase from P.aeruginosa had a low level of amidase activity that could be enhanced by a combinatorial approach.

The wild-type lipase from P.aeruginosa TE3285 was found to have a weak, but detectable, level of activity for the hydrolysis of the naphthylamide 2 (1/35 000 of the activity for the ester 1, Table II). A standard photometric assay using oleoyl 4-nitroanilide as substrate (pH 7.0, 410 nm) failed to detect any amidase activity of this lipase, probably because the chromophoric detection of 4-nitroaniline was not sensitive enough to find the low level of activity. Knowing that the diazo-coupled detection of 2-naphthylamine was effective in measuring the amidase activity for both the colony and the solution assays, the lipase was subjected to directed evolution for improved hydrolytic activity for the amide 2 compared with that for the structurally analogous ester 1. After a single round of random mutagenesis, the first screening by activity staining of the colonies followed by a solution assay for improved amidase/esterase activity ratios (A/E ratio) identified five clones that exhibited improved relative amidase activities (Figure 1). The sequence analysis of these clones found that three mutations (F207S, A213D, F265L) affected the amidase/esterase activities of the lipase. The double and the triple mutants were also prepared by site-directed mutagenesis to see the effect of combination of these mutations.

Mutational and evolutional studies of lipases often encounter difficulties in determining the amount of active lipase expressed in culture media (van Kampen and Egmond, 2000Go; Suen et al., 2004Go). We initially used the esterase activity for 1 to estimate the concentration of each mutant lipase. However, this is valid only when the specific activity for 1 is constant for all the mutants. Of course, this premise is not guaranteed, because mutations may affect the intrinsic esterase activity of the lipase, as well as the amidase activity. In addition, the effect of mutations on the ‘absolute’ activities for the ester 1 and the amide 2 was also our interest. We therefore quantified the active lipase by active-site titration using the fluorescent phosphonate 3 (Fujii et al., 2003Go). This method was effective in determining the molar concentration of the active lipase in a small-scale culture medium, thereby allowing the calculation of molecular activities of each mutant for the ester 1 and the amide 2 without purification of the lipase (Table II). The molecular activities determined for the ester 1 and the amide 2 enabled direct comparison of the activities among the mutants to reveal some interesting facets as to the effect of mutations on the esterase and amidase activities. First, the amidase activity of each mutant (F207S, A213D and F265L) was increased uniformly, although the mutants were screened for increased A/E ratios. No mutants increased the esterase activity compared with the wild-type. This is probably because the first screening (colony assay) was based on the apparent hydrolytic activity for the amide 2 and the preselected mutants (1068 clones) were then screened for increased A/E ratios. It is interesting, however, that the first screening successfully enriched the mutants with increased amidase activity, because the expression level of the mutants was not considered in the colony assay, where ‘well-stained’ colonies were selected irrespective of the significantly varied expression level (see Figure 1A). Second, each mutation (F207S, A213D and F265L) has a different effect on the amidase and the esterase activities. For example, the mutation A213D increased the amidase activity without affecting the activity for the ester, whereas the mutations F207S and F265L increased the activity for the amide, but with concomitant loss of the esterase activity. We anticipated initially that the amidase activity of the lipase would be parallel to its esterase activity, because the ester 1 and the amide 2 are structurally homologous and are hydrolyzed in the same active site by a similar mechanism. However, this was not the case at least for these mutants. The amidase activities were increased without increasing the activity for the ester, suggesting that both activities can vary independently. This result, however, does not rule out the possibility of finding mutants that increase the amidase activity by concomitant increase in the activity for the ester, because the mutations F207S, A213D and F265L were found in the clones that had been selected under a selective pressure for generating different reaction specificities (increased A/E ratios). Other clones such as those found in the upper right region of a plot of v0, amide vs. v0, ester (Figure 1A) might show such a parallelism between the esterase and amidase activities. Finally, the mutation A213D was most significant in increasing the molecular activity for the amide, whereas the other mutations had a limited effect on the amidase activity when combined with A213D.

A structural model based on the crystal structure of P.aeruginosa PAO1 (Nardini et al., 2000Go) revealed that the mutational sites F207, A213 and F265 were located on the surface near the calcium binding site (Figure 4). The binding site for a calcium ion is ~15 Å away from the catalytic serine (S82), but the calcium ion bridges helix {alpha}8 to the catalytic histidine (H251), thereby contributing to keeping H251 at the correct position in the active site (Nardini et al., 2000Go). It should be noted that the most important mutation A213D was found on helix {alpha}8, which forms part of the wall of the active site cleft. This finding suggests that the mutation A213D may influence the orientation of the catalytic H251 through the calcium ion bridge. The importance of catalytic histidine as a general acid–base catalyst is highlighted in the acylation–deacylation double displacement mechanism of serine hydrolases for amide hydrolysis (Wharton, 1998Go; Hedstrom, 2002Go). In the acylation step, the histidine abstracts a proton from the catalytic serine to increase its nucleophilicity and donates the proton to the leaving group nitrogen to facilitate the breakdown of the tetrahedral intermediate to form the acyl enzyme intermediate. In serine proteases, the formation of the acyl enzyme intermediate (acylation step) is usually the rate-limiting step for amide substrates (Zerner et al., 1964Go; Whitaker and Bender, 1965Go; Walsh, 1979Go; Wharton, 1998Go; Hedstrom, 2002Go). If the lipase hydrolyzes the amide 2 via an acyl enzyme intermediate, the molecular activities shown here are dependent on the acylation rate (k2), because they are approximately equal to kcat/Km[S] under the standard assay conditions ([S] << Km, see Results) (Walsh, 1979Go). Therefore, the increase in the amidase activity is attributed to the increase in the acylation rate and the mutation A213D may facilitate the acylation step by influencing the proton-abstracting and/or -donating ability of the catalytic H251. The participation of the catalytic H251 in leaving group protonation is demonstrated by the X-ray crystal structure of a lipase from P.aeruginosa PAO1 complexed with a phosphonate transition-state analog inhibitor covalently bound to the catalytic serine (Nardini et al., 2000Go). A hydrogen bond between the catalytic histidine and the leaving group oxygen is also shown to be critical in enantioselective hydrolysis of a racemic ester by a lipase, where the slow-reacting enantiomer lacks a hydrogen bond between N{varepsilon}2 of the imidazole ring of the catalytic histidine and the leaving group oxygen (Cygler et al., 1994Go). Interestingly, a few hydrolases have been reported to show catalytic properties that do not conform to the general reactivity of amides and esters (Wu et al., 1995Go; Patricelli and Cravatt, 1999Go). A serine enzyme fatty acid amide hydrolase, for example, hydrolyzes oleamide, oleoyl methylamide and oleoyl methyl ester with equivalent catalytic efficiencies (Patricelli and Cravatt, 1999Go). The unusual catalytic properties of this hydrolase are explained by a strong degree of leaving group protonation by a key lysine residue acting as a general acid–base catalyst. The relative importance of general acid-catalyzed leaving group protonation for amide and ester hydrolysis was emphasized by Fersht (1971)Go. Hence it seems reasonable to attribute the increased amidase activity of the lipase to the improved leaving group protonation.

In conclusion, a sensitive screening system for improving amide-hydrolyzing activity of lipases was constructed by combining diazo-coupled activity staining and active-site titration. The amidase activity of a lipase from P.aeruginosa was shown to improve by ~2-fold after a single round of random mutagenesis and the combination of mutations. This study lays the basis for further improvement of amidase activity of lipases that could give rise to novel biocatalysts for amide hydrolysis. In the future, the analysis of mutant lipases with significantly improved amidase activity should broaden our understanding of the underlying principles for enzymatic hydrolysis of esters and amides.


    Notes
 
2 Present address: Enzyme Laboratory, National Food Research Institute, 2–1–12 Kannondai, Tsukuba, Ibaraki, 305-8642, Japan Back


    Acknowledgments
 
This study was supported in part by a Grant-in-Aid for Scientific Research in Priority Areas from the Ministry of Education, Culture, Sports, Science and Technology, Japan (contract 13125203) for Professor Nobuyoshi Esaki at this Institute. We thank him for his generous financial support. We also thank Yuji Utsunomiya for his technical assistance with the preparation of the phosphonate 3.


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Received December 14, 2004; revised January 13, 2005; accepted January 20, 2005.

Edited by Mirek Cygler





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