Characterizing LipR from Pseudomonas sp. R0-14 and Applying in Enrichment of Polyunsaturated Fatty Acids from Algal Oil

Abstract

In this study, Pseudomonas R0-14, which was isolated from Arctic soil samples, showed a clear halo when grown on M9 medium agarose plates containing olive oil-rhodamine B as substrate, suggesting that it expressed putative lipase(s). A putative lipase gene, lipR, was cloned from R0-14 by genome walking and Touchdown PCR. lipR encodes a 562-amino-acid polypeptide showing a typical α/β hydrolase structure with a catalytic triad consisting of Ser₁₅₃-Asp₂₀₂-His₂₆₀ and one α-helical lid (residues 103-113). A phylogenetic analysis revealed that LipR belongs to the lipase subfamily I.3. LipR was successfully expressed in Escherichia coli, purified, and biochemically characterized. Recombinant LipR exhibited its maximum activity towards p-nitrophenyl butyrate at pH 8.5 and 60℃ with a K_m of 0.37 mM and a k_cat of 6.42 s^-1. It retained over 90% of its original activity after incubation at 50℃ for 12 h. In addition, LipR was activated by Ca²⁺, Mg²⁺, Ba²⁺, and Sr²⁺, while strongly inhibited by Cu²⁺, Zn²⁺, Mn²⁺, and ethylenediaminetetraacetic acid. Moreover, it showed a certain tolerance to organic solvents, including acetonitrile, isopropanol, acetone, methanol, and tert-butanol. When algal oil was hydrolyzed by LipR for 24 h, there was an enrichment of n-3 long-chain polyunsaturated fatty acids, including eicosapentaenoic acid (1.22%, 1.65-fold), docosapentaenoic acid (21.24%, 2.04-fold), and docosahexaenoic acid (36.98%, 1.33-fold), and even a certain amount of diacylglycerols was also produced. As a result, LipR has great prospect in industrial applications, especially in food and/or cosmetics applications.

Introduction

Enzyme catalysis is superior to chemical catalysis owing to its greater selectivity and environmental friendliness, as well as its lower energy consumption [11]. Lipases (triacylglycerol acylhydrolase, E.C. 3.1.1.3), a group of important industrial enzymes, can catalyze hydrolysis, esterification, and transesterification reactions on a wide range of substrates [12]. Among them, microbial lipases are prominently applied in the food, detergent, paper-making, biodiesel, fine chemicals, and pharmaceutical industries [39].

Microbial lipases have been extracted from bacteria, yeast, and fungi. Although fungal and yeast lipases are very useful and can be produced in abundance, they are mostly unstable in harsh conditions, such as elevated temperatures, organic solvents, and/or detergents that are used in various industrial applications. In contrast, bacterial lipases exhibit higher thermostability and tolerance to denaturing reagents and/or organic solvents [31]. In addition, they can be easily produced via genetic manipulation, and are usually characterized by some unique features, which expand their application in industry, the biomedical sciences, and academic research [2]. To date, many bacterial lipases with excellent properties have been identified. However, the widest exploration is being focused on the genus Pseudomonas [23,38]. Increasingly versatile and highly tolerant Pseudomonas lipases have been discovered and characterized, and some of them are being applied in various industries [38]. Additionally, because of the rapid development of the modern biochemical industry and significant research in bioengineering, the existing and/or commercial lipases cannot meet increasing demands [23], and thus, more lipases with unique characteristics are badly needed. Therefore, investigations for isolating such lipases from bacteria, especially from Pseudomonas, are urgent and are of great significance.

Moreover, omega-3 (n-3) long-chain polyunsaturated fatty acids (n-3 LCPUFAs), such as eicosapentaenoic acid (EPA), docosapentaenoic acid (DPA), and docosahexaenoic acid (DHA), have been used to prevent and treat cancer, arteriosclerosis, inflammation, and hyperlipemia [9], and they even exhibit neuroprotective properties and therefore represent a potential treatment for a variety of neurodegenerative and neurological disorders [7]. However, they always exist in oils with low contents, and they cannot meet market and medical requirements. Thus, it is very important to enrich the content of n-3 LCPUFAs in oils.

Lipases generally have higher hydrolytic activity towards short- and medium-chain fatty acids than towards long-chain fatty acids, especially towards these n-3 LCPUFAs [6]. Microbial lipases from Aspergillus niger [22], Geotrichum candidum [36], and Yarrowia lipolytica [37] were used to hydrolyze fish oil or algal oil to enrich these n-3 LCPUFAs. Nevertheless, they are easily inactivated during production, and it is assumed that their counterparts from bacteria may have better performance and more potential prospect. Oils, including both n-3 LCPUFAs and diacylglycerols (DAGs), have significant health benefits and have become increasingly popular [9,20]. Therefore, it is of great importance to identify and characterize more microbial lipases from nature to meet various demands [16].

In this study, Pseudomonas sp. R0-14 was selected from Arctic soil samples according to its ability to exhibit clear halos when grown on M9 medium agar plates containing olive oil and rhodamine B. Thus, strain R0-14 was a candidate for the cloning of a putative lipase(s). Subsequently, a whole lipase gene, lipR, was successfully cloned via touchdown PCR and genome walking from the genome DNA of Pseudomonas sp. R0-14 and expressed in Escherichia coli. Then, its enzymatic properties were characterized. In addition, its hydrolysis of algal oil was tested to examine its unique characteristics.

 

Materials and Methods

Bacterial Strains, Plasmids, and Chemicals

Pseudomonas sp. strain R0-14 was isolated from Arctic soil samples and preserved in the China Center for Type Culture Collection (CCTCC), College of Life Sciences of Wuhan University, and its strain preservation number is CCTCC AB 2012778. Strain R0-14 was grown at 28℃ in Luria–Bertani (LB) broth or on agar plates containing 100 μg/ml ampicillin. The E. coli strains DH5α and BL21 (DE3) (Novagen, Darmstadt, Germany) were maintained at 37℃ in LB broth or on agar plates for recombinant plasmid amplification and protein heterologous overexpression, respectively. Ampicillin was added to the medium to screen the transformants. The vectors T-Vector pMD19 (Simple) (Takara, Otsu, Japan) and pET-22b (+) (Novagen, Madison, WI, USA) were used for gene cloning and expression, respectively. Kits for genome extraction, plasmid extraction, and gel extraction were purchased from Omega Bio-tek (Norcross, GA, USA). Genome walking kits, restriction endonucleases, T4 DNA ligase, and Taq DNA polymerase were all purchased from Takara. To validate the accuracy of gene insertions, DNA sequencing was performed by Invitrogen Biotechnology Company (Carlsbad, CA, USA). Substrate p-nitrophenyl (p-NP) esters were purchased from Sigma-Aldrich (USA). All other chemicals used were of analytical grade and were commercially available from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China).

Cloning of lipR via Touchdown PCR and Genome Walking

All primers used are listed in Table 1. A partial lipase sequence was amplified from Pseudomonas sp. R0-14 genomic DNA via Touchdown PCR [39] with degenerate primers T5 and T3 designed using CODEHOP [28]. The PCR conditions were 94℃ for 5 min; 20 cycles of 94℃ for 30 sec, 60℃ to 55℃ with a decrease of 0.5℃ after each cycle, and 72℃ for 50 sec, followed by a final extension at 72℃ for 10 min. The amplified PCR product was cloned into pMD19 (Simple) and sequenced.

Table 1.N represents A, C, G, or T; Y identifies C or T; R represents A or G; W identifies A or T. NdeI and XhoI restriction sites in primers are underlined, respectively.

To obtain the upstream and downstream sequences of the partial lipase gene, a genome walking PCR was performed using a genome walking kit according to the manufacturer’s instructions. Sequence assembly was analyzed via BioEdit software. Sequence alignments of DNA and protein sequences were performed using BLASTN and BLASTP, respectively (http://www.ncbi.nlm.nih.gov/BLAST/). A multiple sequence alignment was conducted using Clustal W2 (http://www.ebi.ac.uk/Tools/msa/clustalw2/) [30] and presented using ESPript 2.2 (http://espript.ibcp.fr/ESPript/ESPript/). A phylogenetic analysis was conducted with MEGA 5.0 software using the neighbor-joining method. A bootstrap analysis with 1,000 replicates was used to estimate the reliability of the tree [10]. A three-dimensional structure of the target protein was constructed by SWISS-MODEL (http://swissmodel.expasy.org/) and presented using CCP4MG 2.5 [24]. A signal peptide was predicted using the SignalP 4.1 server (http://www.cbs.dtu.dk/services /SignalP/) [31].

Nucleotide Sequence Accession Number

The lipR nucleotide sequence and amino acid sequence have been submitted to the GenBank database under the accession numbers KF620115 and AHB29479, respectively.

Expression and Purification of Recombinant LipR

The PCR product amplified by primers lipFm and lipRm was inserted into pET-22b (+) that was digested with NdeI and XhoI, such that LipR had a carboxyl terminal six-histidine (his)-tag. The recombinant plasmid pET-22b-lipR was transformed into E. coli BL21 (DE3) cells. Transformed cells from a single colony were grown overnight at 37℃ in a shake flask containing 5 ml of LB broth supplemented with 100 μg/ml ampicillin. Then, the overnight culture was diluted 1:100 into one liter of fresh LB medium containing ampicillin and grown aerobically at 37℃. Recombinant LipR expression was induced with 0.1 mM isopropyl β-ᴅ-1-thiogalactopyranoside (IPTG) at 16℃ for 20 h after the optical density at 600 nm reached 0.6.

LipR was purified from E. coli cell extracts according to a previously described method for inclusion body solubilization [1] with minor modifications. Eight liters of cells was harvested by centrifugation at 8.000 ×g for 10 min at 4℃ and resuspended in lysis buffer (20 mM Tris–HCl buffer, 0.5 M NaCl, pH 8.0), and then disrupted by a One Shot Cell Disrupter (Constant Systems, Daventry, UK). Insoluble material was isolated by centrifugation at 8,000 ×g for 30 min at 4℃ and then dissolved in 20 mM Tris–HCl (pH 8.0) containing 1 mM ethylenediaminetetraacetic (EDTA), 5% glycerol, 10 mM dithiothreitol (DTT), and 8 M urea. The resultant supernatant was collected by centrifugation at 8,000 ×g for 30 min at 4℃ and then transferred to a Ni-NTA affinity chromatography column (GE Healthcare, Pittsburgh, PA, USA) that had been previously equilibrated with washing buffer (20 mM Tris–HCl, pH 8.0, 0.5M NaCl). Then, the target recombinant enzymes were eluted using an imidazole concentration gradient (0, 30, 60, 100, and 200 mM) in washing buffer. Thirty milliliter of NTA-200 eluent was dialyzed in 20 mM Tris–HCl buffer (pH 8.0) containing 5 mM CaCl2 and then concentrated to 7 ml by ultrafiltration with a 30 kDa molecular mass cutoff filter, and finally analyzed by 12% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and stained with Coomassie brilliant blue R-250. Unstained and pre-stained protein molecular weight markers (Fermentas, SM0431 and SM0671, Thermo Fisher Scientific, Waltham, MA, USA) were used as references. Protein concentrations were measured spectrophotometrically according to the method of Bradford using bovine serum albumin as a standard [5]. For western blot analysis, proteins were transferred onto a polyvinylidene fluoride membrane and immunologically detected with mouse monoclonal IgG2b anti-His antibodies (Tiangen, Beijing, China) as described by the manufacturer.

Biochemical Characterization of LipR

Substrate specificity was analyzed using different p-NP-fatty acyl esters as substrates. In a standard assay, the total 1 ml reaction system contained 940 μl of Tris–HCl buffer (50 mM, pH 8.0), 10 μl of p-NP ester (100 mM), 40 μl of ethanol, and 10 μl of the diluted enzyme solution [39]. The blank contained the same components except the enzyme solution. All experiments were performed in triplicate. One unit of lipase activity (U) was defined as the amount of enzyme that liberated 1 μmol of p-NP per minute under the assay conditions. In addition, the activity of the purified lipase was also measured by titration of free fatty acids (FFAs) released by the hydrolysis of olive oil using the pH-stat method [19]. The measurements were conducted in triplicates. One unit was defined as the amount of enzyme liberating 1 μmol of fatty acid per minute.

The optimal pH was determined by examining the enzyme activity at 60℃ in different buffers with pH values ranging from 4.0 to 10.0: citrate-phosphate buffer (pH 4.0–6.5), Tris–HCl buffer (pH 7.0–8.5), and glycine-NaOH buffer (pH 9.0–10.0). The optimal temperature was determined by varying the temperature from 0 to 100℃ with an interval of 5℃ (except between 0℃ and 10℃) in Tris–HCl buffer (pH 8.5). Enzyme stability was also examined after incubation at different temperatures (50℃, 60℃, 70℃, and 80℃) for 12 h. The residual lipase activity was tested under the standard assay conditions. The kinetic parameters of LipR were tested in 50 mM Tris–HCl (pH 8.5) at 60℃ using p-NP butyrate at different concentrations (0.01, 0.02, 0.05, 0.1, 0.2, 0.4, 0.8, 1, and 2 mM). The Km and Vmax were calculated from the Lineweaver-Burk plot using Microsoft Excel software (Microsoft Corporation, Redmond, WA, USA).

The effects of additives on LipR were also examined. The reactions contained various metal ions (listed in Table 3), EDTA, and the inhibitors phenylmethanesulfonyl fluoride (PMSF), DTT, and β-mercaptoethanol (β-ME) at final concentrations of 1 or 10 mM. Reactions containing 15% (v/v; i.e., mixing 0.45 ml of organic solvent in 3 ml of the enzyme solution) or 30% (v/v) organic solvents were incubated at 60℃ with shaking at 150 rpm in an orbital shaker for 2 h. Reactions containing 0.05% or 0.1% (w/v) of commercial detergents were incubated at 60℃ for 30 min. After these incubations, residual lipase activities were assayed in 50 mM Tris–HCl (pH 8.5) at 60℃ for 10 min. The activity of LipR determined in the buffer with no additives was set as 100%. Values are means with standard errors from three independent experiments.

Hydrolysis of Algal Oil

The ability of LipR to hydrolyze algal oil was determined. The total hydrolysis reaction system contained 2 ml of 50 mM Tris–HCl (pH 8.0), 1 g of algal oil, and 2 ml of the purified LipR solution (2 mg/ml in 50 mM Tris–HCl, pH 8.0). After nitrogen purging for 1 min, the system was reacted at 60℃ with shaking speed at 200 rpm for each 12 h and 24 h, respectively. Another system, which consisted of the above components but with Tris–HCl buffer instead of enzyme solution, served as the control. The oil phase of the reacted mixtures was analyzed by thin-layer chromatography (TLC). The developing solvents of TLC included hexane, diethyl ether, and acetic acid (85:15:1 (v/v/v)). Additionally, the methyl esters of the raw algal oil and fatty acids in the glycerides were identified and quantified using an Agilent 7890A/5975C gas chromatograph (GC) and a mass spectrometer(MS) equipped with a capillary column (DB-WAX, 30 m × 250 μm × 0.25 μm).

The parameters of the GC-MS program were as follows: the oven temperature was increased from 150 to 250℃ at a rate of 10℃/min, and then maintained at 250℃ for 10 min; the temperatures of the injector and connector were set at 250℃ and 260℃, respectively. Split sampling, with an injection volume of 1 μl and a split ratio of 50:1, was employed. The MS program was operated as follows: the solvent delay time was 3 min with a He flow rate of 1 ml/min; the temperatures of the MS source and MS quad were 230℃ and 150℃, respectively. The range of detectable molecular masses was 30–500 Da.

Methyl heptadecanoate was used as the internal standard. The relative contents of each fatty acid in the glycerides were calculated with the following equation:

where Asample is the peak area of each fatty acid in the glyceride sample; Atotal is the total peak area of all fatty acids in the glyceride sample and the internal standard; and Ainternal is the peak area of the internal standard.

 

Results

Gene Cloning and Sequence Analysis

Degenerate primers T5 and T3 were designed according to the conserved NVLNIGYE (Fig. 1, residues 193–200) and NTFLFSGAF motifs (Fig. 1, residues 488–496), respectively. With these degenerate primers, a 915 bp DNA fragment was obtained via touchdown PCR. The BLAST analyses confirmed that this nucleotide fragment is homologous to reported lipase genes of subfamily I.3. Moreover, the 1,000 bp upstream and 750 bp downstream sequences were amplified via genome walking. A 1,689 bp lipase open reading frame (designated as lipR) was identified after assembling the three DNA fragments by using BioEdit. Subsequently, the entire lipase gene lipR was amplified from its genome by primers lipR-F and lipR-R to further confirm its splicing nucleotide sequence.

Fig. 1.Multiple sequence alignment between LipR and other family I.3 Lipases.

The lipR gene encodes a 562-amino-acid polypeptide with a theoretical molecular mass of 58.91 kDa and an isoelectric point (pI) of 4.59. LipR showed sequence similarities with lipases from Pseudomonas mandelii JR-1 (GenBank: WP_010460388, 94%) [14], an uncultured bacterium (GenBank: AAP76488, 85%) [33], Pseudomonas fluorescens Pf0-1 (GenBank: WP_011334098, 83%) (http://genome.jgi-psf.org/draft_microbes/psefl/psefl.home.html), Pseudomonas sp. lip35 (GenBank: ABY86751, 76%) [34], P. fluorescens Pf-5 (GenBank: WP_015635756, 65%) [24], and Pseudomonas sp. MIS38 (PDB: 2z8x_A, 56%) [1]. Phylogenetic analysis revealed that LipR belonged to lipase/esterase family I.3 (Fig. 2). A multiple sequence alignment showed that LipR contains a catalytic triad composed of Ser153, Asp202, and His260, with Ser153 located in a conserved GHSLG motif (Fig. 1, residues 151–155). LipR does not contain a cysteine residue. Like other lipases [1,15,18] in subfamily I.3, LipR also contains a C-terminal targeting signal region consisting of six tandem repeats of the nine-residue motif GGXGXDXUX (U, hydrophobic amino acids; X, any amino acid residue; Fig. 1, residues 319–336; 329-346; 474–491), an 18-residue amphipathic α-helix (Fig. 1, residues 530–547), and an extreme C-terminal motif consisting of a negatively charged residue followed by four hydrophobic residues (Fig. 1, residues 557–560).

Fig. 2.Phylogenetic analysis of LipR and other closely related lipases.

Expression and Purification of LipR

Recombinant LipR was purified from the supernatants of cell lysates by affinity chromatography using a Ni-NTA column, dialyzed, and concentrated. Because of the low solubility of subfamily I.3 lipases in E. coli [1], the study selected 16℃ for 20 h as the IPTG induction condition [35]. As shown in Table 2, this purification protocol for LipR resulted in a 48.6-fold purification with a recovery rate of 6.55%, and it yielded 252 mg of LipR from each 8 L of cell culture. According to SDS-PAGE (Fig. 3A) and Western blotting (Fig. 3B), purified LipR was detected as a single band with a molecular mass of ~60 kDa, which coincides well with its theoretical value.

Table 2.Lipase activity was determined with p-nitrophenyl butyrate as substrate.

Fig. 3.SDS-PAGE and Western blot analysis of LipR.

Substrate Specificity and Effects of pH and Temperature on LipR activity

As shown in Fig. 4A, LipR had the ability to hydrolyze p-NP esters with acyl fatty acid chain lengths ranging from C2 to C14, and it showed the maximum lipase activity towards p-NP butyrate (79.70 U/mg) and olive oil (26.82 U/mg). Compared with its hydrolysis of p-NP butyrate, the relative activities of LipR towards p-NP caprylate (C8) and p-NP caprate (C10) were 53.6% and 31.4%, respectively. The substrate profile of LipR is similar to those of PML (PDB: 2z8x) from Pseudomonas sp. MIS38 [1] and BPL1-3 (Korea Polar Research Institute: No. 22952, No. 22850, No. 23867) from Bacillus pumilus [3]. In contrast, most other lipases from Pseudomonas and Bacillus were reported to function better towards the esters of short and/or medium-chain fatty acids compared with those of longer-chain fatty acids [15,23].

Fig. 4.Characterization of LipR.

Recombinant LipR exhibited its maximum activity towards p-NP butyrate at pH 8.5 and 60℃ (Figs. 4B and 4C) with a Km of 0.3 7 mM and a kcat. of 6.42 s–1. LipR retained more than 50% of its maximal activity in a pH range from 7.0 to 9.5 (Fig. 4B), indicating that it is well adapted to a wide pH range. As shown in Fig. 4C, LipR retained more than 60% of its maximal activity at temperatures ranging from 40℃ to 80℃, and it retained over 30% of its activity between 20℃ and 90℃, which suggests that LipR is active over a wide temperature range. After incubation at 50℃ and 60℃ for 12 h, the residual LipR activities were 90% and 80%, respectively, of the original activity (Fig. 4D). Additionally, LipR retained over 60% of its original activity after incubation for 12 h at 70℃ or 1 h at 80℃, and it retained 30% of its original activity after incubation for 3 h at 80℃.

Effects of Metal Ions, Inhibitors, Organic Solvents, and Detergents on LipR Activity

The effects of different metal ions and inhibitors on LipR activity are listed in Table 3. The activity of the control (100%) was 79.70 U/mg. At a final additive concentration of 10 mM, LipR activity was obviously enhanced by Ca2+ (195.03%), Mg2+ (163.69%), Ba2+ (152.45%), and Sr2+ (170.18%), respectively; however, it was strongly inhibited by Mn2+ (42.25%), Ni2+ (57.99%), Cu2+ (21.21%), Zn2+ (22%), and EDTA (58.61%). At final concentrations of 1 mM or 10 mM, K+ and Na+ had little effect on LipR activity. LipR activity was not affected by PMSF, DTT, and β-ME when they were present at a 1 mM concentration. However, when their concentration was 10 mM, the residual activities of LipR declined to 69.81%, 92.36%, and 92.32%, respectively, of the original activity.

Table 3.Values are means ± SD from three independent experiments.

The effects of 10 organic solvents, at a 15% final concentration, on LipR activity are listed in Table 4. The activity of LipR was markedly affected by most solvents, except tert-butanol (77.71%), acetonitrile (84.55%), and methanol (91.08%). When the concentration of the organic solvents was 30%, LipR was extremely affected, but its residual activity towards methanol still remained at 77.36% of its original activity. LipR had a certain tolerance to Tween-20 (79.48%), Tween-80 (68.48%), SDS (55.97%), cetyltrimethylammonium bromide (62.93%), NP-40 (50.59%), and Triton X-100 (61.67%) at a concentration of 0.05%. After exposure to eight detergents at a 0.1% final concentration, the residual activity of LipR decreased to less than 50% of its original activity.

Table 4.Values are means ± SD from three independent experiments.

Hydrolysis of Algal Oil

According to the GC-MS analysis (data not shown), the algal oil contained 10.94% C14:0 (tetradecanoic acid), 23.95% C16:0 (hexadecanoic acid), 1.8% C16:1 (hexadecenoic acid), 0.76% C18:0 (stearic acid), 12.46% C18:1 (octadecenoic acid), 6.03% C18:2 (octadecadienoic acid), 0.74% C20:5 (EPA), 10.41% C22:5 (DPA), and 27.79% C22:6 (DHA), and approximately 95% of the nine major fatty acids were triacylglycerols (TAGs). The TLC analysis showed that LipR could hydrolyze algal oil to produce the corresponding 1,2-DAGs, 1,3-DAGs, and FFAs, including TAGs (Fig. 5A). Even the total contents of 1,2-DAGs and 1,3-DAGs were almost equivalent to the content of residual TAGs in the derivative glycerides. The GC-MS analysis (Fig. 5B) showed that the n-3 LCPUFAs EPA, DPA, and DHA were gradually enriched when C14–C18 long-chain fatty acids were continually released from the glycerin skeleton. After hydrolysis for 12 h, the relative contents of EPA, DPA, and DHA in the produced glycerides increased to 1% (1.35-fold), 18.32% (1.76-fold), and 33.88% (1.22-fold), respectively. At the same time, the relative contents of C14, C16, and C18 fatty acids decreased to 9.11%, 21.08%, and 16.61%, respectively. When the hydrolysis was extended to 24 h, the relative EPA, DPA, and DHA contents of derived glycerides increased to 1.22% (1.65-fold), 21.24% (2.04-fold), and 36.98% (1.33-fold), respectively. The relative contents of medium-long-chain fatty acids C14, C16, and C18 in the glycerides continued to decrease to 8.60%, 18.11%, and 13.85%, respectively.

Fig. 5.Analysis of the hydrolysis of algal oil.

 

Discussion

In general, degenerate primers were designed according to the conserved motif GHSLG [2,4]. According to the sequence alignment of lipases from Pseudomonas spp. and Bacillus spp., several conserved motifs were discovered, including GHSLG, GGXGXDXUX (a calcium-binding domain), an 18-residue amphipathic α-helix, and four hydrophobic residues at the extreme C-terminus [1]. We successfully obtained a 915 bp sequence fragment of lipase gene lipR using degenerate primers T3 and T5, which were aimed at the NVLNIGYE (residues 193–200) and NTFLFSGAF (residues 488–496) motifs. Meanwhile, touchdown PCR can offer a simple and rapid means to optimize PCRs, without the need for lengthy optimizations and/or the redesigning of primers, thereby increasing the specificity, sensitivity, and yield of PCR amplifications [16]. Similar to lipA from Acinetobacter sp. XMZ-26 [39], the entire lipase gene lipR also was successfully obtained via touchdown PCR and genome walking. Thus, it can be seen that the two domains in lipase subfamily I.3 can also be used for degenerate primer design, and touchdown PCR and genome walking could be combined to obtain the entire lipase gene in a future study.

There are some conserved motifs, including an 18-residue amphipathic α-helix (residues 530–547), four hydrophobic residues (residues 557–560), and six tandem repeats of the repetitive nine-residue motif GGXGXDXUX in LipR. These tandem repeats are characterized by a glycine- and aspartic-acid-rich region, which is also found in toxin motifs. These regions have also been verified to be involved in the ABC-transporter system for lipase transport in Pseudomonas sp. MIS38 (PDB: 2z8z) [17], and Serratia marcescens (GenBank: BAA02519) [21]. Meanwhile, the RTX motifs of LipR are proposed to function as an intramolecular chaperone. This suggests that LipR is a secretory lipase involved in ABC transporter systems.

According to Rost [29], enzyme function starts to diverge quickly when the sequence identity is below 70%. However, except for substrate preferences for p-NP butyrate and octanoate, there was no related report about the function of lipase LipT from P. mandelii JR-1 [14]. Another lipase from uncultured bacterium only showed lipolytic activity towards triolein/tributyrin and no further research was conducted on its applications [33]. Two other lipases from P. fluorescens Pf0-1 and P. fluorescens Pf-5 were putative ones deduced from the complete genome sequences [24]. Therefore, so far, there has been no function research on these four lipases. Moreover, the Enzyme Commission (EC) numbers only capture one aspect of protein function. When considering two enzymes to be similar or belonging to the same class, the full EC number does not describe all details about the function of a particular enzyme, for there may be considerable variations (i.e., function transfer) between two proteins with identical EC numbers [29]. Thus, more studies will be needed to reveal the relationship between the above-mentioned lipases, and this study on LipR may provide a beneficial reference for future studies on functions and/or functions transfer of other subfamily lipases.

By molecular structural modeling, we found that the two crystal structures (PDB: 2z8x [17]; and PDB: 3a70) of lipases from Pseudomonas sp. MIS38 were the most appropriate templates for homology modeling. Like the chosen template in the closed conformation, the final structure of LipR contains eight α-helices and 35 β-sheets, which are composed of three common domains, named as the catalytic domain, the α/β domain, and the regulatory domain (Fig. 1). The catalytic triad consists of Ser153, Asp202, and His206, which are located on the loops between β5-α5, β7-TT, and η4-α7, respectively, and are clustered together at the bottom of the active site. Homology studies (Fig. 6) indicate that the catalytic domain is located at the most proximal amino acids of the protein, whereas the C-terminal region of the protein is occupied by a well-defined β-roll structure that comprises several antiparallel β-sheets acting as calciumbinding sites [17,21]. A lid comprising residues 101–113 in LipR, which corresponds to the well-known lid of lipases, assumes a closed conformation in Fig. 6A and an open conformation in Fig. 6B. With the lid open, the number of α-helices increased (shown in Fig. 6).

Fig. 6.Three-dimensional model of LipR.

Most of the lipases from the genera Pseudomonas and Bacillus have been reported to have molecular masses ranging from 30 to 65 kDa, and previously reported bacterial subfamily I.3 lipases from these two genera have optimum temperatures ranging from 35℃ to 55℃, and optimum pH ranging from 7.5 to 9.0 [2]. The optimal temperature and pH of LipR were slightly higher than those of LipAAc (50℃, pH 7.0) from Acidobacteria sp. [8] and PSL (45℃, pH 8.5) from P. fluorescens SIK W1 [18]. In addition, LipAAc from Acidobacteria sp., which is a member of moderately thermostable bacterial lipases, retained 53% and 33% of its activity after incubation at 60℃ and 70℃, respectively, for 1 h [8]. In contrast to LipAAc, after incubation for 12 h at 50℃, 60℃, and 70℃, LipR still retained more than 90%, 77%, and 67%, respectively, of its original activity. Thus, it can be seen that the thermostability of LipR is better than that of LipAAc. Moreover, LipR was active in a wide temperature range from 0℃ to 100℃. These results demonstrate that LipR is a eurythermal lipase that would be suitable for biotechnological applications undertaken at high temperatures, such as the removal of pitch from pulp in the paper industry [32] and even at low temperature as additives in food industries, such as fermentation, cheese manufacturing, baking, and meat tenderizing [39].

LipA from S. marcescens has been verified to have tandem repeats, which are involved in asymmetric Ca2+ binding, within two β-roll structures, which are essential for its activity [21]. Similar to LipA, the activity of LipR was also heavily dependent on calcium. The activation by Ca2+ proved that the calcium-binding domain is important for the activity of LipR. The chelating agent EDTA could inhibit the activity of LipR to a great extent, which suggests that LipR is a metalloenzyme and that metal ions may be necessary for its catalytic activity, since EDTA can compete for the metal ion that stabilizes LipR [31]. The fact that some divalent metal ions, including Ca2+, Mg2+, Ba2+, and Sr2+, could activate LipR suggests that metal ions may be replaceable. At a 10 mM final concentration, the typical serine inhibitor PMSF obviously inhibited the activity of LipR, further confirming that it is a serine α/β hydrolase [8,10]. The reducing agents DTT and β-ME can reduce the disulfide bonds of proteins and prevent the formation of intra- or intermolecular disulfide bonds between cysteine residues [31]. However, the hydrolytic activity of LipR was not adversely affected by the two reducing agents. These results are consistent with the fact that LipR does not contain a cysteine residue and, therefore, cannot form a disulfide bond.

Table 5.Values are means ± SD from three independent experiments.

As a rule, non-ionic surfactants, such as Tween-20, Tween-40, Tween-60, Tween-80, Triton X-100, and NP-40, and ionic surfactants, including SDS and CTAB, do not stimulate, but instead inhibit, the activity of lipases [31]. LipC12 retained less than 30% of its original activity after treatment with these detergents [10]. A lipase from Fusarium oxysporum [26] and LipAAc belonging to an organism of the Acidobacteria phylum [8] were stable in the above surfactants and retained 60–70% of their activity after 1 h of incubation at 30ºC. Additionally, DR0334, DR1485, DR2078, and DR2522 from the extremely radioresistant bacterium Deinococcus radiodurans had exceeded 50% of their maximum activity in the presence of Tween-20 and Triton X-100 [31]. Similar to these reported lipases, LipR had a certain degree of resistance towards the listed detergents. On the one hand, the effects of commercial detergents showed different rates of lipase inhibition under the same conditions as the surfactants, probably because of the composition of commercial detergents [26,31]. On the other hand, SDS could cause local conformational changes in the active site that result in inhibition, partial reversible unfolding, and subsequent inactivation [26]. In general, lipases with good tolerance to organic solvents are mainly derived from Pseudomonas and Bacillus species [27]. LipR showed better tolerance to acetonitrile (84.55%) and tert-butanol (77.71%) at a 15% concentration. Moreover, after treatment with 15% and 30% methanol, LipR still retained 91.08% and 77.36% of its original activity, respectively. These results suggest that LipR is a potential candidate for application in organic syntheses, transesterifications, and enantioselective resolutions [11,14].

Similar to reported lipases from A. niger [22], G. candidum [36], and Y. lipolytica [37], LipR could gradually split other fatty acids (C14, C16, and C18) from the glycerin skeleton. There are some differences, as a lipase from A. niger slightly increased the DHA content from 13.63% to 18.72% by hydrolyzing sardine oil for 3h. Additionally, lipase from G. candidum mildly increased the total of EPA and DHA contents from 38.5% to 48.7%, respectively, after hydrolyzing fish oil for 16 h, whereas immobilized LIP2 from Y. lipolytica could only enrich the DHA content from 19.32% to 31.53% by hydrolyzing Chlorella protothecoides oil for 4 h. Even though LipR needed 24 h to hydrolyze algal oil, there was a synchronous enrichment of the n-3 LCPUFAs EPA, DPA, and DHA (from 38.94% to 59.44%) with a certain amount of DAGs during the production of glycerides. In comparison with previous reports [22,36,37] dealing with enrichment of EPA and/or DHA and/or DPA, the variation in terms of concentrated effect could be explained by different original oil sources, different sources of lipases, distinct specificities for various substrates, different types of lipase preparations, the optimization of parameters, and the use of different reaction conditions.

Evidence for the unique effects of n-3 LCPUFAs in the treatment of a variety of neurodegenerative and neurological disorders and as prevention and treatment function against cancer, arteriosclerosis, inflammation, and hyperlipemia are growing [7,9,36]. However, the recommended individual intake of the n-3 LCPUFAs is not met by dietary sources such as fish. Additionally, a survey of the literature has shown that DAG oil has beneficial effects on obesity and weight-related disorders [20]. The attractive feature of LipR is its specificity with respect to the glyceride position and fatty acid type, which could seldom be obtained by chemical catalysis, which suggests that LipR may be useful for processing edible fats and oils containing EPA, DPA, and DHA in the food industry, and/or the production of DAGs in the food or cosmetic industries.

In conclusion, culturable microorganisms are important sources of versatile lipases. The lipase LipR from Pseudomonas sp. R0-14 in our study may serve as a eurythermal and/or thermostable catalyst in industrial applications. Our results regarding the hydrolysis of algal oil by LipR also provide important information about the enrichment of n-3 LCPUFAs and the production of certain DAGs in the derivative glycerides. These results suggest that LipR has potential in various industrial applications, especially in food and/or cosmetic applications.