Gene Cloning, Purification, and Characterization of a Cold-Active Alkaline Lipase from Bacillus cereus U2

Abstract

Lipases are important industrial enzymes with a wide range of applications across various sectors. Cold-active lipases are particularly well suited for industrial processes that operate at low temperatures (such as food processing and environmental remediation) due to their high catalytic efficiency and energy-saving benefits. In this study, a novel lipase—LipU (GenBank accession: PV094892)—was heterologously expressed from Bacillus cereus U2 and characterized for its low-temperature adaptability and alkaline resistance. LipU belongs to the lipase Subfamily I.5 and shares the highest amino acid sequence identity (53.32%) with known homologs. Enzymatic assays revealed that LipU exhibits optimal activity at 20 °C and pH 11. It retained 95% of its initial activity after 24 h of incubation at 4 °C and pH 11.0. Furthermore, the activity of LipU was enhanced by Ca²⁺, Na⁺, Tween 20, and Tween 80, whereas it was inhibited by Cu²⁺, Zn²⁺, Mn²⁺, and sodium dodecyl sulfate (SDS). LipU demonstrated tolerance to various organic solvents of differing polarity; after 1 h of exposure to 15% (v/v) ethanol, n-butanol, isoamyl alcohol, dimethyl sulfoxide, or glycerol, it retained over 78.6% of its activity. These properties make LipU a promising candidate for industrial applications, including for leather degreasing, alkaline wastewater treatment, and low-temperature biocatalysis.

1. Introduction

Lipases (EC 3.1.1.3), also known as triacylglycerol hydrolases, are members of the serine hydrolase superfamily. In addition to hydrolyzing carboxylic acid esters, lipases exhibit diverse catalytic activities, including acidolysis, alcoholysis, ammonolysis, and transesterification [1,2]. The majority of currently characterized microbial lipases are derived from bacteria and fungi [3]. Lipase-producing microorganisms primarily include Bacillus, Psychrobacter sp., Pseudomonas sp., Serratia, Rhizopus, and Aspergillus. Compared to fungal lipases, bacterial lipases are generally easier to produce through fermentation, yielding higher outputs at lower production costs [4]. Additionally, bacterial lipases demonstrate greater stability in organic solvents, along with enhanced catalytic activity and substrate specificity [5]. As a result, bacterial lipases are widely utilized in food processing, medicine, environmental management, and bioenergy applications [6].

The genus Bacillus is a well-known source of industrial enzyme-producing microorganisms, and its species are widely distributed in soil, water, and extreme environments, such as cold habitats. Bacillus cereus is capable of producing cold-active lipase, and its active site can adapt to high-pH conditions through surface charge regulation. Furthermore, although Bacillus cereus is not typically considered a psychrophilic bacterium, studies have reported successful expression of a low-temperature lipase gene, suggesting that Bacillus cereus strains isolated from cold environments can acquire low-temperature adaptation through genetic transfer [7].

Cold-active lipases are a class of enzymes that exhibit a high catalytic activity at low temperatures (<25 °C) and possess a more flexible three-dimensional structure compared to mesophilic or thermophilic lipases. This flexible structure enables the enzyme to maintain conformational stability at low temperatures through hydrogen bonding and hydration. The enzyme’s flexible loop region, rich in glycine and proline, lowers the activation energy of the reaction. The high catalytic activity of cryolipases and their strong affinity for substrates at low temperatures render them highly promising for industrial applications. Furthermore, the reduced energy consumption and broad adaptability of these enzymes enable them to function effectively under unfavorable conditions in industrial processes.

In alkaline industrial processes such as detergent production, leather degreasing, and fish oil hydrolysis, conventional chemical methods rely on high temperatures (60–80 °C) and strong alkali (pH 10–12) conditions to drive the reaction [8,9]. While these conditions improve efficiency, they also present significant drawbacks: the high-temperature environment promotes saponification of fats and oils, degradation of heat-sensitive products (e.g., ω-3 fatty acids), and increased energy consumption. Enzymatic catalysis is considered an ideal alternative due to its environmentally friendly and efficient nature. However, conventional lipases are unstable and prone to inactivation in high-temperature alkaline systems and may even exacerbate side reactions, such as substrate oxidation and saponification, under these conditions, thereby hindering their industrial application. While the flexible protein structure of low-temperature lipases reduces the activation energy of reactions [8] and enables efficient catalysis at room temperature—thereby avoiding the negative side effects of high temperatures—most existing low-temperature lipases suffer from insufficient alkaline resistance. For instance, the cryolipase reported by Su et al. [9] exhibited a more than 50% reduction in activity at pH 9.5, while the cold-tolerant lipase developed by Abdella et al. [10] retained only 10% of its activity at pH 10.0, severely limiting its application in alkaline industrial environments. Alkaline conditions (pH 9–11) significantly enhance enzymatic efficiency by activating the catalytic centers of lipase (e.g., deprotonation of serine residues) and facilitating the spontaneous emulsification of oleyl esters (e.g., fish oils). Additionally, these conditions inhibit oxidative side reactions and shift the reaction equilibrium toward hydrolysis. Therefore, developing a novel lipase that combines high activity at low temperatures with robust alkali tolerance (e.g., pH 11.0) is crucial. The development of such an enzyme will drive innovation in low-temperature detergent decontamination, green hydrolysis of fish oil, and leather cleaning and degreasing while providing technical support for sustainable development.

In this study, a lipase derived from Bacillus cereus was characterized, and an NCBI comparison revealed a low identity with known lipase genes, indicating that it is a relatively novel enzyme. The lipase was clonally expressed and purified using Escherichia coli. The purified enzyme exhibited a high activity at low temperatures, remained stable after 24 h of incubation under highly alkaline conditions, demonstrated tolerance to organic solvents, showed enhanced catalytic activity in the presence of surfactants, and exhibited regioselectivity, indicating its broad applicability in oil hydrolysis. These properties endow it with significant potential for alkaline industrial applications.

2. Materials and Methods

2.1. Bacterial Strains, Plasmids, and Chemicals

Bacillus cereus U2 was isolated from soil samples collected near the cafeteria of the Guangxi Academy of Sciences, Guangxi Zhuang Autonomous Region, China, and preserved at the Wuhan Conservation Center, China, under the strain conservation number CCTCC M20242887. Strain U2 was cultured on LB medium and agar plates at 37 °C. The genome extraction kit, restriction endonuclease, T4 DNA ligase, and Taq DNA polymerase were obtained from Takara (Kusatsu, Japan). p-Nitrophenyl (p-NP) esters were purchased from Sigma (St. Louis, MO, USA). All other chemicals used were of analytical grade and procured from Sinopharm Chemical Reagent Co. Ltd. (Shanghai, China).

2.2. Prediction of Genes

DNA and protein sequence comparisons were conducted using blastn and blastp tools, respectively, available at NCBI BLAST (http://www.ncbi.nlm.nih.gov/BLAST/) (accessed on 11 March 2025). Multiple sequence alignments were performed using Clustal W2 (http://www.ebi.ac.uk/Tools/msa/Clustalw2/) and visualized with ESPript 3.0 (https://espript.ibcp.fr/ESPript/ESPript/) (accessed on 12 March 2025). Phylogenetic analysis was performed using MEGA 11.0, employing the neighbor-joining method, with tree reliability estimated via bootstrap analysis with 1000 replicates [11]. Protein physicochemical parameters were analyzed using the ProtParam tool from the ExPASy Proteomics Server (https://web.expasy.org/protparam/) (accessed on 13 March 2025). Signal peptides were predicted using the SignalP 5.0 server (SignalP). The 3D structure of the target protein LipU was modeled based on homology to the structure of Pelosinus fermentans, as reported by SWISS-MODEL (http://swissmodel.expasy.org/) (accessed on 14 March 2025), using DSM 17108 lipase (PDB: 5AH0) as a template and visualized in PyMOL 3.1.

2.3. Gene Cloning, Expression, and Protein Purification

To identify the target lipase gene, a whole-genome sequence of Bacillus cereus (GenBank: WP_090976694.1) was obtained from NCBI, and a BLAST pairwise sequence analysis was performed. Based on the sequence of the target lipase, primers were designed to amplify the gene, and signal peptide prediction was conducted using SignalP 5.0. The analysis indicated the presence of a signal peptide at the 5′ end of the gene, which could potentially affect its expression in Escherichia coli. Therefore, the signal peptide was excised. The primers used for amplification were F1 (5′-GGGGTACCGCGAAAATATCATACGCTGAAG-3′) and R1 (5′-CCGCTCGAGTTTCGGGTAAACGTGATAGCTT-3′). The PCR products were then double-digested with Kpn I and Xho I restriction enzymes, and the isolated gene was ligated into the pET-30a(+) vector using T4 DNA ligase. The recombinant plasmid was transformed into E. coli DH5α, and positive clones were verified by colony PCR. The plasmid was subsequently sequenced. The resulting recombinant plasmid, pET30a-LipU, was then transferred into E. coli Rosetta. A 1% (v/v) inoculum of the recombinant E. coli was added to 100 mL of LB medium and incubated at 37 °C until the cells reached log phase. Gene expression was induced by adding 1 mM IPTG, followed by incubation at 16 °C and 120 rpm for 16 h.

Proteins were extracted through ultrasonic cell disruption and were purified using nickel affinity chromatography [12]. Briefly, cells were harvested by centrifugation at 5000 rpm and 10 °C, before being resuspended in 10 mL of lysis buffer containing 10 mM imidazole. Cell disruption was performed in an ice-water bath for approximately 20 min until the lysate became translucent. The lysate was then centrifuged at 12,000 rpm for 30 min to collect the supernatant as the crude enzyme solution. This solution was filtered through a 0.22 μm membrane and incubated with pre-washed Ni-NTA resin for 1 h on ice with gentle shaking. After incubation, the mixture was loaded onto an empty chromatography column, and unbound proteins were removed. The target protein was sequentially eluted with imidazole at concentrations of 10, 30, 60, 100, 200, 300, 400, and 500 mM. Eluted fractions were concentrated and desalted using a 10 kDa ultrafiltration tube and centrifugation at 5000 rpm and 4 °C, with repeated washes using deionized water. The final purified enzyme solution volume was concentrated to approximately 1–1.5 mL.

2.4. Enzyme Activity Determination

Solution A comprised 16.5 mmol/l p-nitrophenol palmitate (pNPC16) isopropanol solution. Solution B comprised 0.05 mol/L glycine–NaOH buffer solution (pH = 11.0) incorporating 1% (w/v) sodium deoxycholate and 0.1% (w/v) gum arabic.

The microplate reader is a high-throughput analytical instrument based on the principle of photoelectric colorimetry. Its core function is to measure the absorbance, emission, or scattering intensity of samples in microtiter plates at specific wavelengths using an integrated optical system. In enzyme assays, a wavelength of 410 nm is commonly used to assess the degree of substrate hydrolysis, thereby indicating enzymatic activity within the reaction system.

The reaction system was as follows: Briefly, 50 μL of solution A was mixed with 450 μL of solution B after preheating at 25 °C for 5 min. The purified enzyme was diluted and reacted for 10 min. The reaction was terminated by adding 500 μL of 10% (w/v) trichloroacetic acid (TCA), and then the pH was adjusted by the addition of 1 mL of 0.5 M Na₂CO₃ to neutralize the TCA and measured by using spectrophotometric methods at 410 nm. The amount of decomposed pNPC16 was measured at 410 nm [12]. The unit of lipase activity, U, was defined as the amount of lipase consumed to produce 1 μmol of nitrophenol per min of pNPC16 decomposition. Each set of experiments was repeated three times.

2.5. Effect of Temperature on Lipase Activity and Stability

The relative activity of lipase was determined at different temperatures (10–50 °C) to obtain the optimum reaction temperature. The diluted enzyme was tested for residual activity after keeping it at a temperature of 20–50 °C for 15–60 min. Thermal stability of lipase was analyzed by comparison with an untreated control enzyme.

2.6. Effect of pH on Lipase Activity and Stability

Buffers of different pH were prepared using acid–Na₂HPO₄ buffer (pH 6.0–7.5), Tris-HCl buffer (pH 7.5–9.5), and glycine–NaOH buffer (pH 9.5–12.0). After dissolving the substrate pNPC16 in buffer solutions of different pH (6.0–12.0), the relative lipase activity was measured, and the optimal pH for the reaction was determined [13]. After dilution of lipase using buffer solutions with pH ranging from 6.0 to 12.0, the lipase was kept at 4 °C and 30 °C for 24 h, and then the residual activity was measured. The pH stability of lipase was analyzed by comparing it with an untreated control enzyme without added buffer solution.

2.7. Effects of Organic Solvents on Lipase Activity

We used a pH 11.0 glycine–NaOH buffer solution to dilute the lipase before adding methanol, ethanol, n-propanol, n-butanol, isoamyl alcohol, ethyl acetate, acetone, dimethyl sulfoxide, acetonitrile, glycerin, and chloroform, respectively, so that the final concentration of the organic solvent was 15% and 30% (v/v). After incubating the mixture at 25 °C and pH 11.0 (glycine–NaOH buffer) for 1 h, the relative lipase activity was determined. The effect of organic solvent on lipase was calculated by comparing the enzymes with and without the addition of organic solvent.

2.8. Effects of Metal Ions and Surfactants on Lipase Activity

To investigate the influence of metal ions on lipase activity, metal salts containing K⁺, Li⁺, Na⁺, Co²⁺, Cu²⁺, Ca²⁺, Fe²⁺, Mg²⁺, Mn²⁺, Zn²⁺, Fe³⁺, and Al³⁺ were added to the standard enzymatic reaction at final concentrations of 1 mM, 2.5 mM, 5 mM, and 10 mM. Additionally, the effect of various chemical reagents on the activity of LipU was assessed by pre-incubating the lipase with surfactants at concentrations of 0.5% and 1% (v/v) for 30 min at 25 °C and pH 11 prior to the activity assay. An untreated enzyme sample, processed under identical conditions, served as the control. Residual lipase activity was then measured to evaluate the effects of metal ions and chemical additives.

2.9. Substrate-Specific Analysis of Lipases

To evaluate the substrate specificity of the lipase, p-nitrophenyl fatty acid esters with varying carbon chain lengths (C6, C8, C10, C12, C14, C16, and C18) were used as substrates at a final concentration of 16.5 mmol/L [12]. The relative activity of the lipase toward each substrate was assessed at pH 11.0 and 25 °C. Lipase activity was determined according to the previously described spectrophotometric method.

2.10. Selective Specificity Analysis of Lipases

The reaction system consisted of 2 mL of glycine–NaOH buffer (50 mM, pH 11.0), 1 g of fish oil, and 1 mL of crude LipU enzyme solution. After purging with nitrogen for 1 min, the reaction vessel was sealed and incubated at 20 °C, with shaking taking place afterwards at 200 rpm for 12 h. The reaction was terminated by placing the mixture in a boiling water bath for 10 min. To prepare the samples, the reaction mixture was dissolved in 50 mL of hexane, followed by the addition of 20 mL of 0.5 M ethanolic KOH solution and vigorous mixing. The upper organic phase, containing glycerides, was washed three times with 20 mL of purified water, and the solvent was removed by rotary evaporation at 55 °C and 50 rpm for 20 min. The products were analyzed using thin-layer chromatography (TLC) on 0.2 mm silica gel plates (10 cm × 20 cm). Samples dissolved in hexane were applied to the TLC plates, which were developed in a solvent system of hexane/ether/acetic acid (v/v/v, 85:15:1), dried with heat, and compared with standard compounds from Sigma (St. Louis, MO, USA).

2.11. Nucleotide Sequence Entry Number

The LipU nucleotide sequence was submitted to the GenBank database with entry number PV094892.

3. Results and Discussion

3.1. Gene Cloning, Sequence Analysis, and Molecular Modeling

A bioinformatics analysis of the LipU gene was initially conducted. The LipU gene was found to be 1178 base pairs long, encoding a protein of 392 amino acids, with a predicted molecular weight of 43.65 kDa and an isoelectric point of 6.60. BLASTP analysis revealed that LipU shared the highest identity (53.32%) with the Pelosinus fermentans DSM 17108 lipase (AIX10936.1), followed by the Geobacillus thermoleovorans lipase (AAD30278.1, 46.34%), Staphylococcus aureus lipase (AAA26633.1, 39.90%), and Staphylococcus hyicus lipase (CAA26602.1, 37.22%). Therefore, LipU represents a novel lipase that may possess unique properties.

Regarding the classification of lipase families, according to the system proposed by Arpigny and Jaeger [14] in 1999, Family I represents the typical true lipases, primarily derived from the genera Pseudomonas and Bacillus. Lipases in this family exhibit considerable sequence diversity. Family II lipases carry the characteristic GDSL domain, and some utilize a Ser-His catalytic structure, differing from the classical Ser-Asp-His catalytic triad. Family III lipases strictly adhere to the Ser-Asp-His catalytic structure and possess the standard alpha/beta hydrolase folding configuration. Family IV lipases are closely related to mammalian lipases and possess the standard alpha/beta hydrolase folding configuration. Family IV lipases share more than 60% sequence homology with mammalian lipases, and most are low-temperature-active enzymes. Family V lipases are primarily isolated from thermophilic microorganisms (e.g., Thermomyces lanuginosus) and exhibit excellent thermal stability. Family VI lipases typically have molecular weights lower than 30 kDa and exhibit specificity for short-chain fatty acid esters. Family VII includes eukaryotic homologous enzymes. Family I is further subdivided into six subfamilies.

As shown in Figure 1, phylogenetic tree analysis confirmed that LipU is classified as belonging to bacterial lipase Subfamily I.5 of Family I, indicating that LipU is a typical lipase. The uniqueness of Subfamily I.5 is distinguished by its surface cap structure, which is more hydrophobic compared to other subfamilies. Figure 2 shows that the sequence comparison reveals the catalytic triad of LipU, consisting of Ser121, His363, and Asp323. The residues from 183 to 264 and 288 to 322 in the α/β-hydrolase core of LipU form the cap structure domain, which covers the active site and catalytic triad. Figure 3 illustrates the 3D homology model of LipU, showing that Zn²⁺ is tetrahedrally coordinated by the side chains of Asp71, His91, His97, and Asp248.

3.2. Expression and Purification of LipU

The expressed proteins were analyzed using sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) (Figure 3B) [15]. Following nickel column purification and ultrafiltration, recombinant LipU was further analyzed using SDS-PAGE. As shown in Figure 3B, protein bands with molecular weights close to the theoretical value of LipU (calculated MW = 43.92 kDa) were observed, confirming the successful expression of LipU. A high yield is generally favorable for large-scale protein preparation. Therefore, the purification fold of LipU was calculated. A total of 25.6 mg of crude enzyme protein was obtained from 100 mL of E. coli culture medium, and 3.68 mg of pure enzyme protein was obtained after purification. As shown in

Table 1. Purification of recombinant LipU.

Steps	Total Protein (mg)	Total Activity (U)	Specific Activity (U/mg)	Purification (Fold)	Recovery (%)
Supernatant	25.6	2576	100.625	1	100
Purified LipU	3.68	1334	362.5	3.602	51.78

, the purification fold was 3.602, and the recovery was 51.78%.

3.3. Effect of pH and Temperature on LipU Activity

The catalytic temperature of lipases limits their industrial applications [8]. Therefore, the optimum temperature and thermal stability of recombinant LipU were determined. The results indicated that lipase LipU from Bacillus cereus had a low optimal temperature and exhibited good activity in the range of 10–35 °C. As shown in Figure 4A, LipU exhibited the highest activity at 20 °C and retained more than 80% of its initial activity after 24 h of incubation at 10–35 °C. However, after 24 h of incubation at 40 °C, the residual activity significantly decreased. These results indicate that recombinant LipU is a low-temperature lipase. The optimal temperature of lipases impacts their industrial applications, and the optimal temperature varies among lipases from different bacterial sources. Pria Laot Sabang strain 80, isolated from the underwater hot spring area of Pria Laot Sabang, Indonesia, produced the PLSA lipase with an optimum temperature of 70 °C [16]. The LipBST lipase from Bacillus stratophilus L1 has an optimum temperature of 35 °C [5]. AMS8, a low-temperature lipase from Pseudomonas sp., has an optimum temperature of 30 °C [17]. Compared to the lipases mentioned above, LipU has a lower optimum temperature. Compared to enzymes with higher optimal temperatures, low-temperature lipases have fewer α-helices, more β-folds, and a higher proportion of hydrophilic residues on their surfaces. This structure reduces intramolecular rigid interactions, such as hydrophobic stacking and salt bridges [8]. This structural flexibility enables the lipase to adapt its conformation and bind substrates effectively, even at low temperatures. Temperature stability measurements of LipU were performed, as shown in Figure 4B. LipU maintained high stability after 1 h of incubation at 30 °C and 35 °C. However, after 1 h of incubation at 45 °C, the residual activity decreased dramatically, retaining only 19.82% of its initial activity, indicating that the recombinant lipase has good low-temperature stability.

The pH profile of LipU was investigated. As shown in Figure 4C, LipU activity increased and then decreased within the pH range of 6.0–12.0, with optimal activity observed at pH 11.0. In terms of pH stability, recombinant LipU remained stable in the pH range of 7.0 to 11.0. LipU retained more than 95% of its enzyme activity after 24 h of incubation at 4 °C, pH 11.0, and more than 75% of its residual activity after 24 h of incubation at 30 °C, pH 7.0–11.0. In comparison to other cryolipases successfully expressed in heterologous hosts, LipU exhibits a higher optimal pH. As shown in

Table 2. Characteristics of low-temperature lipases previously expressed in heterologous hosts.

Source Strains	Optimum Temperature (℃)	Optimum pH	References
Bacillus cereus	20	11.0	This study
Acinetobacter sp. XMZ-26	15	10.0	[20]
Psychrobacter sp. G	35	8.0	[21]
Bacillus	35	8.0	[22]
Stenotrophomonas maltophilia GS11	35	8.0	[23]
Sorangium cellulosum Strain So0157-2	30	8.0	[24]
Psychrobacter sp. C18	30	8.0	[25]
Staphylococcus epidermidis AT2	25	8.0	[26]
psychrotrophic Yersinia enterocolitica	25	7.0	[27]
Rhizomucor endophyticus	40	6.0	[28]
Malassezia globosa	15	6.0	[29]
Candida albicans	15	5.0	[30]

, the optimal pH of LipU is higher than that of most reported cryolipases. This ability to function in high-pH environments provides a foundation for its participation in chemical reactions under strongly alkaline conditions. The stability of LipU, which is an alkaline lipase, suggests its potential for use in various industrial applications, including biodiesel synthesis [16], polyunsaturated fatty acid enrichment [17], and leather manufacturing [18,19].

3.4. Substrate Specificity of LipU

Defining the substrate selectivity of lipases is essential for their industrial application. In this study, the hydrolytic activity of p-nitrophenyl esters, ranging from C6 to C18, was assessed to elucidate their catalytic properties. This study found that purified LipU was able to hydrolyze p-nitrophenyl esters with fatty acyl chains ranging from C6 to C18 (Figure 4E), exhibiting maximum lipase activity (100%) against p-NP C10. The relative activities of LipU towards other substrates, compared to p-NP C10, were as follows: p-NP C6, 36.66%; p-NP C8, 35.67%; p-NP C12, 96.38%; p-NP C14, 93.10%; p-NP C16, 90.43%; and p-NP C18, 52.72%. Therefore, LipU exhibited higher activity towards medium- and long-chain substrates (C12, C14, and C16). The analysis of LipU’s hydrolytic activity towards C6–C18 p-nitrophenyl esters revealed efficient catalytic ability (>90% relative activity) for medium- and long-chain substrates (C12–C16), while activity significantly decreased for C18 (52.72%). A similar substrate selectivity has been reported for the lipase from Aspergillus niger GZUF36 [31]. However, the present findings contrast with those of the lipase from Bacillus stratosphericus L1 [5]. Typically, lipases exhibit the highest activity towards water-insoluble long-chain triglycerides, whereas esterases tend to catalyze esters with short-chain fatty acids. Therefore, the results of the substrate-specific characterization indicate that LipU is a true lipase. This substrate preference may be attributed to the structural properties of the enzyme’s active pocket, where the moderate hydrophobic lumen volume allows for flexible binding of medium- and long-chain acyls, while the C18 substrate may cause reduced catalytic efficiency due to spatial site-blocking or conformational rigidity [32]. This property makes LipU suitable for industrial applications in the degradation of medium- and long-chain fats and oils [33], such as food processing waste streams or biodiesel feedstocks.

3.5. Effect of Metal Ions on the Activity of Recombinant LipU

In industrial applications, reaction mixtures often contain metal ions that can significantly affect lipase activity. To investigate this, 12 metal ion solutions were tested for their effects on LipU activity. As shown in Figure 5, LipU activity was enhanced in the presence of Cu²⁺, Co²⁺, Na⁺, Ca²⁺, Mg²⁺, and Li⁺. Specifically, Ca²⁺ and Mg²⁺ increased enzyme activity by 172% and 126%, respectively, aligning with previous studies where Ca²⁺ and Mg²⁺ were found to be major contributors to the activity of lipase from Pseudomonas fluorescens [34]. This activation is likely due to conformational changes in the lipase upon metal ion binding, which form salt bridges. These interactions facilitate the release of fatty acids during hydrolysis, creating insoluble salts that enhance both enzyme stability and catalytic activity [35,36]. Interestingly, Cu²⁺ did not inhibit LipU activity, as typically observed with conventional lipases; instead, it promoted enzyme activity. Similar findings have been reported for lipases from Candida rugosa [37], where Cu²⁺ also served as an activator. The increased negative surface charge of LipU under alkaline conditions may reduce the binding of Cu²⁺ to key carboxylic acid residues due to electrostatic repulsion.

Furthermore, LipU retained 78.76–90.18% of its initial activity after incubation with 1 mM, 2.5 mM, and 5 mM of Al³⁺, Mn²⁺, Zn²⁺, Fe²⁺, and Fe³⁺. Low concentrations of Fe²⁺ and Fe³⁺ did not significantly inhibit enzyme activity, a finding consistent with studies on lipase from Bacillus subtilis EH 37 and Bacillus cereus C7. This lack of inhibition is likely due to the formation of metal ion complexes with OH⁻ in alkaline environments, reducing the concentration of free metal ions and weakening their direct interaction with the enzyme’s active site. However, lipase activity was significantly inhibited at a 10 mM concentration of these metal ions. A similar inhibitory effect was observed in lipase from Pseudomonas fluorescens SBW25 at high metal ion concentrations [34]. The negative impact of metal ions on lipase activity typically results from the direct inhibition of the catalytic site, as is the case with many other enzymes [38]. Additionally, the 3D model in Figure 3A shows that LipU is ligated with Zn²⁺. However, in the experimental measurements, enzyme activity decreased as the Zn²⁺ concentration increased. This may be due to the fact that Zn²⁺ ligation could be structural rather than catalytic, and Zn²⁺ may interfere with the enzyme’s water binding, inducing the partial denaturation of the protein, consequently reducing its activity [39].

3.6. Effects of Surfactants and Organic Solvents on LipU

Surfactants influenced LipU activity at concentrations of 0.5% and 1.0%. As shown in Figure 6A, the presence of 0.5% surfactant significantly enhanced LipU enzyme activity, with Tween20 and Tween80 increasing the enzyme activity to 159.73% and 185.82%, respectively. When the surfactant concentration was increased to 1.0%, the enzyme activity of Tween20 increased to 167.79%, whereas that of Tween80 decreased to 104.20%. The enzyme activity induced by X-100 transitioned from an enhancement to a significant inhibition. The enhancement of enzyme activity by Tween20 and Tween80 has been previously reported in the literature. In lipid hydrolysis applications, Tween20 enhanced the enzymatic activity of lipase from Rhizopus homothallicus [40] by 25%, and Tween80 increased the activity of lipase from Kid pregastric [41] lipase by 35%. In comparison, Tween20 and Tween80 showed greater enhancement of LipU activity. This enhancement is primarily attributed to the surfactants’ interaction with the enzyme via hydrogen bonding and hydrophobic interactions, which stabilize the enzyme [42]. Furthermore, the surfactants preferentially bind to specific enzyme binding sites, forming stronger hydrophobic interactions that induce a conformational change in the enzyme, opening its active site [43]. In contrast, the addition of SDS and sodium deoxycholate resulted in the significant inhibition of LipU activity at both concentrations, with little or no enzyme activity observed. The negative effect of SDS and sodium deoxycholate on LipU is likely due to their interference with substrate access to the enzyme’s active center [26,44]. Additionally, the inhibition of enzyme activity at high concentrations of X-100 is consistent with previous findings, as Bacillus species show weak tolerance to X-100 [45,46]. At low concentrations, activation is weak, while at higher concentrations, enzyme activity is inhibited. This reduction in activity may result from surfactant saturation near or above the critical micelle concentration (CMC), which limits lipase binding at the oil/water interface, leading to enzyme inactivation [47]. These findings suggest that LipU may have potential applications in the detergent industry and leather manufacturing.

We also investigated the effect of organic solvents in aqueous mixtures on the activity of LipU. As shown in Figure 6B, after treatment with ethanol, n-butanol, isoamyl alcohol, dimethyl sulfoxide, and glycerol at a concentration of 15%, LipU retained more than 78.6% of its residual activity. However, after treatment with ethyl acetate, acetone, and chloroform at the same concentration, LipU activity was drastically inhibited, remaining below 39.60%. When the solvent concentration was increased to 30%, lipase activity was further inhibited in all experimental groups except glycerol, with residual activity ranging from 7.30% to 70.54% of its original activity. The significant decrease in lipase activity in the presence of most organic solvents is consistent with previous findings. A similar phenomenon was observed for lipase from Staphylococcus aureus [48]. This reduction in activity may be attributed to the decreased water activity around the protein, which promotes structural denaturation [49,50]. LipU demonstrated moderate tolerance to some organic solvents, such as dimethyl sulfoxide and glycerol. Treatment with glycerol at concentrations of 15% and 30% increased LipU enzyme activity to 119.87% and 106.56%, respectively. The stabilization of protein molecules by glycerol can occur through two mechanisms: firstly, glycerol changes the properties of the protein environment, such as viscosity and dielectric constant. The presence of glycerol increases the viscosity of the solution, which reduces the mobility of the protein molecule, thereby enhancing its stability. Secondly, glycerol influences the conformation of the protein and promotes interactions that favor the stability of the protein structure.

LipU enzyme activity was slightly reduced to 91.50% after treatment with 15% dimethyl sulfoxide. However, when the concentration of dimethyl sulfoxide was increased to 30%, the residual activity decreased less, with 70.54% of its original activity remaining. Lipases tolerant to polar organic solvents have been reported in the literature. For instance, the lipase from Ureibacillus thermosphaericus [51] showed 62% residual enzyme activity after one hour of incubation in 30% dimethyl sulfoxide. Organic solvents in the reaction medium typically enhance the solubility of reactants, reduce reaction viscosity, and minimize the inhibitory effect of chemicals on the enzyme, thereby improving conversion efficiency. In summary, LipU exhibits moderate tolerance to organic solvents. The stability and activity of LipU can be attributed to its flexible, open structure, which allows organic solvents to interact with hydrophobic amino acid residues in the lid region, thereby facilitating substrate access to the active site. Furthermore, hydrophobic solvents may help maintain the enzyme’s natural conformation and reduce side reactions by limiting the redistribution of water molecules in the enzyme’s hydration layer. In industrial applications, the high stability and activity of bacterial lipases in organic solvents is a significant advantage. Therefore, the stability of LipU in solvents such as isoamyl alcohol, n-butanol, ethanol, and dimethyl sulfoxide makes it suitable for biocatalysis applications, including the optical splitting of chiral compounds, organic synthesis, and other uses.

3.7. Regional Selectivity of LipU

The regioselectivity of lipase is a crucial property of its catalytic function, directly influencing its applicability in industries such as food production and pharmaceuticals. Lipases with 1,3-selective specificity have active sites that are better suited to ester bonds at the sn-1 and sn-3 positions of triglycerides, facilitating hydrolysis at these positions more efficiently. Conversely, some lipases exhibit a looser active site structure that can accommodate ester bonds at multiple positions of the triglyceride, thereby catalyzing hydrolysis at both the 1,2 and 1,3 positions. Lipases derived from Aspergillus carneus [52] and Aspergillus niger AN0512 [53] are selective for the sn-1,3 positions, while lipases from Pseudozyma tsukubaensis [54] and Acinetobacter calcoaceticus LP009 [55] are non-regioselective and catalyze hydrolysis at both the 1,2 and 1,3 positions. In this study, the purified lipase was capable of hydrolyzing glycerol trioleate into glycerol monooleate, 1,2(2,3)-dioleate, 1,3-dioleate, oleic acid, and glycerol, demonstrating non-regioselectivity, as shown in Figure 7. The non-regioselectivity of this lipase suggests that its active site structure may be more flexible, allowing it to accommodate multiple ester bonds on triglycerides. This characteristic provides potential value for its application in industrial catalysis and serves as a foundation for further studies on the enzyme’s catalytic mechanism and the structure–function relationship of lipases.

4. Conclusions

LipU belongs to the lipolysis enzyme Family I.5, with the highest identity (53.32%), which means it is a relatively novel lipase. In this study, the optimal temperature of LipU was found to be 20 °C and the optimal pH was 11.0. The results show that this enzyme has a high catalytic activity at low temperatures and that it maintains a high activity in high-pH environments. In addition, LipU is, to a certain extent, resistant to organic solvents while also being resistant to metal ions and being equipped with the potential to be partially enhanced by surfactants. Its non-regional selectivity could also be widely useful in various industries. These properties make LipU an attractive potential candidate for alkaline industrial applications.