Advertisement
Browse Subject Areas
?

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Bacillus sp. JR3 esterase LipJ: A new mesophilic enzyme showing traces of a thermophilic past

  • Judit Ribera,

    Roles Investigation, Methodology

    Affiliation Department of Genetics, Microbiology and Statistics, Faculty of Biology, University of Barcelona, Av. Diagonal 643, Barcelona, Spain

  • Mónica Estupiñán,

    Roles Data curation, Formal analysis, Investigation, Methodology, Writing – original draft

    Affiliations Department of Genetics, Microbiology and Statistics, Faculty of Biology, University of Barcelona, Av. Diagonal 643, Barcelona, Spain, Institute of Nanoscience and Nanotechnology (IN2UB), University of Barcelona, Av. Diagonal 643, Barcelona, Spain

  • Alba Fuentes,

    Roles Investigation, Methodology

    Affiliation Department of Genetics, Microbiology and Statistics, Faculty of Biology, University of Barcelona, Av. Diagonal 643, Barcelona, Spain

  • Amanda Fillat,

    Roles Conceptualization, Formal analysis, Supervision, Validation

    Affiliation Department of Genetics, Microbiology and Statistics, Faculty of Biology, University of Barcelona, Av. Diagonal 643, Barcelona, Spain

  • Josefina Martínez,

    Roles Data curation, Formal analysis, Resources

    Affiliations Department of Genetics, Microbiology and Statistics, Faculty of Biology, University of Barcelona, Av. Diagonal 643, Barcelona, Spain, Institute of Nanoscience and Nanotechnology (IN2UB), University of Barcelona, Av. Diagonal 643, Barcelona, Spain

  • Pilar Diaz

    Roles Conceptualization, Funding acquisition, Resources, Supervision, Validation, Writing – original draft, Writing – review & editing

    pdiaz@ub.edu

    Affiliations Department of Genetics, Microbiology and Statistics, Faculty of Biology, University of Barcelona, Av. Diagonal 643, Barcelona, Spain, Institute of Nanoscience and Nanotechnology (IN2UB), University of Barcelona, Av. Diagonal 643, Barcelona, Spain

Bacillus sp. JR3 esterase LipJ: A new mesophilic enzyme showing traces of a thermophilic past

  • Judit Ribera, 
  • Mónica Estupiñán, 
  • Alba Fuentes, 
  • Amanda Fillat, 
  • Josefina Martínez, 
  • Pilar Diaz
PLOS
x

Abstract

A search for extremophile enzymes from ancient volcanic soils in El Hierro Island (Canary Islands, Spain) allowed isolation of a microbial sporulated strain collection from which several enzymatic activities were tested. Isolates were obtained after sample cultivation under several conditions of nutrient contents and temperature. Among the bacterial isolates, supernatants from the strain designated JR3 displayed high esterase activity at temperatures ranging from 30 to 100°C, suggesting the presence of at least a hyper-thermophilic extracellular lipase. Sequence alignment of known thermophilic lipases allowed design of degenerated consensus primers for amplification and cloning of the corresponding lipase, named LipJ. However, the cloned enzyme displayed maximum activity at 30°C and pH 7, showing a different profile from that observed in supernatants of the parental strain. Sequence analysis of the cloned protein showed a pentapeptide motif -GHSMG- distinct from that of thermophilic lipases, and much closer to that of esterases. Nevertheless, the 3D structural model of LipJ displayed the same folding as that of thermophilic lipases, suggesting a common evolutionary origin. A phylogenetic study confirmed this possibility, positioning LipJ as a new member of the thermophilic family of bacterial lipases I.5. However, LipJ clusters in a clade close but separated from that of Geobacillus sp. thermophilic lipases. Comprehensive analysis of the cloned enzyme suggests a common origin of LipJ and other bacterial thermophilic lipases, and highlights the most probable divergent evolutionary pathway followed by LipJ, which during the harsh past times would have probably been a thermophilic enzyme, having lost these properties when the environment changed to more benign conditions.

Introduction

Lipolytic enzymes (EC 3.1.1.-), widely distributed in nature, are a diverse group of hydrolases catalysing the cleavage or formation of ester bonds [1,2]. Grouped under the general term of lipases, lipolytic enzymes include “true” lipases (EC 3.1.1.3, triacylglycerol hydrolases) and esterases (EC 3.1.1.1, carboxyl ester hydrolases), which differ in both, their kinetics and chain-length substrate preferences [1,2]. Esterases are active on short-chain length esters partially soluble in water, and lipases have optimal activity towards long-chain triacylglycerides, not soluble in aqueous environments [2]. Therefore, esterases display a typical Michaelis-Menten behaviour, whereas many lipases show interfacial activation when lipids reach an equilibrium between the monomeric, micellar and emulsified states [2]. Although the physiological role of many microbial lipases remains still unclear, in bacteria most esterases are intracellular enzymes involved in lipid metabolism and turnover [3,4]. On the contrary, most bacterial true lipases are secreted enzymes and perform their role extracellularly, probably by procuring nutrients to the cell or by executing synthesis reactions aimed at releasing modified lipid compounds that could act as extracellular cell communication signals, or have a role in detoxification of biocides [36].

The 3D structure of both, lipases and esterases, consists on a defined alternation of α-helices and β-sheets forming the characteristic α/β–fold also found in other hydrolases [68]. Their active site is constituted by a catalytic triad containing a serine as the catalytic residue, an aspartic or glutamic acid, and a histidine, usually being the nucleophile serine embedded in the GXSXG consensus pentapeptide [2,9]. Bacterial lipolytic enzymes were initially classified into eight families [10], with successive revisions up to more than 16 families described nowadays [9,11,12]. These classifications are based on amino acid sequence homology and the presence of conserved motifs, although more recently, certain structural features and phylogenetic analysis have come to complement this classification [912].

During the last years the interest generated by lipases has grown exponentially due to both, their broad array of substrate specificity and their versatility in catalysed reactions [1316]. Therefore, these enzymes are considered powerful biocatalysts with many biotechnological applications such as food technology, detergent formulation, flavour and drug production, or in the synthesis of optically pure compounds, among other uses in fine chemistry [2,13,1518]. Moreover, lipases do not usually require cofactors, are quite robust and stable, and can be active in organic solvents [1,1820]. However, not all lipolytic enzymes bear the properties required for a defined biotechnological process, and availability of new catalysts with striking properties is still a challenge [21,22]. For those reactions requiring harsh conditions like high temperature or extreme pH, different strategies can be followed. Among them, microbial isolation from natural environments is one of the most used strategies to discover new enzymes which can fit the conditions required by the industrial processes [2325]. This is the case for certain environments that supported high temperatures or extreme conditions in old times. Therefore, ancient volcanic soils from Laurel forest (Laurisilva) of El Hierro Island (Canary Islands, Spain) are a good example of such environments, as they suffered from harsh conditions in the past, when the volcanic island appeared approx. 1 million years ago, but evolved to acquire milder conditions nowadays. Evolution of the volcanic island from an extreme environment to its present benign state, a cloudy forest with plenty of organic matter, endemic vegetation and mild temperatures due to the north-east trade winds and the Föhn effect, is an exceptional niche for isolation of undescribed strains with reminiscent extreme biocatalysts. Therefore, a search for new strains was performed, focussing our interest on lipolytic enzymes from sporulated bacteria. Several microbial strains were isolated from an ancient volcanic soil sample, and their catalytic activities analysed. Strain JR3, displaying extracellular lipolytic activity up to 100°C was selected for further prospection and characterization of novel thermophilic lipases. The mixed molecular and biochemical properties of the cloned enzyme LipJ are described and discussed here.

Materials and methods

Strains, plasmids and growth conditions

A collection of sporulated strains (Table 1) was isolated from an ancient volcanic soil, now a Laurel forest at El Hierro Island (coordinates 27.735163–17.993231; sampling performed in a public area of the municipality of El Pinar, with permission of the National Institute of Geography of the Canary Islands, Spain), using the conditions stated in Table 1 and following a previously described isolation protocol with modifications [25]. Microorganisms were extracted by suspending 1 g of soil in 10 mL Ringer ¼. After 10 min vigorous stirring and additional sedimentation, aliquots of the upper liquid phase were collected and used for isolation of aerobic and facultative anaerobic spore-forming bacteria by means of serial dilutions in Ringer 1/4. 0.2 mL of each dilution were spread on Luria-Bertani medium (LB) agar plates and incubated at 30°C and 55°C for 1 to 7 days. Those colonies displaying different morphological properties were isolated, their Gram and spore stain performed, and their hydrolytic activity on several substrates at the optimum growth temperature was analysed on Trypticase Soy Agar (TSA) (Pronadisa) plates supplemented with sterile skimmed milk (1% v/v, Sharlau), LB agar plates supplemented with carboxymethyl cellulose (CMC) (0.5% w/v, Sigma), CeNAN (ASDA Micro) supplemented with olive oil (1% w/v, Carbonell) or tributyrine (1% w/v, Sigma) emulsified with 0.1% arabic gum (w/v, Sigma) and 0.0002% Rhodamine B (v/v, Sigma) [26], and incubated at the isolation temperature of each strain, as previously described [25]. Strain Paenibacillus barcinonensis BP-23 (CECT 7022) [27] was used as hydrolytic positive control. Pure cultures of each newly isolated strain were maintained at 4°C and stored in glycerol stocks at −80°C.

thumbnail
Table 1. Strains isolated from a forest at the volcanic El Hierro Island (Canary Islands, Spain).

Activity assays were performed at 30°C on LB-agar plates supplemented with either milk (protease), carboxymethylcellulose (cellulose), olive oil (lipase) or tributyrine (esterase). According to the 16S rDNA sequence and physiological tests, strain JR3 was identified as Bacillus sp. JR3, close but not identical to B. cereus.

https://doi.org/10.1371/journal.pone.0181029.t001

Selected isolate Bacillus sp. JR3 strain (CECT 9334) and control strain P. barcinonensis BP-23 were grown in LB for 24 hours at 30°C, under aerobic conditions. E. coli DH5α and BL21 star (DE3) strains were routinely cultured overnight at 37°C in LB broth or on LB agar plates, and were used as host strains for cloning and expression of the enzyme-encoding gene. Plasmids pGEM®-T Easy and pET28a/pET-101-D-TOPO® (Promega/Novagen/Invitrogen®) were used as cloning/expression vectors producing strains E. coli DH5α/pGEMT-LipJ, E. coli BL21/pET28a-LipJ, and E. coli BL21/pET101D-LipJ-HisTag. Media were supplemented with antibiotics (ampicillin 100 μg ml-1; kanamycin 50 μg ml-1) or IPTG (isopropyl-β-D-thiogalactopyranoside; 1mM) and X-gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside; 80 μg ml-1) when required for recombinant clone selection or gene expression [28].

DNA procedures

Genomic DNA was extracted using the GeneJet Genomic DNA Purification Kit (Thermo Scientific) according to the manufacter’s instructions. Plasmid DNA was purified using commercial kits (NucleoSpin Plasmid, Macherey-Nagel). DNA synthesis and restriction enzymes (Biotools/Thermo Scientific) were used following the manufacturer’s recommendations. PCR amplifications were performed in a Gene Amp® PCR System 2400 (Perkin Elmer) and T100™ Thermal Cycler (Bio-Rad) using different cycling periods. PCR resulting DNA was purified with Gel Band Purification kit (Thermo Scientific). To obtain the nucleotide sequences of DNA, PCR amplified fragments were analysed using the ABI PRISM® BigDye® Terminator v.3.1 Cycle Sequencing Kit (Applied Biosystems), and the analytical system CEQTM 8000 (Beckman-Coulter) available at the Serveis Científics i Tecnològics of the University of Barcelona. DNA samples were routinely analysed by 0.8% (w/v) agarose gel electrophoresis [28], and stained with 0.001% Nancy-520 DNA gel stain (v/v, Sigma-Aldrich). Nucleic acid concentration and purity was measured using a Spectrophotometer ND-100 NanoDrop®.

Identification of strain JR3

Universal primers FW27F/BW1525R [29] were used for PCR amplification of the 16S rDNA gene from strain JR3. The complete 16S rDNA gene homology search was achieved using GenBank database. Additional information was obtained after observation of bacterial morphology and endospore location plus biochemical characterization tests including carbohydrate utilization patterns (API®20E and API®50CH systems, BioMérieux) for comparison with other strains of Bacillus species showing 99% identity and 100% coverage in 16S rDNA sequence (GenBank).

Extracellular fraction preparation

Extracellular medium samples of Bacillus sp. JR3 were obtained from 24 h grown cultures of the strain in LB after centrifugation at 8,000 x rpm for 20 minutes. Supernatants were recovered and kept at 4°C for extracellular activity evaluation. Concentrated (50 X) supernatant samples were obtained using ultrafiltration CentriconTM devices (Amicon®) of 10 kDa cutoff and analysed by 12% SDS-PAGE [30] after short time (0 to 5 minutes) treatments at 100°C. Zymogram analysis was performed as previously described [31,32] using 100 μM MUF-butyrate prepared in Tris HCl 20 mM buffer (pH 7) at 30 and 60°C, and visualized under UV light in a Gel Doc XR System (Biorad) before staining with Coomasie Brilliant Blue R-250 (BioRad).

Primers design for thermophilic lipase isolation

Degenerate primers (Table 2) were designed to amplify different conserved regions found in Bacillus sp. and Geobacillus sp. thermophilic lipases, including the N- and C-terminal ends, the protein core, the active site and the oxyanion hole homolog regions. Using Bacillus sp. JR3 genome as a template, all possible primer combinations were used for amplification in a gradient thermocycler T100™ Thermal Cycler (Bio-Rad). 16S rDNA specific primers were used as positive control for amplification assays. Reliable PCR products were obtained with primers FWintLipGBCdeg/BWintLipGBCdeg and FWintLipBCdeg/BwintLipBCdeg, which were further sequenced and analyzed by BLASTn [33]. Those sequences showing the highest identity values were used to produce a lipase consensus sequence.

Cloning Bacillus sp. JR3 LipJ gene

From the consensus sequence obtained, new degenerate primers LipBFwInNcoI/LipBFwOutHindIII (Table 2) were designed and used for amplification (Expand High Fidelity polymerase, Roche) of a putative thermophilic lipase, designated LipJ, using JR3 genomic DNA as a template. The resulting PCR fragment was ligated to pGEM-T® Easy vector (Promega) and transformed into E. coli DH5α. For overexpression, pGEMT-LipJ was digested with NcoI/HindIII, ligated to the doubly digested expression vector pET28a and transformed in E. coli BL21 star (DE3). Additionally, lipJ gene was amplified (Kappa-HiFi polymerase, Kapa Biosystems) using specific primers FwLipJTOPO/BwLipJTOPO, and ligated to pET101/D-TOPO, producing recombinant plasmid pET101-LipJ-Histag, which contains the full-length enzyme linked to a C-terminal His6-tag. All constructions were verified by sequencing. The DNA sequence of lipJ was submitted to NCBI and given the GenBank Accession Number KU747177.

Expression and purification of LipJ

For LipJ production, exponential growth cultures (OD600nm = 0.6–0.8) of recombinant E. coli BL21/pET28a-LipJ in LB medium supplemented with kanamycin (50 μg/ml) were induced with 1 mM IPTG at different temperatures for 24 h. Cells were collected by centrifugation at 6,000 x rpm for 15 min, suspended in 20 mM Tris-HCl buffer pH 7.0 and disrupted using a SLM Aminco French Press. Clarified cell extracts were then recovered for activity determination after centrifugation at 10,000 x rpm for 10 minutes. For determination of the enzyme kinetics parameters (Km, Vmax), recombinant His6tag LipJ was purified from clarified cell extracts by immobilized metal affinity chromatography (IMAC) using HisTrap HP columns of 1 ml (GE Healthcare), and eluted in 20 mM Tris-HCl buffer (pH 7) with 500 mM NaCl using a 0 to 500 mM imidazol gradient on a fast protein liquid chromatography system (ÄKTA FPLC; GE Healthcare). Activity on pNP-butyrate (see below) was detected in a single elution fraction. Buffer exchange was performed by dialysis in 20 mM Tris-HCl buffer, shaking overnight at 4°C. Protein concentration was achieved in Centricon centrifugal filter units of 30-kDa molecular mass cutoff (Millipore). Bradford method was performed for protein concentration determination [34], using bovine serum albumin (BSA) as the standard.

Activity assays

Lipolytic activity of crude cell extract, supernatant or purified samples was analysed by measuring the release of para-nitrophenol (pNP) from pNP-derivative fatty acid substrates (1mM for C2-C5 substrates and 0.5 mM for C10-C16 substrates, Sigma), as previously reported [35,36] using a Bio-Rad 3550 microplate reader. One unit of activity was defined as the amount of enzyme that released 1μmol of pNP per minute under the assay conditions used. All determinations of enzyme activity were performed by two replicas of triplicates (6 determinations per sample).

Optimum temperature of Bacillus sp. JR3 and P. barcinonensis supernatants was determined by analysis of the activity over a range from 4 to 100°C, at pH 7, using 1mM pNP-butyrate as a substrate [35]. Thermal stability was determined by incubating cell extracts or supernatant samples at temperatures from 4 to 100°C for 1 to 96 h; residual activity was measured under standard assay conditions once samples reached the optimum temperature. pH stability of samples was determined by measuring the residual activity on pNP-butyrate after 1 h incubation at different pH. The effect of temperature or pH on LipJ lipolytic activity was evaluated by response surface methodology (RSM) [37], as stated below. To evaluate the effect of metal ions or inhibitors on activity, assays were performed on pNP-butyrate in the presence of several metal chlorides (Al+3, Ba+2, Ca+2, Cu+2, Fe+2, Hg+2, K+, Li+2, Mg+2, Mn+2, Na+, NH4+, Zn+2) and EDTA, used at different concentrations (1 and 10 mM). Moreover, Ca+2 (1–30 mM) and Zn2+ (0–5 mM) were assayed at 30°C and 60°C and in the range 60–100°C. Residual activity was measured at 30°C and pH 7 using a conventional assay on pNP-butyrate, and expressed as a percentage of activity without ions or inhibitors, respectively. Supernatant samples of Bacillus sp. JR3 were also incubated in the presence of 1 mM PMSF (phenylmethylsulfonyl fluoride, Sigma) to measure the effect of this serine inhibitor on enzyme activity. Kinetic parameters (Vmax and Km) were determined under optimal assay conditions by fitting hyperbolic Michaelis-Menten curves with GraphPad Prism ® software version 6.

Statistical analysis

For optimum pH and temperature determination, response surface methodology (RSM) [37] was applied, using a design of experiment of 22 Central Composite Designs (CCD) with 5 levels leading to 11 sets of experiments with 3 replicates of the central position. Temperatures ranging from 16 to 50°C and the Britton-Robinson 50 mM buffer adjusted to pH ranging from 4.2 to 9.8 were used. The experimental results of CCD, represented by the mean of at least 6 samples each, were fitted to the following second order polynomial equation: (z = −186.5841+48.6774x−3.4315x2+2.9364y−0.0500y2−0.0173xy). Quality of this equation was evaluated by the coefficient of determination R-sqr, which was 0.94557, indicating that 94.56% response data can be justified by the chosen model. Table 3 shows that pH and temperature (°C), and the interaction of both of them (1L by 2L), do not fit a linear regression, as P > 0.005, and both variables are independent. However, these variables can be adjusted to a quadratic model regression with a 90% confidence level, as P < 0.05. Therefore, the model used allowed prediction of any activity value, providing values of pH and temperature with a precision of 94.56% and 90% confidence. The statistical significance of the Central Composite Design model was determined by the F-test ANOVA. Statistical analysis and RSM values were calculated with STATISTICA software version 8.

Bioinformatics tools

BLAST searches were routinely performed for DNA or protein sequence analysis and to retrieve identity and similarity percentages by pairwise alignment [33]. Multiple sequence alignments were obtained using T-Coffee (http://tcoffee.crg.cat/) [38] or ClustalO (http://www.ebi.ac.uk/Tools/msa/clustalo/) [39]. BioEdit v.7.0.1 [40] was used for restriction and consensus pattern determination. Identification of putative signal peptide [41] was performed through SignalP versions 3.0 or 4.0 (http://www.cbs.dtu.dk/services/SignalP/), and transmembrane regions determined through TMHMM tools (http://www.cbs.dtu.dk/services/TMHMM-2.0). ExPASy proteomics server was used to analyse the protein physico-chemical parameters (ProtParam tool; http://web.expasy.org/protparam/), and to predict isoelectric point and molecular mass of the cloned protein [42]. Detection of aromatic amino acids was performed using GPMAW tool (http://www.alphalyse.com/customer-support/gpmaw-lite-bioinformatics-tool/start-gpmaw-lite/) [42]. ProDom (http://prodom.prabi.fr/prodom/), Pfam (http://pfam.xfam.org/), and InterProScan 5.2 (http://www.ebi.ac.uk/interpro/) were used for domain identification [43], and a Hidden Markov Model was created with HMMER (http://www.ebi.ac.uk/Tools/hmmer/) in order to detect conserved protein motifs [44]. LipJ amino acid sequence was compared with PDB database to identify possible template proteins with available 3D structure. Pelosinus fermentans lipase PfL1 (pdb 5AH0, 92% coverage and 56% identity) [45] and Geobacillus stearothermophilus lipase L1 (pdb 1KU0, 92% coverage and 49% identity) [46] were selected for homology model construction and validation using Phyre2 server (http://www.sbg.bio.ic.ac.uk/~phyre2/html/page.cgi?id=index) [47]. 3D structure superposition and search for conserved amino acid motifs or ion binding cavities was performed using the visualization tool Pymol Molecular Graphics System, Version 1.5.0.4, Schrödinger, LLC (http://www.pymol.org). For family assignment, a phylogenetic analysis was conducted using MEGA 7 software including lipase sequences of functionally characterized enzymes bearing similar catalytic mechanisms, available in the literature and in UniprotKB (http://www.uniprot.org/) or in Protein Data Bank (http://www.rcsb.org/pdb/) databases, always considering their catalytic and conserved residues [48,49]. A phylogenetic tree was constructed applying the Maximum Likelihood (PALM) method, employing WAG+G as the amino acid substitution model [50]. A bootstrap consensus tree was achieved after 1000 repeats, with a cut off of 50% [50]. Family assignment was confirmed using ESTHER database (http://bioweb.ensam.inra.fr/esther), specific for α/β-hydrolase fold protein superfamily [51].

Results and discussion

Bacillus sp. JR3 isolation

A green laurel forest located at the volcanic island of El Hierro (Canary Islands, Spain) was chosen as a source for isolation of spore-forming bacteria coding for exceptional enzymatic activities resulting from natural evolution of volcanic soils, and due to their capacity to persist in extreme environments. Seven Gram-positive bacterial sporulated isolates (JR1 to JR7) showing either protease, cellulase, lipase or esterase activity were obtained after growth at mild temperature (30°C) for 24 h in aerobic conditions (Table 1). Among them, strain JR3 was selected for further identification and characterization due to its differential morphology and strikingly high esterase activity when culture supernatants were assayed on pNP-butyrate at 60°C. BLAST analysis of the 16S rDNA sequence of strain JR3 allowed assigning this isolate to the genus Bacillus, in close proximity to the B. cereus and B. thuringiensis group (query cover 100%, maximum identity 99%). Additional API®20E i API®50CH tests were performed and the results obtained again assigned the strain to the genus Bacillus, almost identical to B. cereus, although no complete match was obtained for gelatine utilization and acetoin production, which resulted positive and negative, respectively. Therefore, the strain was named Bacillus sp. JR3.

Extracellular esterase activity of Bacillus sp. JR3

Extracellular lipase activity of strain JR3 was initially tested on different chain-length pNP-derivative substrates at 37°C. Supernatants of the strain grown at 30°C for 24 h showed activity on C2 to C7 substrates (Fig 1A). However, under the conditions used here (15 minute incubation assays), no significant activity could be detected on substrates of longer chain length (C10 to C18). Maximum activity at 37°C was obtained on pNP-butyrate (21.23 U g-1), whereas activity on pNP-acetate and pNP-valerate was 4.72 and 0.43 U g-1, respectively. Activity assays performed using crude cell extracts of the strain grown under the same conditions revealed also the presence of a faint intracellular activity, with a maximum value at 37°C of 2.11 U g-1 on pNP-butyrate (Fig 1A*).

thumbnail
Fig 1. Activity assays of Bacillus JR3 supernatants.

(A) Substrate specificity profile; *cell extract activity was only assayed on p-NP-butyrate. (B) Activity at different temperatures of strain JR3 (blue diamonds) and P. barcinonensis (red squares), the latter used for comparison of mesophilic activity.

https://doi.org/10.1371/journal.pone.0181029.g001

The apparent kinetic parameters of supernatant samples from Bacillus sp JR3 were determined at 37°C using pNP-butyrate. A typical Michaelis-Menten kinetics plot was obtained (not shown), with a Km of 0.076 mM and a Vmax of 0.522 U mg-1, showing lack of interfacial activation, therefore suggesting that the major extracellular lipolytic activity of the strain corresponds to esterase, according to the proposed classification based on substrate specificity [2].

Optimum temperature of strain JR3 supernatants was determined as described in Materials and Methods, resulting in a strikingly high activity at temperatures over 60°C (Fig 1B), reaching maximum activity at 80–100°C (ca. 90 U g-1). To assess that the results obtained were not due to an artefact caused by the effect of such high temperatures on substrate or buffer stability, supernatants from P. barcinonensis BP-23 [27] were used as internal control and assayed for activity over the same range of temperatures. P. barcinonensis displayed a typical mesophilic lipase activity profile compared with that of supernatants from Bacillus sp. JR3 (Fig 1B), suggesting the existence of a thermophilic lipolytic extracellular system in JR3. To further confirm this interesting behaviour, new activity assays were performed at high temperatures in the presence of PMSF. PMSF is a common serine inhibitor of enzymes bearing a nucleophilic serine at the active site [15], as is the case for lipases [2,10]. Assays performed for only 15 minutes at 80°C showed almost complete inhibition (81% activity loss) of activity in supernatant samples of strain JR3 in the presence of PMSF, whereas full activity was achieved when PMSF was not present in the reaction medium (Fig 2A). These results unambiguously confirm that the activity found at high temperatures in supernatants of strain JR3 was not artefactual but it was indeed due to the presence of at least a thermophilic lipolytic enzyme. However, it must be stated that the strain was isolated at 30°C and could not grow when incubated at temperatures over 50°C, indicating that it is in fact a mesophilic strain, yet bearing thermophilic esterase activity, probably as a reminiscence of its life under the ancient extreme environmental conditions of the volcanic soil.

thumbnail
Fig 2. Temperature assays of Bacillus JR3 extracellular activity.

(A) Activity assay at 100°C with and without PMSF. (B) Residual activity found after 15 min incubation in a temperature range from 4 to 100°C. (C) Long-term thermal stability of JR3 (blue diamonds, blue squares, blue triangles) and P. barcinonensis (black +, blue Ӿ, red dots) supernatants, incubated for 90h at 60 (blue diamonds, black +), 80 (blue squares, blue Ӿ) and 100°C (blue triangles, red dots). P. barcinonensis supernatant activity measured for comparison of mesophilic activity.

https://doi.org/10.1371/journal.pone.0181029.g002

Thermal stability was assayed by measuring the residual activity after incubation of supernatant samples of strain JR3 in a temperature range from 4 to 100°C for 15 minutes. Almost complete activity recovery was obtained at all assayed temperatures (Fig 2B). Moreover, thermal stability assays performed at 60, 80 and 100°C for 96 hours showed a 50% activity loss only after 50 and 96 hours incubation at 100 and 80°C, respectively, while supernatants of P. barcinonensis rapidly lost their activity at high temperatures (Fig 2C). Interestingly, only a 15% loss of activity was observed after 96 hours incubation at 60°C, thus confirming the interesting thermoresistance and thermophilicity of JR3 extracellular lipolytic activity (Fig 2C), which could meet the conditions of biotechnological processes requiring high temperatures and long incubation times [52]. Therefore, we focussed on the prospection of thermophilic lipases/esterases in the genome of strain JR-3.

Gene isolation and cloning

Search for thermophilic Bacillus sp. or Geobacillus sp. lipase sequences in the databases revealed that they all belong to the bacterial lipase family I.5 [10]. Among the lipases from these genera, two clusters were established upon multiple sequence alignment (Fig 3A and 3B): those belonging to the B. cereus group (20–35 kDa), and those of Geobacillus group (40–45 kDa), a genus that was created to include thermophilic members of former Bacillus species [53]. Both clusters share a common central region of ca. 480 bp (Fig 3A and 3B) for which the set of degenerate primers, FWintLipGBCdeg/ BWintLipGBCdeg (conserved motifs SSNWDRACE and YDFKLDQW) was designed (Table 2). On the other hand, lipases from the Geobacillus cluster are longer in size, and contain two conserved sequence fragments flanking the central common region. A new set of degenerate primers, FwintLipGdeg/ BWintLipGdeg (conserved motifs GWGREEM and NDGIVNT) was designed (Table 2) for amplification of a putative thermophilic Geobacillus-like lipase sequence. Both sets of primers produced amplicons of ca. 450 and 900 bp, respectively, which were aligned with the closest sequences in the databases to produce a consensus sequence that was used to design two new sets of degenerate primers (Table 2), one inside (LipBFwInNcoI, LipBBwInHindIII) and another outside (LipBFwOutNcoI, LipBBwOutHindIII) the consensus sequence coding region. The four primer combinations were tested for amplification using Bacillus sp. JR3 genomic DNA as a template, and a band of ca. 1300 bp was obtained with primers LipBFwInNcoI and LipBBwOutHindIII (Table 2). The amplified 1300 bp DNA fragment was sequenced and analysed by BLASTn, showing 91–96% identity and 92% coverage with predicted lipase sequences belonging to the B. cereus group. This fragment, designated lipJ, was further digested with NcoI and HindIII, and cloned in E. coli BL21 using vectors pGEM-T®, TOPO and pET28. Presence of lipJ insert in the resulting recombinant clones was confirmed by restriction digestion and sequencing, showing the presence of an ORF of 1242 bp coding for a hypothetical protein of 413 amino acids, with a predicted molecular mass of 46 kDa and a pI of 6.5, displaying activity on tributyrin (Fig 3C). The complete sequence of lipJ was submitted to GeneBank and given the accession number KU747177.

thumbnail
Fig 3.

Conserved motifs (A) and regions (B) of short Bacillus and long Geobacillus lipases found in the databases, used for the design of consensus degenerated primers employed for amplification of an internal lipase coding DNA fragment from strain JR3. (C) Tributyrin-supplemented plate assay showing the hydrolysis haloes produced by cloned LipJ (left) and strain JR3 (right).

https://doi.org/10.1371/journal.pone.0181029.g003

LipJ purification and characterization

Expression assays of E. coli BL21/pET28-LipJ were done after IPTG induction at different temperatures (10°C, 21°C, 30°C and 37°C). Activity of soluble and insoluble fractions of induced cell extracts was analysed by zymogram [31,32], showing a band of ca. 40 kDa with activity on MUF-butyrate at 30°C (Fig 4A). The highest expression was obtained in soluble fractions of cell extracts induced at 21°C with 1 mM IPTG.

thumbnail
Fig 4. Expression and activity profile of cloned LipJ.

(A) Coomassie-stained (left) and zymogram [31,32] (right) SDS-PAGE of crude cell extracts (E) and cell debris (P) of cloned LipJ induced with 1 mM IPTG at different temperatures. (B) Michaelis-Menten kinetics profile of cloned LipJ. (C) Substrate specificity of cloned LipJ assayed on pNP-derivative substrates of different chain length.

https://doi.org/10.1371/journal.pone.0181029.g004

For characterization purposes, LipJ was purified by fast protein liquid chromatography from 20-fold concentrated crude cell extracts of recombinant E. coli BL21/pET101D-LipJ-HisTag. The purification process rendered a low yield (5.3%) that allowed isolation of a small sample of semi-purified protein. Unexpectedly, the resulting semi-purified enzyme did not display activity at 80°C, showing also very low activity at 37°C, thus allowing only the study of the kinetic parameters of LipJ. When assayed at 37°C on pNP-butyrate, the enzyme displayed a typical Michaelis-Menten plot (Fig 4B) without interfacial activation, like most esterases, with a calculated apparent Km and Vmax of 1.7 mM and 21.6 U g-1, respectively. Further characterization of LipJ was therefore performed using crude cell extracts of E. coli BL21/pET28-LipJ, prepared as described in Materials and Methods.

LipJ substrate specificity was assayed at 37°C on several pNP-derivatives, showing preference for short chain-length fatty acid substrates (Fig 4C), and exhibiting the highest activity (52.1 U g-1; 100%) on pNP-butyrate (C4:0). The enzyme maintained almost 95% activity on pNP-acetate (C2:0; 45.7 U g-1) and 20% on pNP-valerate (C5:0; 10.3 U g-1). However, a dramatic activity reduction was observed when long chain length substrates were used under the same conditions (Fig 4C).

The effect of temperature and pH on the activity of LipJ was determined on pNP-butyrate (Fig 5A), using a Surface Response Methodology strategy [37]. Surprisingly, LipJ displayed maximum activity at pH 7.0 and 28.13°C, whereas no activity could be detected at extreme pH or high temperatures. Moreover, thermal stability assays performed at optimum pH and temperature on pNP-butyrate demonstrated that the cloned enzyme was rapidly inactivated when incubated at temperatures over 30°C (Fig 5B). These results are in clear contradiction with the extracellular thermophilic profile found in the supernatant of strain Bacillus sp. JR3, and suggest that the cloned enzyme LipJ is not the thermophilic enzyme we intended to clone. On the contrary, the catalytic behaviour of LipJ points to a mesophilic esterase that could even be intracellular, based on the fact that most secreted Bacillus-related lipases are alkaliphilic [12,54,55], while LipJ only shows activity at neutral pH, as happens for most intracellular esterases [5658].

thumbnail
Fig 5. Properties of cloned LipJ.

(A) RSM results plot for optimum temperature and pH determination. (B) Thermal stability of LipJ cell extracts incubated for 1h at different temperatures.

https://doi.org/10.1371/journal.pone.0181029.g005

Thus, taking into consideration the previous results and on view of the kinetic parameters displayed by LipJ, with lack of interfacial activation, we propose that LipJ should be considered an esterase showing similar properties to those described for other Bacillus-related species carboxylesterases like B. subtilis PnbA [59], P. barcinonensis BP-23 EstA [56], or Bacillus sp. BP-7 EstA1 [57]. Nevertheless, these results are also in contradiction with those expected from the PCR prospection procedure because gene lipJ displays similar length and high sequence identity with thermophilic lipases of both, Bacillus and Geobacillus genera, and the gene was isolated from a strain (JR3) bearing proven thermophilic lipolytic activity. To find an explanation for the results obtained, zymogram analysis [31,32] and a bioinformatics approach were addressed.

Zymogram analysis

Zymogram analysis [31,32] of cloned LipJ and concentrated samples of Bacillus sp. JR3 supernatant and cell extracts was performed at 30°C and 60°C on MUF-butyrate (Fig 6). For technical reasons, no higher temperatures were tested. As internal controls, soluble and insoluble fractions of cell extracts from E.coli bearing the same plasmid but without the lipJ insert were analysed (Fig 6, lanes 3 and 4), together with concentrated supernatant samples of B. cereus 131 (Fig 6, lane 2). As shown in Fig 6A, cloned LipJ appears as a band of ca. 40 kDa, whereas supernatants of Bacillus sp. JR3 display a complex lipolytic system including a set of bands of ca. 44, 39, and a fainter band of 22 kDa when assayed at 30°C (Fig 6A). Non boiled cell extracts of strain JR3 show a faint 49 kDa band plus a high molecular weight activity band of ca. 120 kDa probably due to aggregates, a very common trait among lipases [58,60]. It is interesting to note the differences in the activity band pattern of B. cereus (CECT131) and that of strain JR3, showing similar but not identical bands, in agreement with the above data indicating that both strains are close to each other but not identical. When the same samples were analysed in zymograms performed at 60°C [31,32], the band corresponding to LipJ disappeared, indicating loss of activity of the cloned enzyme at this temperature (Fig 6B). On the contrary, both supernatant and cell extract samples of strain JR3 show bands with activity at this temperature. The extracellular 39 kDa band almost disappears but the 44 kDa band still displays high activity at 60°C, confirming the presence of at least an extracellular thermophilic lipase (Fig 6B). Also the 120 kDa and the faint 49 kDa band found in cell extracts of JR3 show higher intensity when assayed at 60°C. The apparent high molecular mass of these bands suggest that they might correspond to the aggregated and unprocessed form of the extracellular thermophilic lipase. The results obtained here confirm the mesophilic profile of LipJ and its differential behaviour in comparison to the extracellular thermophilic lipase activity of strain JR3 and are in agreement with the previous observation of inactivation of LipJ at temperatures over 30°C (Fig 5B). Additionally, the 22 kDa band could be the common low molecular mass lipase of most Bacillus-related species, a well-known ubiquous lipase previously reported [61,62], which has seldom been described as a thermotolerant enzyme [63,64]. On the other hand, considering its size and activity at 60°C, the 44 kDa band could be assigned to the thermophilic activity found in supernatants of the strain JR3, being different from cloned LipJ. Although this band was isolated and analysed by MALDI-TOF, no alpha/beta-hydrolase (neither lipase nor esterase) domains could be identified, except for a fragment corresponding indeed to a section of LipJ sequence.

thumbnail
Fig 6.

Zymogram analysis [31,32] of cell extract and supernatant samples of cloned LipJ and Bacillus JR3, performed at 30°C (A) and 60°C (B). Lane 1: crude cell extract of E.coli expressing LipJ; Lane 2: supernatant of B. cereus 131, used for comparison; Lanes 3–4: soluble (3) and insoluble (4) cell extract of E.coli without the lipJ insert, used as a negative control; Lanes 5–6: concentrated supernatant (5) and crude cell extract (6) of Bacillus strain JR3.

https://doi.org/10.1371/journal.pone.0181029.g006

LipJ sequence and structure analysis

As stated above, the nucleotide sequence of gene lipJ showed a single open reading frame of 1242 bp coding for a hypothetical protein of 413 amino acids, with a predicted molecular mass of 46.2 kDa and a pI of 6.46. A putative Shine-Dalgarno (AGTGA) sequence was found 9 nucleotides upstream the initiation codon [40], along with a -10 (AATTCA) putative promoter region. Three TAA contiguous repeats were found downstream the stop codon but neither inverted repeats nor significant secondary structures that could act as transcription terminators appeared at the available intergenic region, suggesting that other signals located downstream could serve for transcription termination [40].

Being strain Bacillus sp. JR3 closely related to B. cereus and B. thuringiensis group, a BLASTp search for LipJ-homologous proteins in these genera (B. cereus, tax id. 1396 and B. thuringiensis, tax id. 1428) was performed. An annotated lipase (100% coverage and 97% identity) and a hypothetical protein (100% coverage and 93% identity) were found, respectively, both uncharacterized so far. This positions LipJ as the first functionally characterized enzyme with this specific amino acid sequence in the B. cereus group.

A typical carboxylesterase consensus pentapeptide–GXSXG–was found in the deduced amino acid sequence of the cloned enzyme, including Ser138 as a part of the predicted catalytic triad, together with residues Asp340 and His380, assigned by similarity [45,46,65]. Interestingly, an additional pentapeptide-like motif (AAS214FG) bearing an alanine at the first position, like in most secreted Bacillus lipases, was detected. However, further structure analysis of LipJ (see below) positioned this motif far from a putative active site, suggesting that it is not indeed a functional lipase pentapeptide. Although being an infrequent trait among lipases, presence of more than one pentapeptide-like motifs has already been described for Pseudomonas CR611 Lip I.3 acidic lipase [66] and is also found in many Bacillus/Geobacillus-related thermophilic lipase sequences from the databases (UniProt). Also, a bacterial oxyanion hole-like motif (P35IILVNG) was identified [67], which matched well with those of Geobacillus thermophilic lipases [68].

Knowing that most Bacillus-Geobacillus lipases are secreted extracellularly, a search for a signal peptide was performed on the amino acid sequence of LipJ. Curiously, depending on the SignalP program version used [69], different results were obtained: when SignalP 4.1 was used with default settings, no apparent signal peptide existed in LipJ; however, using the default cut-off values of SignalP 3.0 for Gram-positive bacteria, a signal peptide was identified with a probability of 0.994 and the processing point located between amino acids 28 and 29 (AEE-K) [41,70]. This observation could explain the difference in size of recombinant LipJ expressed in E. coli when assayed in zymograms (Fig 6) and the predicted molecular weight.

To get a better knowledge of LipJ properties, we analysed and compared the sequences of related enzymes, extracted from the databases. According to protein domain databases, LipJ is a single domain, globular protein, containing the signature of α/β-fold hydrolases with the consensus pattern _PS00120, unambiguously identified as a lipolytic enzyme [8,71]. This assignment is supported by the high content of non-polar amino acids (hydrophobicity index -0.34) found in the protein sequence [72]. Secondary structure prediction confirmed the typical α/β-fold of carboxylesterases and location of the conserved pentapeptide constituting the “nucleophilic elbow” between strand β3 and the following α helix [2,72].

Assignment of LipJ to a defined family or cluster was performed by inspecting the sequence/function similarities [10], and construction of a phylogenetic tree. LipJ displays significant similarities to the motifs that define the bacterial lipase family I.5, which includes thermophilic lipases from Bacillus, Geobacillus and Staphylococcus species [9,10], suggesting that LipJ might belong to this family. In fact, a phylogenetic tree constructed using the described family I.5 lipases [10,73], positioned LipJ in the same cluster, close to Bacillus/Geobacillus lipases (Fig 7A). Moreover, the database ESTHER, specific for α/β-hydrolases, also assigned LipJ to the Bacan-BA2607 group (Bacterial_lip_FamI.5, including B. anthracis, B. thuringiensis and B. cereus) [51], giving support to this assumption.

thumbnail
Fig 7. LipJ in silico analysis.

(A) Phylogenetic tree of LipJ obtained using MEGA 7 software [50] and the described family I.5 lipases plus the closest Pelosinus fermentans lipase PfL1. The box highlights the position of LipJ, grouping in the same cluster as family I.5 thermophilic lipases. A Rhodococcus sp. CR53 family X lipase [11] was included in the tree for external rooting. (B) 3D model structure obtained for LipJ (purple), aligned and superposed with those of G. stearothermophilus L1 (green) and Pelosinus fermentans PfL1 (grey). The three structures show almost a complete match except for two external β-sheets emerging from the structure of P. fermentans PfL1 (left). The residues involved in catalysis appear at the same position in the three structures and have been highlighted in cyan. Zn and Ca ions (yellow and blue spheres) show the position of the putative binding cavities, which might also be present in LipJ.

https://doi.org/10.1371/journal.pone.0181029.g007

Two validated 3D homology models of LipJ were constructed using pdb 5AH0, a family I.5 lipase from the anaerobic groundwater organism Pelosinus fermentans showing 56.3% sequence identity and 90% coverage [45], and pdb 1KU0, corresponding to L1, a thermoalkaliphilic secreted family I.5 lipase from Geobacillus stearothermophilus [65]. In both cases the 3D models were identical, being validated with 100% residues modelled at >90% confidence. Position of the catalytic triad Ser138, Asp340 and His380, and the putative oxyanion hole (P35IILVHG) [68,74,75] at the expected positions was confirmed. Interestingly, superposition of the 5AH0 and 1KU0 structures with that of LipJ 3D model provided evidence of the large overlapping between the three hydrolases (Fig 7B), indicating that they share a great structural homology. Moreover, putative Ca2+ and Zn2+-binding cavities [45,46,65,76] could also be predicted for LipJ after superposition of the three enzyme structures (Fig 7B), suggesting that LipJ could either be activated or dependent on such ions for activity. In fact, most described thermophilic and thermoalkaliphilic lipases bear either a Ca2+ or Zn2+-binding site, or both (seldom other ions; [12]), acting through a specific net of salt bridges [46,77] that have been shown to be essential for thermophilic activity and thermoresistance [21,65].

Effect of metal ions on LipJ activity

From the previous in silico analysis it was shown that LipJ displays many traits of thermophilic lipases, including its phylogenetic relationship with typically thermophilic family I.5 lipases, the conserved structure and folding of thermophilic lipases, and the presence of putative cavities for accepting ions. Presence of such cavities could indicate that either Ca2+ or Zn2+ (or both) are required for activity at high temperature. This might justify the lack of activity shown by LipJ when assayed over 30°C in the absence of extra ions. To test this possibility, activity assays were carried out at 30 and 60°C in the presence of different concentrations of Ca2+ or Zn2+ or both, and other ions. As shown in Fig 8A, a substantial gain in LipJ activity at 60°C was achieved when the reaction mix was supplemented with Ca2+, obtaining maximum activity at 20 mM Ca2+. However, the highest activity values of LipJ at 60°C with Ca2+ were always lower than those found at 30°C with or without Ca2+, indicating that presence of this ion in the reaction mixture improves in fact the thermal range of LipJ but it does not shift the enzyme to a completely thermophilic behaviour. This was further confirmed when assays were performed in the presence of 20 mM Ca2+ in a range of temperatures from 60 to 100°C. Although an important loss of activity was observed at temperatures above 60°C, LipJ still retained some activity in a wider range of temperatures, showing approximately 65% and 20% residual activity when assayed for 15 minutes at 80 and 100°C, respectively (Fig 8B).

thumbnail
Fig 8. LipJ catalytic behaviour in the presence of Ca2+ and Zn2+ ions.

(A) Activity of LipJ measured at 30 (blue diamonds) and 60°C (red squares) in the presence of different Ca2+ concentrations. (B) Activity of LipJ in the range of temperatures from 60 to 100°C in the presence of 20 mM Ca2+. (C) Loss of activity of LipJ assayed at 60°C in the presence of different Zn2+ concentrations.

https://doi.org/10.1371/journal.pone.0181029.g008

When activity was tested at 60°C in the presence of different concentrations of Zn2+ or both, Ca2+ and Zn2+, complete loss of activity resulted in all cases at Zn2+ concentrations over 0.15 mM, showing only 40% activity at 0.05 mM Zn2+ with respect to that found without ions at 30°C (Fig 8C). When other ions and EDTA were assayed (Fig 9A and 9B), it was shown that Ba2+ and Mn2+ contribute also to increase activity at high temperatures. In fact, a 3-fold activity increase at 80°C was obtained with a combination of 20 mM Ca2+ and 5 mM Mn2+ (Fig 9C). However, neither the enhanced activity at high temperatures produced by Ca2+, nor that of Ba2+ and Mn2+ could completely restore the high activity at 100°C, indicating that cloned LipJ is most probably a different enzyme from that found in the supernatant of Bacillus JR3.

thumbnail
Fig 9.

Effect of different metal ions (assayed at 1 and 10 mM) and EDTA on LipJ activity, measured at 30 (A) and 60°C (B). (C) Enhanced activity of LipJ by a combination of 20 mM Ca2+ and 5 mM Mn2+, assayed at 80°C.

https://doi.org/10.1371/journal.pone.0181029.g009

From the above results we conclude that although LipJ is not responsible for the thermophilic activity found in supernatants of strain JR3, it displays many features of thermophilic lipases like the close phylogenetic proximity to family I.5 lipases, the high similarity of the 3D model structure, plus the observation of lipolytic activity at higher temperatures with the same ions required by thermophilic lipases. Moreover, large amino acid signatures of thermophilic lipases are conserved in LipJ, which was cloned using degenerated primers designed for thermophilic lipases of related genera. Altogether, the results obtained here allow to hypothesize that LipJ could have been in fact a thermophilic extracellular lipase/esterase in ancestral times, when the environmental conditions were more extreme, having evolved to the present mesophilic profile. The change from extreme to mild conditions in the volcanic island of El Hierro could have prompted evolution of Bacillus JR3 towards a mesophilic strain (optimum growth temperature 30°C), yet retaining certain thermophilic traits like the secreted thermophilic lipase/esterase activity observed. In ancestral times, the extracellular thermophilic lipolytic system of Bacillus JR3 (range of temperatures from 30 to 100°C) could have been useful when search for nutrients was harsh and required the presence of duplicated specialised enzymes for survival. However, once the environmental conditions changed, mutations of the enzyme could have been positively selected for mesophilic adaptation, and LipJ (range of temperature from 15 to 40°C) could have evolved from a thermophilic scenario to adapt to the present mild, mesophilic conditions.

Conclusions

We provide here evidence that the newly isolated strain Bacillus sp. JR3 from the Laurisilva Forest of El Hierro Island displays a complex thermophilic and thermostable extracellular lipolytic system, with great potential for industrial applications requiring biocatalysts adapted to harsh temperature conditions. PCR prospection of JR3 genome using degenerated consensus primers for thermophilic lipases allowed cloning and characterization of the new esterase LipJ. The cloned enzyme displays a mesophilic profile, showing preference for short chain-length substrates, with a kinetics profile typical of an esterase. Although being a member of the bacterial lipase family I.5 and bearing several traits of thermophilic lipases, identified in the amino acid sequence and the 3D structure model, LipJ is indeed a mesophilic enzyme. The mixture of thermophilic plus mesophilic features shown by LipJ allows to hypothesise that this enzyme could have evolved from a thermophilic lipase after adaptation to the present mild environment.

Acknowledgments

We thank the Serveis Cientifico-Tècnics of the University of Barcelona for technical support in sequencing. Dr. Mª José Blanco, Chair of the National Institute of Geography at the Canary Islands is also acknowledged for her contribution in sample collection.

Ethical statement

This article does not contain any studies with human participants or animals performed by any of the authors.

References

  1. 1. Bornscheuer UT (2002) Microbial carboxyl esterases: classification, properties and application in biocatalysis. FEMS Microbiol Rev 26: 73–81. pmid:12007643
  2. 2. Jaeger KE, Dijkstra BW, Reetz MT (1999) Bacterial biocatalysts: Molecular biology, three-dimensional structures, and biotechnological applications of lipases. Annu Rev Microbiol 53: 315. pmid:10547694
  3. 3. Ferrer M, Bargiela R, Martínez-Martínez M, Mir J, Koch R, et al. (2015) Biodiversity for biocatalysis: A review of the α/β-hydrolase fold superfamily of esterases-lipases discovered in metagenomes. Biocatal Biotransformation 33: 235–249.
  4. 4. Rauwerdink A, Kazlauskas RJ (2015) How the Same Core Catalytic Machinery Catalyzes 17 Different Reactions: The Serine-Histidine-Aspartate Catalytic Triad of α/β-Hydrolase Fold Enzymes. ACS Catal 5: 6153–6176. pmid:28580193
  5. 5. Khalameyzer V, Fischer I, Bornscheuer UT, Altenbuchner J (1999) Screening, nucleotide sequence and biochemical characterization of an esterase from Pseudomonas fluorescens with high activity towards lactones. Appl Environ Microbiol 65: 477–482. pmid:9925571
  6. 6. Barriuso J, Vaquero ME, Prieto A, Martínez MJ (2016) Structural traits and catalytic versatility of the lipases from the Candida rugosa-like family: A review. Biotechnol Adv 34: 874–885. pmid:27188926
  7. 7. Ollis DL, Cheah E, Cygler M, Dijkstra B, Frolow F, et al. (1992) The α/β hydrolase fold. Protein Eng 5: 197–211. pmid:1409539
  8. 8. Carr P, Ollis D (2009) Alpha/beta hydrolase fold: an update". Protein Pept Lett 16: 1137–48. pmid:19508187
  9. 9. Jaeger K-EE, Eggert T (2002) Lipases for biotechnology. Curr Opin Biotechnol 13: 390–397. pmid:12323363
  10. 10. Arpigny JL, Jaeger KE (1999) Bacterial lipolytic enzymes: classification and properties. Biochem J 343: 177–183. pmid:10493927
  11. 11. Bassegoda A, Pastor FIJ, Diaz P (2012) Rhodococcus sp. strain CR-53 lipR, the first member of a new bacterial lipase family (Family X) displaying an unusual Y-type oxyanion hole, similar to the Candida antarctica lipase clan. Appl Environ Microbiol 78: 1724–1732. pmid:22226953
  12. 12. Castilla A, Panizza P, Rodríguez D, Bonino L, Díaz P, et al. (2017) A novel thermophilic and halophilic esterase from Janibacter sp. R02, the first member of a new lipase family (Family XVII). Enzyme Microb Technol 98: 86–95. pmid:28110668
  13. 13. Drepper T, Eggert T, Hummel W, Leggewie C, Pohl M, et al. (2006) Novel biocatalysts for white biotechnology. Biotechnol J 1: 777–786. pmid:16927261
  14. 14. Kapoor M, Gupta MN (2012) Lipase promiscuity and its biochemical applications. Process Biochem 47: 555–569.
  15. 15. Gupta R, Gupta N, Rathi P (2004) Bacterial lipases: an overview of production, purification and biochemical properties. Appl Microbiol Biotechnol 64: 763–781. pmid:14966663
  16. 16. Gupta R, Kumari A, Syal P, Singh Y (2015) Molecular and functional diversity of yeast and fungal lipases: their role in biotechnology and cellular physiology. Prog Lipid Res 57: 40–54. pmid:25573113
  17. 17. Jaeger K-E, Reetz MT (1998) Microbial lipases form versatile tools for biotechnology. Trends Biotechnol 16: 396–403. pmid:9744114
  18. 18. Angajala G, Pavan P, Subashini R (2016) Lipases: An overview of its current challenges and prospectives in the revolution of biocatalysis. Biocatal Agric Biotechnol 7: 257–270.
  19. 19. Antranikian G, Bornscheuer UT, Liese A (2010) Highlights in Biocatalysis. ChemCatChem 2: 879–880.
  20. 20. Bornscheuer UT, Kazlauskas RJ (2006) Hydrolases in organic synthesis: regio- and stereoselective biotransformations. Wiley-VCH.
  21. 21. Siddiqui KS (2015) Some like it hot, some like it cold: Temperature dependent biotechnological applications and improvements in extremophilic enzymes. Biotechnol Adv 33: 1912–1922. pmid:26585268
  22. 22. Narancic T, Davis R, Nikodinovic-Runic J, O’ Connor KE (2015) Recent developments in biocatalysis beyond the laboratory. Biotechnol Lett 37: 943–954. pmid:25555685
  23. 23. Falcocchio S, Ruiz C, Pastor FIJ, Saso L, Diaz P (2005) Identification of a carboxylesterase-producing Rhodococcus soil isolate. Can J Microbiol 51: 753–758. pmid:16391653
  24. 24. Prim N, Ruiz C, Bofill C, Pastor FIJ, Diaz P (2005) Isolation of lipolytic microorganisms from subtropical soils. cloning, purification and characterization of a novel esterase from strain Pseudomonas sp CR-611. J Biotechnol 118: S120–S120.
  25. 25. Ruiz C, Pastor FIJ, Diaz P (2005) Isolation of lipid- and polysaccharide-degrading micro-organisms from subtropical forest soil, and analysis of lipolytic strain Bacillus sp CR-179. Lett Appl Microbiol 40: 218–227. pmid:15715648
  26. 26. Kouker G, Jaeger KE (1987) Specific and sensitive plate assay for bacterial lipases. Appl Environ Microbiol 53: 211–213. pmid:3103532
  27. 27. Sánchez MM, Fritze D, Blanco A, Spröer C, Tindall BJ, et al. (2005) Paenibacillus barcinonensis sp. nov., a xylanase-producing bacterium isolated from a rice field in the Ebro River delta. Int J Syst Evol Microbiol 55: 935–939. pmid:15774688
  28. 28. Sambrook J, Russell DW (2001) Molecular cloning: A laboratory manual. Cold Spring Harbor Laboratory Press {a}, 10 Skyline Drive, Plainview, NY, 11803–2500, USA.
  29. 29. Weisburg WG, Barns SM, Pelletier DA, Lane DJ (1991) 16S ribosomal DNA amplification for phylogenetic study. J Bacteriol 173: 697–703. pmid:1987160
  30. 30. Laemmli UK (1970) Cleavage of structural proteins during assembly of head of bacteriophage-T4. Nature 227: 680–. pmid:5432063
  31. 31. Diaz P, Prim N, Pastor FIJ (1999) Direct fluorescence-based lipase activity assay. Biotechniques 27: 696–699. pmid:10524309
  32. 32. Prim N, Sánchez M, Ruiz C, Pastor FIJ, Diaz P (2003) Use of methylumbeliferyl-derivative substrates for lipase activity characterization. J Mol Catal B Enzym 22: 339–346.
  33. 33. Altschul SF, Madden TL, Schäffer AA, Zhang J, Zhang Z, et al. (1997) Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucleic Acids Res 25: 3389–3402. pmid:9254694
  34. 34. Bradford MM (1976) Rapid and sensitive method for quantitation of microgram quantities of protein utilizing principle of protein dye binding. Anal Biochem 72: 248–254. pmid:942051
  35. 35. Ruiz C, Falcocchio S, Xoxi E, Javier Pastor FI, Diaz P, et al. (2004) Activation and inhibition of Candida rugosa and Bacillus-related lipases by saturated fatty acids, evaluated by a new colorimetric microassay. Biochim Biophys Acta-General Subj 1672: 184–191.
  36. 36. Bofill C, Prim N, Mormeneo M, Manresa A, Javier Pastor FI, et al. (2010) Differential behaviour of Pseudomonas sp. 42A2 LipC, a lipase showing greater versatility than its counterpart LipA. Biochimie 92: 307–316. pmid:19944735
  37. 37. Box GEP, Hunter WG, Hunter JS (1978) Statistics for experimenters: an introduction to design, data analysis, and model building. Wiley. 653 p.
  38. 38. Notredame C, Higgins DG, Heringa J (2000) T-Coffee: A novel method for fast and accurate multiple sequence alignment. J Mol Biol 302: 205–217. pmid:10964570
  39. 39. Sievers F, Wilm A, Dineen D, Gibson TJ, Karplus K, et al. (2011) Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Mol Syst Biol 7: 539. pmid:21988835
  40. 40. Hall T (1999) BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp Ser 41: 95–98.
  41. 41. Petersen TN, Brunak S, von Heijne G, Nielsen H (2011) SignalP 4.0: discriminating signal peptides from transmembrane regions. Nat Methods 8: 785–786. pmid:21959131
  42. 42. Gasteiger E, Hoogland C, Gattiker A, Duvaud S, Wilkins MR, et al. (2005) Protein Identification and Analysis Tools on the ExPASy Server. The Proteomics Protocols Handbook. Totowa, NJ: Humana Press. pp. 571–607. https://doi.org/10.1385/1-59259-890-0:571
  43. 43. Quevillon E, Silventoinen V, Pillai S, Harte N, Mulder N, et al. (2005) InterProScan: protein domains identifier. Nucleic Acids Res 33: W116–20. pmid:15980438
  44. 44. Finn RD, Clements J, Eddy SR (2011) HMMER web server: interactive sequence similarity searching. Nucleic Acids Res 39: W29–37. pmid:21593126
  45. 45. Biundo A, Hromic A, Pavkov-Keller T, Gruber K, Quartinello F, et al. (2016) Characterization of a poly(butylene adipate-co-terephthalate)-hydrolyzing lipase from Pelosinus fermentans. Appl Microbiol Biotechnol 100: 1753–1764. pmid:26490551
  46. 46. Jeong S-T, Kim H-K, Kim S-J, Pan J-G, Oh T-K, et al. (2001) Crystallization and preliminary X-ray analysis of a thermoalkalophilic lipase from Bacillus stearothermophilus L1. Acta Crystallogr Sect D Biol Crystallogr 57: 1300–1302.
  47. 47. Kelley LA, Mezulis S, Yates CM, Wass MN, E Sternberg MJ (2015) The Phyre2 web portal for protein modeling, prediction and analysis. Nat Protoc 10. pmid:25950237
  48. 48. Magrane M, Consortium U (2011) UniProt Knowledgebase: a hub of integrated protein data. Database (Oxford) 2011: bar009. pmid:21447597
  49. 49. Bernstein FC, Koetzle TF, Williams GJB, Meyer EF, Brice MD, et al. (1978) The protein data bank: A computer-based archival file for macromolecular structures. Arch Biochem Biophys 185: 584–591. pmid:626512
  50. 50. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S (2013) MEGA6: Molecular Evolutionary Genetics Analysis version 6.0. Mol Biol Evol 30: 2725–2729. pmid:24132122
  51. 51. Lenfant N, Hotelier T, Velluet E, Bourne Y, Marchot P, et al. (2013) ESTHER, the database of the / -hydrolase fold superfamily of proteins: tools to explore diversity of functions. Nucleic Acids Res 41: D423–D429. pmid:23193256
  52. 52. Bornscheuer U (2012) From Commercial Enzymes to Biocatalysts Designed by Protein Engineering. Synlett 24: 150–156.
  53. 53. Nazina TN, Tourova TP, Poltaraus AB, Novikova E V, Grigoryan AA, et al. (2001) Taxonomic study of aerobic thermophilic bacilli: descriptions of Geobacillus subterraneus gen. nov., sp. nov. and Geobacillus uzenensis sp. nov. from petroleum reservoirs and transfer of Bacillus stearothermophilus, Bacillus thermocatenulatus, Bacillus th. Int J Syst Evol Microbiol 51: 433–446. pmid:11321089
  54. 54. Quintana-Castro R, Diaz P, Valerio-Alfaro G, Garcia HS, Oliart-Ros R (2009) Gene Cloning, Expression, and Characterization of the Geobacillus Thermoleovorans CCR11 Thermoalkaliphilic Lipase. Mol Biotechnol 42: 75–83. pmid:19107605
  55. 55. Espinosa-Luna G, Sánchez-Otero MG, Quintana-Castro R, Matus-Toledo RE, Oliart-Ros RM (2016) Gene Cloning and Characterization of the Geobacillus thermoleovorans CCR11 Carboxylesterase CaesCCR11, a New Member of Family XV. Mol Biotechnol 58: 37–46. pmid:26603441
  56. 56. Prim N, Blanco A, Martinez J, Pastor FIJ, Diaz P (2000) estA, a gene coding for a cell-bound esterase from Paenibacillus sp BP-23, is a new member of the bacterial subclass of type B carboxylesterases. Res Microbiol 151: 303–312. pmid:10875287
  57. 57. Prim N, Pastor FIJ, Diaz P (2001) Cloning and Characterization of a Bacterial Cell-Bound Type B Carboxylesterase from Bacillus sp. BP-7. Curr Microbiol 42: 237–240. pmid:11178722
  58. 58. Prim N, Bofill C, Pastor FIJ, Diaz P (2006) Esterase EstA6 from Pseudomonas sp CR-611 is a novel member in the utmost conserved cluster of family VI bacterial lipolytic enzymes. Biochimie 88: 859–867. pmid:16600467
  59. 59. Zock J, Cantwell C, Swartling J, Hodges R, Pohl T, et al. (1994) The Bacillus subtilis pnb A gene encoding p-nitrobenzyl esterase: cloning, sequence and highlevel expression in Escherichia coli. Gene 151: 37–43. pmid:7828905
  60. 60. Adlercreutz P, Raku T, Song E, Park S-H, Yoo D, et al. (2013) Immobilisation and application of lipases in organic media. Chem Soc Rev 42: 6406. pmid:23403895
  61. 61. Ruiz C, Blanco A, Pastor FIJ, Diaz P (2002) Analysis of Bacillus megaterium lipolytic system and cloning of LipA, a novel subfamily I.4 bacterial lipase. Fems Microbiol Lett 217: 263–267. pmid:12480114
  62. 62. Ruiz C, Pastor FIJ, Diaz P (2003) Isolation and characterization of Bacillus sp BP-6 LipA, a ubiquitous lipase among mesophilic Bacillus species. Lett Appl Microbiol 37: 354–359. pmid:12969503
  63. 63. Ewis HE, Abdelal AT, Lu C-D (2004) Molecular cloning and characterization of two thermostable carboxyl esterases from Geobacillus stearothermophilus. Gene 329: 187–195. pmid:15033540
  64. 64. Imamura S, Kitaura S (2000) Purification and Characterization of a Monoacylglycerol Lipase from the Moderately Thermophilic Bacillus sp. H-257. J Biochem 127: 419–425. pmid:10731713
  65. 65. Jeong S-T, Kim H-K, Kim S-J, Chi S-W, Pan J-G, et al. (2002) Novel Zinc-binding Center and a Temperature Switch in theBacillus stearothermophilus L1 Lipase. J Biol Chem 277: 17041–17047. pmid:11859083
  66. 66. Panizza P, Syfantou N, Pastor FIJ, Rodríguez S, Díaz P (2013) Acidic lipase Lip I.3 from a Pseudomonas fluorescens-like strain displays unusual properties and shows activity on secondary alcohols. J Appl Microbiol 114: 722–732. pmid:23190193
  67. 67. Jaeger K-EE, Ransac S, Dijkstra BW, Colson C, van Heuvel M, et al. (1994) Bacterial Lipases. Fems Microbiol Rev 15: 29–63. pmid:7946464
  68. 68. Bell PJL, Sunna A, Curach NC, Bergquist PL, Gibbs MD, et al. (2002) Prospecting for novel lipase genes using PCR a. Microbiology 148: 2283–2291. pmid:12177322
  69. 69. Gardy JL, Laird MR, Chen F, Rey S, Walsh CJ, et al. (2005) PSORTb v.2.0: expanded prediction of bacterial protein subcellular localization and insights gained from comparative proteome analysis. Bioinformatics 21: 617–623. pmid:15501914
  70. 70. Emanuelsson O, Brunak S, von Heijne G, Nielsen H (2007) Locating proteins in the cell using TargetP, SignalP and related tools. Nat Protoc 2: 953–971. pmid:17446895
  71. 71. Nardini M, Dijkstra BW (1999) α/β Hydrolase fold enzymes: the family keeps growing. Curr Opin Struct Biol 9: 732–737. pmid:10607665
  72. 72. Fojan P, Jonson PH, Petersen MTN, Petersen SB (2000) What distinguishes an esterase from a lipase: A novel structural approach. Biochimie 82: 1033–1041. pmid:11099800
  73. 73. Lee Lee C.H., Oh T.K., Song J.K., and Yoon MH J.H. (2006) Isolation and Characterization of a Novel Lipase from a Metagenomic Library of Tidal Flat Sediments: Evidence for a New Family of Bacterial Lipases. Appl Env Microbiol 72: 7406–7409.
  74. 74. Holmquist M (2000) Alpha/Beta-hydrolase fold enzymes: structures, functions and mechanisms. Curr Protein Pept Sci 1: 209–235. pmid:12369917
  75. 75. Rengachari S, Aschauer P, Schittmayer M, Mayer N, Gruber K, et al. (2013) Conformational plasticity and ligand binding of bacterial monoacylglycerol lipase. J Biol Chem 288: 31093–31104. pmid:24014019
  76. 76. Kim K, Kim S-J, Chi S-W, Pan J-G, Oh T-K, et al. (2002) Novel Zinc-binding Center and a Temperature Switch in the Bacillus stearothermophilus L1 Lipase* Seong-Tae Jeong, Hyung. 10.1074/jbc.M200640200.
  77. 77. Charbonneau DM, Beauregard M, DeGrado D, Schildbach J, Deal M, et al. (2013) Role of Key Salt Bridges in Thermostability of G. thermodenitrificans EstGtA2: Distinctive Patterns within the New Bacterial Lipolytic Enzyme Family XV. PLoS One 8: e76675.