Selection of Putative Polyester Hydrolases from the Metagenome of Los Humeros Geothermal Field by Means of In Silico Probes

Abstract

Hydrolases are the most popular enzymes, and among the most valuable in biotechnological applications. Some hydrolases, such as lipases, esterases, proteases, cellulases and amylases, are used in the food industry and the production of biopharmaceuticals, biofuels, biopolymers and detergents. Of special interest are those obtained from thermophilic microorganisms. Although there is great microbial diversity in extreme environments, the investigations aimed at detecting and isolating enzymes with potential for polyester degradation such as polyethylene terephthalate (PET) are limited. In this work, we explored the metagenomic library of an oil-enriched soil sample from the “Los Humeros” geothermal field by means of in silico probes in search for enzymes potentially able to degrade polyesters. Using conserved motifs and activity-relevant sites of reported polyester hydrolases, we designed probes that allowed us to identify 6 potential polyester hydrolases in the metagenome. Three-dimensional structure prediction revealed a canonical α/β fold and a cap covering the active site of the enzymes. The catalytic triads were composed of Ser, His and Asp. Structural comparison, substrate binding site analysis and molecular docking suggested their potential as polyester hydrolases, particularly cutinases and PETases. An enzyme, REC98271, was cloned, expressed and characterized, showing thermophilic properties and preference for short-chain substrates. These findings contribute to our understanding of enzyme diversity in “Los Humeros” metagenome and their potential applications in biodegradation and recycling processes.

1. Introduction

Extremophiles are microorganisms that inhabit environments with extreme physicochemical conditions, such as the earth’s poles, glaciers, hot springs, geysers, volcanoes, geothermal fields, deserts, solfatara fields, soda lakes and deep ocean hydrothermal vents, amongst others [1,2]. Of particular interest are the enzymes produced by extremophiles, termed extremozymes, due to their stability and optimal activity under the extreme conditions of their environment, which make them ideal for biotechnological and industrial applications [3]. Examples are the enzymes of thermophilic microorganisms that display activity at high temperatures and show a good thermostability compared to those of mesophilic microorganisms [4].

The difficulty of emulating the extreme conditions necessary for the growth of extremophilic microorganisms means that only a small fraction of extremophiles are cultured and exploited. The development of culture independent “omic” technologies (metagenomics, transcriptomics, proteomics and metabolomics), has helped to overcome this limitation and has facilitated the description of microbial communities, metabolic capabilities and their interactions within, and with, the environment [5].

Metagenomics refers to obtaining and analyzing genome sequences of microbial communities from a particular environment without a previous culturing process [6]. Sequences are usually obtained by massive gene sequencing (next-generation sequencing, NGS) using technological platforms such as Ilumina [7], followed by their assembly in contigs and analysis by binning [8] and alignment (basic local alignment tool, BLAST) for comparing them with known genes deposited in databases [9]. Utilizing bioinformatic tools, the identification of protein domains can be undertaken, as well as the assignment into protein families (Pfam) [10] and the prediction of protein structure, chemical properties, function (Pfam, SwissModel, Robetta, AlphaFold2, UniProt) [11] and phylogenetic location (MEGA, PHYLIP) [12]. These methodologies can be applied to explore new or uncharacterized enzymes by means of the identification of conserved regions, such as protein motifs that might be involved in protein function, and the definition of unique activity-attributed sequences or active sites residues and their variants that might help to assign a function to an unknown gene sequence [13,14]. In this way, thermophilic enzymes with new or superior catalytic abilities can be obtained and used for improved biotechnological or industrial applications where activity at high temperatures and thermostability are desired properties [3,15,16].

Hydrolases are the most popular enzymes and among the most valuable in biotechnological applications. Some hydrolases, such as lipases, esterases, proteases, cellulases and amylases, are extensively used in the food and paper industries and the production of biopharmaceuticals, biofuels, biopolymers and detergents [17]. However, reports related to the detection and isolation of extremozymes capable of degrading polyesters are limited. Polyesters are polymers composed of monomers linked by ester bonds [18]. Plastics are polymers synthetized industrially from fossil fuels, such as polycaprolactone (PCL), or obtained from natural sources such as polyhydroxyalkanoates (PHAs), and might be biodegradable or recalcitrant, depending on their chemical composition and physical and physico-mechanical properties [19]. Biodegradable plastics are polyesters that can be degraded by hydrolysis and then recycled and reused [20]; however, the rate of recycling or incineration is well below the level of production, so it is estimated that more than half ends up in landfills or in the environment [21]. In consequence, new or improved recycling strategies need to be developed. In that respect, enzymatic degradation possesses irreplaceable advantages over chemical depolymerization, especially at high temperatures where the polymer strength is diminished and the solubility and bioavailability of the polymeric substrates are increased [21,22,23].

Polyester hydrolases are classified as carboxylic ester hydrolases (EC 3.1.1), which lipases (EC 3.1.1.3), cutinases (EC 3.1.1.74) and carboxylesterases (EC 3.1.1.1) are grouped as. In recent years, enzymes that hydrolyze polyethylene terephthalate (PETases, EC 3.1.1.101) and enzymes that hydrolyze monohydroxyethyl terephthalate (MHETases, EC 3.1.1.102) have been included in this classification. All these enzymes belong to the α/β hydrolase superfamily and share structural similarities in terms of the central domain and their catalytic machinery constituted by a catalytic triad composed of Ser, Asp and His that might or might not be covered by a lid domain [24,25,26].

In recent years, various microorganisms able to hydrolyze polyesters such as poly(butylene succinate (PBS), poly(butylene adipate-co-terephtalate) (PBAT), poly(lactic acid) (PLA), poly(ε-caprolactone) (PLA) and poly(ethylene terephthalate) (PET) have been isolated from soil, marine seawater, compost, water, sediments, activated sludges, plant leaves and PET residues and characterized. Some of the bacterial and fungal genera reported are Bacillus, Pseudomonas, Marinobacter, Clostridium, Rhodococcus, Thermomonospora, Peribacillus, Priestia, Streptomyces, Acinetobacter, Brevundimonas, Ideonella, Exiguobacterium, Fusarium, Alcanicorax, Thermobifida, Humicula, Rhizopus, Candida, Aspergillus, Penicillium and Moniliophthora; some of the carboxyl esterases identified in the isolates are arylesterases, PETases and MHETases like enzymes, PBAT hydrolases, cutinases, lipases, PCL depolymerases and MHET hydrolases [19,26,27].

The sequences of genes that code for polyester hydrolases and other enzymes of biotechnological interest have also been obtained by direct sequencing of metagenomic environmental samples from human-affected and relatively pristine sites [28], marine sediments [29], industrial effluents [30], industrial sludge [15] and hot spring sediments and have been identified by homology analysis with sequences present in databases [31]. In addition, candidate genes for polyester hydrolases have been identified in metagenomes from both marine and terrestrial environments by using bioinformatics tools such as search algorithms, identification of conserved sequence motifs and phylogenetic analyses in various databases [19,28,32]. Unfortunately, some of these tools and databases can be expensive and might need powerful hardware or computational resources that are not always available to researchers.

Recently, our research group investigated, by a whole-genome shotgun approach, the microbial diversity and the presence of lipolytic enzyme genes from steam soil of “Los Humeros” geothermal field, located in the Trans-Mexican volcanic belt and with surface temperatures between 50 °C and 90 °C. A high percentage of non-characterized lipolytic enzyme genes was observed in the metagenome of an enrichment culture of soil in an oil-rich medium, which suggest the presence of biotechnologically promising organisms and enzymes that deserve to be investigated [33].

Therefore, the purpose of this study was to explore the metagenomic library of the oil-enriched soil sample from the “Los Humeros” geothermal field in search of genes coding for enzymes potentially able to degrade polyesters using probes designed from conserved motifs and activity relevant sites.

2. Results and Discussion

2.1. In Silico Probe Design

In a previous investigation, our research group reported a high percentage of genes coding for non-characterized lipolytic enzymes in the metagenome obtained from a soil sample from the “Los Humeros” geothermal field (15 cm depth, 80 °C), enriched in a high-oleic acid safflower oil medium [33]. In order to identify new potential polyester hydrolases in the metagenome, we designed probes directed to conserved motifs of polyester hydrolases with experimentally confirmed hydrolytic activity, reported structural data and biochemical characterization (Table 1).

First, using the MultiAlin multiple sequence alignment server, we analyzed the amino acid sequences of seven polyester hydrolases: a depolymerase from Acidovorax delafieldii (Q8RR62), a PETase from Pseudomonas aestusnigri (6SBN), a cutinase from Thermobifida alba (D4Q9N1), two cutinases from Thermobifida cellulosilytica (5LUI and 5LUJ), a PETase from Ideonella sakaiensis (IsPETase) (6ANE) and a cutinase from a metagenome (4EB0) (Figure 1A). IsPETase sequence was used to locate the conserved and distinctive motifs of ester hydrolases, which were as follows: the catalytic triad, which is composed of serine, aspartic or glutamic acid and histidine, and is responsible for the nucleophilic attack on the carbonyl carbon and subsequent hydrolysis of the ester bond; the pentapeptide G-X1-S-X2-G, which includes the catalytic serine; the oxyanion hole, composed of the amino acids of the active site that stabilize the reaction intermediate; and the conserved structure named the “nucleophilic elbow”, where the catalytic Ser is located. Ester hydrolases might also possess a lid covering the active site and Cys residues forming disulfide bridges that give thermostability to the enzyme [40]. In the aligned sequences, seven conserved motifs were identified (Figure 1A): a sequence near the amino terminal (motif 1), a sequence rich in glycine that is part of the oxyanion hole (motif 2), a sequence rich in threonine (motif 3), the pentapeptide (motif 4), the sequence surrounding the catalytic aspartate (motif 5), the sequence surrounding the catalytic histidine (motif 6) and the sequence that contains a cysteine implicated in disulfide bond formation (motif 7). A logo for each motif was created via the online tool Skylign (http://skylign.org/, accessed on 4 April 2022), which included the alternative residues that can be found in some positions (Figure 1B).

All the possible sequence combinations of motifs 2, 4, 5, 6 and 7 were used as probes to search for ester hydrolases within the “Los Humeros” translated metagenomic library. The probes that were successful for ester hydrolases identification were GHSMGG, which matched nine contigs, and GWSMGG, which matched two contigs. The two probes were composed of alternative sequences of motif 4 derived from the pentapeptide sequence. Each contig contained one protein coding gene.

2.2. Protein Identity

The identity of the proteins recognized by the two probes derived from motif 4 was determined using the online Basic Local Alignment Search Tool (blastn and blastp suites). Results are shown in Table 2. The proteins were composed of 250–300 residues in coincidence with their corresponding proteins in the BLAST database. All the proteins were identified as hydrolases, with similarities ranging from 25 to 61% with the following enzymes: four bacterial phospholipases, five α/β hydrolases (three of bacterial origin and two unassigned), one uncharacterized hydrolase (unassigned) and one methyl ester esterase of archaeal origin. The enzymes classified as hydrolases (46241, 48180, 48200, 84022, 98271, 25914 and 41677) were selected for 3D structure analysis.

The use of probes designed according to conserved regions of ester hydrolases, such as the pentapeptide, has been useful to find ester hydrolases in other metagenomes. For example, Danso et al. (2018) [32] used probes based on the pentapeptide motif, on amino acids for oxyanion hole and aromatic clamp formation and regions containing Cys residues, to analyze 108 marine and 25 terrestrial metagenomes from the Integrated Microbial Genome (IMG) database, detecting 349 putative PET hydrolases. The enzymatic activity of four enzymes was experimentally confirmed. In another investigation, Saini et al. (2022) [41] searched for polyester hydrolases in the BLASTP database, using the pentapeptide as a probe, with Trp in X1 and a Cys near the active site. They selected five PETases whose activity over MHET was confirmed by molecular docking.

2.3. Structural Analysis

The three-dimensional structures of the selected enzymes (46241, 48180, 48200, 84022, 98271, 25914 and 41677) were predicted by Alphafold2. All the enzymes showed the canonic α/β-hydrolase fold with a lid covering the active site (Table 3), which are distinctive characteristics of hydrolytic enzymes. The α/β fold is generally composed of a parallel β-sheet of eight beta-strands, connected by α-helices located on the sides of the β-sheet. The lid is usually present in ester hydrolases and is composed of α-helices connected to the core enzyme by a flexible structure that uncovers the active site and enables the entrance of the substrate [40].

The catalytic triads of the active site were all composed of Ser, His and Asp. A Trp residue was positioned at the nucleophilic elbow next to the catalytic Ser in enzymes identified as 25914 and 41677, while a His residue was observed in enzymes 46241, 48200, 84022, 48180 and 98271. No Cys residues likely to form disulfide bonds were found in any of the enzymes. The oxyanion cavities were composed mainly of hydrophobic residues, and their sizes ranged from 51.60 to 376.48 ų. It has been reported that the presence of a Trp (W) at the nucleophilic elbow next to the catalytic Ser (GWSMGG) has the effect of extending the hydrophobic surface adjacent to the active site for an efficient binding of polyester substrates. His (H), despite being more voluminous, has also been located in that position (GHSMGG) in several polyester hydrolases [20].

In order to compare the positions and spatial organization of the catalytic triad residues, the 3-D structures of the selected enzymes were overlapped with four polyester hydrolases’ templates: two cutinases from Thermobifida cellulosityca (PBD ID 5LUJ and ID 5LUI) [37], a cutinase from a compost metagenome (PBD ID 4EB0) [39] and a PETase from Ideonella sakaiensis (PBD ID 6ANE) [38] (Figure 2). The comparation of the catalytic triads’ arrangements with their best overlapping polyester hydrolase is shown in Figure 3 and Table 4.

The catalytic triad of the enzymes identified as 25914, 48180 and 84022 showed the best coincidence with the catalytic triad of a cutinase isolated from a compost metagenome (4EB0) (Figure 2A–C), with a root mean square deviation of atomic positions (RMSD) of 0.183 Å, 0.172 Å and 0.229 Å, respectively (Figure 3A–C and Table 4). The catalytic triad of the enzyme 41677 best overlapped with the catalytic triad of the cutinase of Thermobifida fusca (5LUI), and the catalytic triad of 46241 best overlapped with the catalytic triad of the cutinase of Thermobifida cellulosityca 5LUJ (Figure 2D,E), with a RMSD of 0.088 Å and 0.388 Å, respectively (Figure 3D,E and Table 4). The catalytic triad of the enzyme 98271 showed the best superposition with the catalytic triad of the PETase of Ideonella sakaiensis (6ANE) (Figure 2F), with a RMSD of 0.138 Å (Figure 3F and Table 4). Finally, the catalytic triad of the enzyme 48200 showed no match with any of the templates evaluated. It is important to emphasize that an RMSD < 1 indicates a high similarity in the residues’ arrangements of the regions that are being compared, which suggests that the six selected enzymes could be polyester hydrolases (five cutinases and one PETase).

2.4. Identification of Substrate Binding Site Conformations

Using molecular docking in the UCSF Chimera platform, we predicted the recognition and interaction of the catalytic triad of the putative polyester hydrolases with two substrates: a dimer of the aliphatic polyester polycaprolactone (PCL) (Figure 4A) and a trimer of the aromatic polyester polyethylene terephthalate (PET) (Figure 4B).

Substrate binding site analysis revealed an interaction between the catalytic serine and the carbonylic carbon (Cα) in the ester bond of PET in the enzymes 25914, 41677, 46241, 84022, 98271 and 48180, while in 25914, 46241, 84022 and 98271 the catalytic serine also interacted with the ester bond of PCL. The distances of these interactions were between 2.6 Å and 4.0 Å in all cases (Table 5), which means that they are chemically relevant for the hydrolysis to occur. Likewise, distances less than 3.9 Å were observed between the catalytic serine and histidine, and distances no greater than 3.0 Å were observed between histidine and aspartic acid of the catalytic triads (Table 5), indicating a high probability of an effective interaction between the catalytic residues [42]. Molecular docking also indicated the presence of hydrophobic interactions and hydrogen bond formation between the oxyanionic cavity and the substrate. These results might confirm that the selected enzymes are polyester hydrolases. Molecular docking of the putative polyester hydrolase 98271 using PET and PCL as substrates is shown in Figure 5 as an example. The enzyme 98271 was selected for a further characterization due to its structural similarity to IsPETase.

2.5. Gene Cloning, Expression and Characterization of Enzyme 98271

The gene coding for the enzyme 98271 was cloned in the pHTP8 vector (869 pb, Figure 6A), and the recombinant enzyme REC98271 was expressed and characterized. The enzyme was successfully overexpressed, particularly in the intracellular compartment; it possesses a molecular weight of 46.9 kDa (32.4 kDa from 98271 enzyme plus 14.5 kDa from vector’s thioredoxine), and its activity was confirmed by a zymogram over MUF-butyrate (Figure 6B). The enzyme was purified by affinity chromatography (Figure 6C) and characterized (Figure 7 and Figure 8).

The enzyme REC98271 showed a preference for short-chain substrates (C2–C8) at pH 6.0 (Figure 7), as carboxylesterases do. As shown in Figure 8A, maximum specific activity was obtained at 50 °C (1.72 U mg⁻¹), indicating its thermophilic nature. These properties are similar to other thermophilic polyester hydrolases, such as LC-cutinase (50 °C, C4, pH 8.5), obtained from a metagenome [39], and Tfac (60 °C, C4, pH 6.0) from Thermobifida fusca [43]. It is important to note that REC98271 showed a 54.9% similarity with a non-characterized hydrolase from Anoxybacillus caldiproteolyticus U458, a thermophilic bacterium that grows optimally at 60 °C, pH 6.5 and at 0–0.5% salinity and that was originally isolated from a sewage sludge [44].

After incubating the enzyme in the presence of metal ions for 1 h (Figure 8B), Hg²⁺ decreased up to 95% of residual activity, probably due to structural changes derived from its interaction with the histidine at the active site [45]. Ca²⁺ decreased activity by 42%, which may be due to the interference of potassium with the reaction buffer that contributes to activity; the activity increase in the presence of Ba²⁺ (divalent) and Li¹⁺ (monovalent) (55% and 25%, respectively) might be the result of an interaction with cation binding sites that provide structural stability [46,47].

The enzymatic activity of REC98271 was not considerably affected by the presence of organic solvents (Figure 8C) or detergents (Figure 8D). The increase in activity observed with Triton X-100, Tween-80 and β-mercaptoethanol can be attributed to their disassembling effects over enzyme aggregates, a common phenomenon observed in lipolytic enzymes [48]. Only ethanol and acetone decreased activity by approx. 40%, which is comparable to what has been reported for the IsPETase from Ideonella sakaiensis, whose activity decreased by 30% in the presence of ethanol [46]. These results suggest that REC98271 enzyme is a robust enzyme applicable in biodegradation processes of contaminated soils and effluents, in which these substances might be present.

As shown in Figure 8D, the enzyme REC98271 was inactivated by PMSF, confirming the essential role of the serine residue in the catalytic activity of REC98271. EDTA showed no interference with the activity, indicating that the enzyme is not a metalloprotein.

In summary, by means of in silico analysis and with the aid of bioinformatic tools, we performed a metagenome screening and identified potential polyester hydrolases for further research and experimental validation. It is worth mentioning that the enzymes identified in our study have not been previously reported, since they appear to be classified only as α/β hydrolases in the databases. The enzymes showed a high similarity in sequence and in the active site and nucleophilic elbow conformations to the main polyester hydrolases reported so far, and molecular docking analysis revealed relevant interactions between enzymes and the polyesters PET and PCL. The selected enzyme for experimental validation (98271) showed hydrolytic activity against p-nitrophenyl esters of short chain length at 50 °C and in the presence of organic solvents and detergents, which make it interesting for its application in biotechnological and industrial processes. The experimental validation of its activity against polyesters is currently being carried out in the laboratory. Finally, we want to emphasize the user-friendliness of our bioinformatic approach, since it included only free access databases and software, and no sophisticated computer equipment was needed.

3. Materials and Methods

3.1. In Silico Probe Design

In order to look for new enzymes with the ability to degrade polyesters we analyzed the metagenomic library (5.3 GB, 109,000 contigs) of a soil sample obtained from a steaming location at “Los Humeros” geothermal field (15 cm depth, 80 °C) that was enriched in a high-oleic acid safflower oil medium [33]. The search was undertaken using probes that were designed to specifically recognize the amino acids’ sequences of conserved motifs of polyester hydrolases with experimentally confirmed hydrolytic activity, reported structural data and biochemical characterization [48]. The seven polyester hydrolases that were used for probes’ design are shown in Table 1.

The amino acids sequences of the selected polyester hydrolases were aligned using the multiple sequence alignment server MultAlin (http://multalin.toulouse.inra.fr/multalin/, accessed on 28 March 2022). The sequence of the PETase IsPETAse from Ideonella sakaiensis was used as reference for the identification of the pentapeptide, the catalytic triad and the oxyanion hole. After detailed analysis and comparisons, seven conserved regions were found; the ones corresponding to the nucleophilic elbow and the sequences surrounding aspartic acid and histidine of the catalytic triad were used to design nine probes that were assayed for polyester hydrolases detection in the translated metagenomic library with the MEGAX software version 10.2.6.

3.2. Enzyme Identity and 3D Structure Analysis

The translated contigs that were recognized by the probes were analyzed using the online Basic Local Alignment Search Tool, BLAST (https://blast.ncbi.nlm.nih.gov/Blast.cgi, accessed on 11 May 2022) to obtain protein identity and percentage of similarity. The tridimensional structures of the possible polyester hydrolases found were predicted by Alphafold2 (https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb, accessed on 11 May 2022).

The models were validated using MolProbity (http://molprobity.biochem.duke.edu/, accessed on 17 May 2022), Saves 6.0 (https://saves.mbi.ucla.edu/, accessed on 17 May 2022) and PDBsum (http://www.ebi.ac.uk/thornton-srv/databases/pdbsum/Generate.html, accessed on 17 May 2022). Incomplete sequences were discarded.

3.3. Active Site Analysis

For active site analysis, we used four PDB templates obtained from the Protein Data Bank (RCSB-PDB) (https://www.rcsb.org/, accessed on 25 May 2022) of two cutinases from Thermobifida cellulosityca with PBD IDs 5LUI and 5LUJ [35]; the compost metagenome cutinase with PBD ID 4EB0 [36]; and the PETase from Ideonella sakaiensis with PBD ID 6ANE [39]. The superposition of the templates with the structures of the putative polyester hydrolases was performed through the PyMOL align tool that expresses the structural homology of the active sites as the root mean square deviation of atomic positions (RMSD) [49].

3.4. Identification of Substrate Binding Site Conformations (Molecular Docking)

To predict the recognition and interaction of the putative polyester hydrolases with their substrate(s), molecular docking was carried out. The models of the selected enzymes were parameterized and minimized in UCSF Chimera (AMBER force field ff14SB), as well as the substrates, a dimer of the aliphatic polyester polycaprolactone (PCL) and a trimer of aromatic polyester polyethylene terephthalate (PET). The coordinates center (8.9, −1.8, −3.0) and size (20, 22.9, 26) were used to design the box, which included the active site of the enzymes. In the docking models, we determined the distance between the catalytic Ser residue and the ester bond of the substrate (<5 Å for a relevant interaction to occur) and analyzed the binding affinity, the hydrogen bond formation and the number of interactions.

3.5. Expression and Characterization of 98271 Recombinant Enzyme (REC98271)

3.5.1. Heterologous Expression

The gene identified as 98271 was synthesized by the company Catalysis in the plasmid pD864. Subsequently, primers were designed to transfer the gene to the plasmid pHTP8, resulting in the recombinant vector pHTP8-98271. Competent cells of the E. coli BL21 strain were prepared with 100 mM CaCl₂ and transformed. Colonies were incubated on LB agar with 50 µg/mL kanamycin. Transformants were inoculated into LB medium at 37 °C until the culture reached an optical density (OD) of 0.8. Then, 1 mM IPTG was added to induce protein expression. Cells were collected by centrifugation after 12 h of induction at 37 °C. The cells were washed twice with sterilized water by centrifugation (12,879× g for 15 min at 4 °C), and the cell pellet was resuspended in a wash solution (300 mM NaCl, 50 mM Na₂HPO₄ and 5 mM imidazole, pH 8.0). The cells were lysed by sonication and subsequently centrifuged to obtain the crude enzyme (12,879× g for 15 min at 4 °C).

3.5.2. Recombinant Enzyme (REC98271) Purification

REC98271 enzyme was purified in an IMAC resin, which was packed and loaded with chelating metal (Ni) as indicated in the Profinity™ IMAC Bio-Rad user manual (Hercules, CA, USA). The IMAC resin, loaded with metal and pre-equilibrated with 10 column volumes (CV) of buffer (50 mM NaH₂PO₄, 300 mM NaCl and 5 mM imidazole at pH 8.0), was used to load the sample (10 CV of total protein at 16 mg/mL resin). The column was washed with 15 CV of the same buffer to elute unbound components. Elution of bound proteins was achieved using the elution buffer (50 mM NaH₂PO₄, 300 mM NaCl and 400 mM imidazole, pH 8.0). Flow and wash fractions were collected in fractions of 500 µL, and the concentration was directly measured in the nanodrop until no protein was detected. To estimate the molecular mass of the recombinant protein, samples were analyzed by 12% SDS-PAGE (Bio-Rad, Hercules, CA, USA).

3.5.3. Enzymatic Activity Analysis

After expression and purification, the enzyme REC98271was biochemically characterized. Activities was measured by the technique reported by Nawani et al. (1998) [50]. This technique employs p-nitrophenyl esters as substrates, which, upon hydrolysis, generate a colored compound (p-nitrophenol), which is measured spectrophotometrically at 420 nm. The reaction mixture consisted of 100 μL of purified enzyme (0.05–0.1 mg protein), 800 μL of 0.05 M potassium phosphate buffer (pH 6.0) and 100 μL of 0.01 M p-nitrophenyl esters (C2-C12 in absolute ethanol). It was incubated at the corresponding T for 30 min, adding 250 μL of 0.1 M Na₂CO₃ at the end. The mixture was centrifuged at 12,879× g for 15 min at 4 °C in an Eppendorff^® 5415R microcentrifuge (Westbury, New York, USA), and absorbance at 420 nm was measured in a spectrophotometer (SmartSpec^TM 3000, Bio-Rad, Hercules, CA, USA). A blank without enzyme was used as a control. Each experiment was performed in triplicate. Enzymatic activity was calculated according to the molar extinction coefficient of p-nitrophenol at 405 nm (1.1591 × 10⁻⁶ μM⁻¹ cm⁻¹). One enzyme unit is defined as the amount of enzyme that releases 1 μM of p-nitrophenol under the assay conditions in 30 min.

3.5.4. REC98271 Enzyme Characterization

To determine the substrate chain length preference of the enzyme, the activity assay described in Section 3.5.3. was be performed at different pH values (5.0, 6.0, 7.0, 8.0) with p-nitrophenyl esters of different chain length (C:2, C:4, C:5, C:8, C:10, C:12) in each case. The effect of temperature on REC98271 enzyme activity was determined at 30, 40, 50, 60, 70 and 80 °C. The pH of the reaction buffer was adjusted to pH 6.0 based at each assay temperature. The effect of pH on enzyme activity was determined with 0.05 M sodium acetate (pH 5) and 0.05 M potassium phosphate (pH 6, 7 and 8). In all cases, the pH was adjusted to the temperature of maximum activity. To evaluate the effect of metal ions on enzyme activity, aliquots of 500 μL of enzyme and 500 μL of an aqueous solution of 1 mM of CaCl₂, KCl, MgCl₂, NaCl, BaCl₂, HgCl₂ and LiCl were incubated for 1 h at 30 °C, and then the activity was evaluated. To evaluate the effect of inhibitors and detergents on enzyme activity, aliquots of 500 μL of enzyme and 500 μL of an aqueous solution of EDTA (1 mM), 1% v/v β-mercaptoethanol, Triton X-100^® 1% (v/v), Tween 80^® 1% (v/v), and 0.1% w/v SDS (sodium dodecyl sulfate) were incubated for 1 h at 30 °C, followed by the lipolytic activity assay. To assess enzyme stability in the presence of organic solvents, an incubation of 0.5 mL of enzyme with 0.5 mL of solvent (methanol, ethanol, butanol, 2-propanol, acetone) was performed for 1 h at 30 °C, and then the residual activity was determined. The activity was expressed as relative activity (%).

4. Conclusions

Through the use of probes based on the pentapeptide conserved site (GWSGGG and GHSGGG) of lipases, esterases and PETases, it was possible to identify six enzymes with probable activity on polyesters, since they possess the canonical α/β hydrolase structure, the three-dimensional configuration of the active site and can interact in silico with aromatic and/or aliphatic polyesters (polyethylene terephthalate (PET) and polycaprolactone (PCL)). One of them, REC98271 enzyme, was cloned, expressed and characterized, resulting in a thermophilic carboxyl ester hydrolase, tolerant to the presence of detergents and organic solvents, with potential application in biodegradation processes of contaminated soils and effluents.

The identification of carboxyl ester hydrolases in the metagenomic library obtained from an enriched soil sample of “Los Humeros” geothermal field allowed us to highlight the potential of these extreme environments as a source of enzymes applicable to biotechnological processes. An example is the degradation of various forms of polyesters such as plastics and microplastics that can be found all over the world and possess serious risks to the environment and to humanity.

Author Contributions

Conceptualization, R.M.O.-R. and C.P.-M.; methodology, R.S.-P. and R.M.O.-R.; software, G.E.-L. and M.G.S.-O.; validation, R.Q.-C. and M.G.S.-O.; formal analysis, R.S.-P. and R.M.O.-R.; investigation, R.S.-P. and R.M.O.-R.; resources, R.M.O.-R. and C.P.-M.; writing-original draft preparation, R.S.-P. and R.M.O.-R.; writing—review and editing, R.M.O.-R. and C.P.-M.; supervision, G.E.-L., M.G.S.-O. and R.Q.-C.; project administration, R.M.O.-R. and C.P.-M. All authors have read and agreed to the published version of the manuscript.

Data Availability Statement

Data are contained within the article.

Acknowledgments

Rocio Solis-Palacios acknowledges the National Council of Humanities, Sciences and Technologies (CONAHCyT-México) for her scholarship grant 859073.

Conflicts of Interest

The authors declare no conflicts of interest.

Footnotes

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Figures and Tables

Figure 1. (A) Alignment of the amino acid sequences of the polyester hydrolases used for the identification of conserved regions and positions of the relevant substrate-binding residues and for in silico probes design. The boxes indicate the conserved regions identified. (B) Logos of the conserved regions created via the online tool Skylign (http://skylign.org/, accessed on 4 April 2022).

Figure 2. Structural overlap of the three-dimensional models of the selected sequences (in light blue with the catalytic triad in dark blue) with templates of polyester hydrolases with confirmed activity (in green with the catalytic triad in red). (A) Cutinase from a compost metagenome superposed with the enzyme 25914. (B) Cutinase from a compost metagenome superposed with enzyme 48180. (C) Cutinase from a compost metagenome superposed with enzyme 84022. (D) Thermobifida fusca cutinase superposed with the enzyme 41677 (E) Thermobifida cellulosityca cutinase superposed with the enzyme CI 46241. (F) Ideonella sakaiensis PETase superposed with enzyme 98271.

Figure 3. Superposition of the catalytic triads of the polyester hydrolases (red) and the selected enzymes from the “Los Humeros” metagenome (blue). Cutinase 4EB0 aligned with (A) 25914, (B) 48180 and (C) 84022. Cutinase 5LUI aligned with (D) 41677. Cutinase 5LUJ aligned with (E) 46241. PETase 6ANE aligned with (F) 98271. The positions of the catalytic triad residues of the predicted enzymes are indicated in bold.

Figure 4. Substrates used in molecular docking. (A) Polycaprolactone (PCL). (B) Polyethylene terephthalate (PET).

Figure 5. Active site of 98271. Catalytic triad and the coupling model of the reaction intermediate of (A) PET and (B) PCL. The three residues Ser104, Asp288 and His256 that form the catalytic triad are shown as cyan bars and are appropriately labeled. The interaction distances between the catalytic serine and Cα are also indicated. The structure of 98271 is shown as a cartoon model. The docking pattern of PET and PCL is shown as an orange bar and the interaction site is highlighted as a green box.

Figure 6. Electropherograms. (A) 3% agarose gel showing a band corresponding to the PCR-amplified 98271 gene (869 bp). M: molecular weight markers (NZYDNA Ladder III). 1: Band corresponding to the PCR-amplified 98271 gene. (B) 12% SDS-PAGE gel showing the band corresponding to the REC98271 enzyme after the expression induction with IPTG. M: molecular weight markers (BioRad Dual color), S: soluble fraction, P: insoluble fraction, S1: activity in MUF-Butyrate in soluble fraction, P1: activity in MUF-Butyrate in insoluble fraction. (C) 12% SDS-PAGE gel showing protein patterns after purification of REC98271 enzyme in affinity chromatography. M: molecular weight markers (BioRad Dual color), E: crude extract, F1–F8: eluted fractions (1–8) with 400 mM imidazole.

Figure 7. Relative activity of the enzyme REC98271 on various p-nitrophenyl esters (C-2: p-nitrophenyl acetate, C-4: p-nitrophenyl butyrate, C-5: p-nitrophenyl valerate, C-8: p-nitrophenyl caprylate, C-10: p-nitrophenyl decanoate, C-12 p-nitrophenyl laurate) at pH 5.0, 6.0. 7.0 and 8.0. Activities are expressed as the percentage of that of hydrolyzed p-nitrophenyl butyrate (C4) at pH 6, which was taken as 100%. Different letters (a–h) indicate significant differences between activity measurement conditions, determined by ANOVA-Tukey test (p < 0.005). Results are expressed as mean ± standard deviation of triplicates.

Figure 8. Biochemical characterization of the enzyme REC98271. (A) Effect of temperature on enzymatic activity (30 °C to 80 °C). (B) Effect of ions (1 mM) on enzymatic activity. (C) Effect of organic solvents on enzymatic activity. (D) Effect of inhibitors on enzymatic activity. Enzymatic activity was evaluated in 50 mM potassium phosphate buffer (pH 6.0) using p-nitrophenyl butyrate as substrate at 50 °C (except for the effect of temperature). Different letters (a–g) indicate significant differences between activity measurement conditions, determined by ANOVA–Tukey test (p < 0.005). Results are expressed as mean ± standard deviation of triplicates.

Table 1

Polyester hydrolases used to design probes directed to conserved sites’ recognition.

UniProt or PDB ID	Source	Reference
Q8RR62	Acidovorax delafieldii	Uchida et al. (2002) [34]
6SBN	Pseudomonas aestusnigri	Bolliger et al. (2020) [35]
D4Q9N1	Thermobifida alba	Hu et al. (2010) [36]
5LUI	Thermobifida cellulosilytica	Ribitsch et al. (2016) [37]
5LUJ	Thermobifida cellulosilytica	Ribitsch et al. (2016) [37]
6ANE	Ideonella sakaiensis	Yoshida et al. (2016) [38]
4EB0	Metagenome	Sulaiman et al. (2012) [39]

Table 2

Enzyme identity of the sequences recognized by the two probes derived from motif 4.

Contig	Organism	Protein Identity	Blast Database (# aa)	Los Humeros Metagenome (# aa)	Identity (%)
20637	Anoxybacillus	Phospholipase YtpA	259	265	61.09
27123	Aerybacillus pallidus PI8 DNA	Phospholipase YtpA	259	259	60.23
46241	Unassigned	Hydrolase α/β	271	280	27.24
48180	Aeribacillus pallidus KCTC3564	Hydrolase α/β	273	266	59.46
48200	Archaeon	Pimeloyl-[acyl carrier protein] methyl ester esterase	254	243	26.75
49166	Parageobacillus thermoglucosidasius	Phospholipase YtpA	259	263	61.09
84022	Thermogemmatispora	Hydrolase α/β	258	271	25.88
98271	Anoxybacillus caldiproteoliticus U458	Uncharacterized hydrolase Yug	273	280	54.98
102392	Anoxybacillus caliroteolyticus U458	Phospholipase YtpA	259	265	58.37
25914	Unassigned	Hydrolase α/β	286	301	25.34
41677	Geobacillus sp. GHH01	Hydrolase α/β	272	297	26.1

# aa: number of amino acids.

Table 3

Analysis of the canonical structure of carboxylic ester hydrolases selected from the metagenome.

Enzyme	α/β Hydrolase Domain		Lid Domain	Catalytic Triad
	# α-Helix	# β-Sheet	# α-Helix	Residues Position
25914	8	8	5	Ser100-Asp249-His278
41677	8	8	5	Ser100-Asp249-His278
46241	8	8	4	Ser119-Asp230-His258
84022	7	6	4	Ser78-Asp187-His215
98271	8	8	6	Ser104-Asp228-His256
48180	8	7	6	Ser94-Asp218-His246
48200	8	7	4	Ser82-Asp192-His220

#: number.

Table 4

Enzyme templates used for superposition analysis in PyMOL and the resulting root mean square deviation of atomic positions (RMSD).

Enzyme	Templates	RMSD (Å)
25914	Cutinase 4EB0	0.172
41677	Cutinase 5LUI	0.088
46241	Cutinase 5LUJ	0.388
84022	Cutinase 4EB0	0.229
98271	PETase 6ANE	0.138
48180	Cutinase 4EB0	0.183

Table 5

Distances between the residues of the catalytic triad.

Enzyme	Distance (Å)Ser-His	Distance (Å)His-Asp	Distance (Å)Ser-Asp
25914	3.3	2.8	7.2
41677	3.4	2.8	7.2
46241	3.0	3.0	7.1
84022	3.8	2.9	7.2
98271	3.8	2.9	7.0
48180	3.7	2.9	7.0

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