Insight into improved specificity and thermostability of Geobacillus zalihae T1 lipase by introducing novel molecular interactions via phenylalanin to cysteine substitution.

Insight into improved specificity and thermostability of Geobacillus zalihae T1 lipase by introducing novel molecular interactions via phenylalanin to cysteine substitution - PMC Skip to main content An official website of the United States government Here's how you know Here's how you know Search Log in Primary site navigation Search Log in Search PMC Full-Text Archive Search in PMC Journal List User Guide As a library, NLM provides access to scientific literature. Inclusion in an NLM database does not imply endorsement of, or agreement with, the contents by NLM or the National Institutes of Health. Learn more: PMC Disclaimer | PMC Copyright Notice 3 Biotech . 2025 May 14;15(6):166. doi: 10.1007/s13205-025-04330-5 Insight into improved specificity and thermostability of Geobacillus zalihae T1 lipase by introducing novel molecular interactions via phenylalanin to cysteine substitution Samah Hashim Albayati 1, 2 , Muna Hilal Shabeeb 6 , Oluwasola Michael Akinola 1, 5 , Nima Ghahremani Nezhad 1, 2 , Adam Leow Thean Chor 1, 4, 5 , Mohd Shukuri bin Mohamad Ali 1, 3, 4 , Fairolniza binti Mohd Shariff 1, 2 , Noor Dina binti Muhd Noor 1, 3 , Raja Noor Zaliha Raja Abd Rahman 1, 2, 4, ✉ Author information Article notes Copyright and License information 1 Enzyme and Microbial Technology Research Center, Faculty of Biotechnology and Bimolecular Science, Universiti Putra Malaysia, Serdang, 43400 Selangor, Malaysia 2 Department of Microbiology, Faculty of Biotechnology and Bimolecular Science, Universiti Putra Malaysia, Serdang, 43400 Selangor, Malaysia 3 Department of Biochemistry, Faculty of Biotechnology and Bimolecular Science, Universiti Putra Malaysia, Serdang, 43400 Selangor, Malaysia 4 Institute Bioscience, Universiti Putra Malaysia, Serdang, 43400 Selangor, Malaysia 5 Department of Cell and Molecular Biology, Faculty of Biotechnology and Bimolecular Science, Universiti Putra Malaysia, Serdang, 43400 Selangor, Malaysia 6 Department of Salahuddin Education, Ministry of Education, Baghdad, Iraq ✉ Corresponding author. Received 2024 Dec 13; Accepted 2025 Apr 22; Issue date 2025 Jun. © King Abdulaziz City for Science and Technology 2025. Springer Nature or its licensor (e.g. a society or other partner) holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law. PMC Copyright notice PMCID: PMC12078924  PMID: 40386625 Abstract Lipase specificity is a crucial requirement for industrial employment; however, tuning the lipase specificity in some cases can impair the structure and affect its stability. To enhance the T1 lipase specificity, we targeted the conserved bulky residue Phe180 in the lid domain to eliminate the impact of steric hindrance, as it constrains substrate accession to the enzyme active site and affects specificity. This residue was pre-substituted with a small side chain residue by utilizing DynaMut2 software to ensure that this substitution did not affect lipase stability. Phe180Cys was chosen because it exhibited a stabilizing effect by showing (ΔΔ G Stability ) of 0.74 kcal/mol, which was subsequently substituted by the rational design approach. This variant has successfully exhibited specificity modification toward long fatty acid chains as a result of increasing the distance between the lid domain and catalytic site by 1.2 Å and the volume of active site by 190.2 Å 3 . In addition, this F180C variant exhibited an increase in the optimum temperature and thermal denaturation point to 75 °C and 78 °C, respectively, with an improvement in the lipase stability in the organic solvents. The analysis of the atomic interactions revealed a change in the whole H-bonds, S / π interactions, and salt bridge network. The biophysical study revealed changes in the secondary structure content compared with wt-T1. The MD simulation results displayed lower RMSD, gyration radius, and SASA values for the mutated lipase structure. In conclusion, comparative analysis of the atomic interactions resulting from structural modification can significantly elucidate the specificity and thermostability of enzymes of industrial relevance. Supplementary Information The online version contains supplementary material available at 10.1007/s13205-025-04330-5. Keywords: H-bonds, Salt bridges, S / π interactions, Lipase thermostability, Specificity, Binding site volume Introduction Lipases (triacylglycerol hydrolases, EC 3.1.1.3) are a group of enzymes widely used for hydrolyzing triacylglycerols to produce various products such as free fatty acids, monoacylglycerol, and diacylglycerol. In addition, they play crucial roles in lipid synthesis via esterification, transesterification, and alcoholysis in organic solvent environments (Chandra et al. 2020 ). In general, most naturally occurring fats and oils contain various fatty acids with differing chain lengths, which consequently decrease the concentration of specific FAs. Therefore, it is necessary to enhance the desired FA content in the acylglycerol form (Qin et al. 2021 ). Lipases are widely utilized in various industrial applications, including pharmaceuticals, food, papermaking, detergents, textiles, cosmetic additives, bioremediation, waste oil disposal, and biodiesel production (Kumar et al. 2023a ). However, high temperature is the main factor for successful industrial bioprocessing. Furthermore, executing bioprocesses at elevated temperatures can eliminate the potential contamination caused by mesophilic microbes, minimize the need for external cooling, and improve substrate solubility and reaction rates. Thus, lipases with high thermostable activities at higher optimum temperatures than their counterparts are highly desirable (Liu et al. 2017a ). An important structural feature of many lipases is the mobile subdomain lid or flap located over the active site. The structural domain is crucial for lipase activity because it protects the active site from the environment and is responsible for substrate accessibility when in close conformation. However, only the open-label lipases exhibited catalytic activity (Khan et al. 2017b ). Many studies have reported the role of the lid domain in modifying lipase activity and thermostability (Cui et al. 2022 ; Timucin and Sezerman 2013 ; Yu et al. 2012 ). Various mutagenesis approaches have been applied to enhance the lipase specificity and thermodynamic stability (Jiang et al. 2020 ; Xiang et al. 2023 ; Yoshida et al. 2021 ; Qin et al. 2021 ). However, the most commonly used strategies are directed evolution and rational, and semi-rational designfor enhancing the targeted enzymes characteristics (Guan et al. 2020 ; Cheng and Nian 2023 ; Pongsupasa et al. 2022 ). Recently, the rational design engineering approach has been utilized alongside computational software tools that can enhance the modulation of the biochemical properties of enzymes (Marques et al. 2021 ; Yauheniya and Manish 2019 ). Furthermore, the advantage of employing these approaches is the considerable library size reduction; thus, less effort and time are required (Ding et al. 2024 ). However, novel deep learning software tools are considered fast and accurate siamese networks by predicting the changes in the Gibbs Free Energy upon single and multiple point mutations (Zhou et al. 2023 ). An important factor in protein stability is atomic interactions. These interactions include salt bridges, hydrogen bonds, and aromatic interactions. The salt bridges are electrostatic interactions that arise between opposite charges of amino acid residues. Enhancing the abundance of amino acids with opposite charges can increase the salt-bridge interactions in or between helical domains, which is crucial for maintaining the thermostability of the protein (Yang et al. 2015 ). In addition, maintaining the rigidity of the protein backbones through hydrogen bonds is another factor for enhancing protein thermostability. Another type of atomic interaction is mainly aromatic interactions, which include the cation–aromatic pair (cation-p) interactions and aromatic–aromatic (pi–pi) stacking. Cation-p interactions are stronger than pi-pi stacking interactions. However, the energy generated by the cation-p interaction is greater than that generated by the ionic bonds by approximately two-fold (Yang et al. 2015 ). T1 lipase is an intracellular lipase isolated from the strain Geobacillus zalihae , derived from a palm oil mill in Malaysia (Rahman et al. 2007 ). Recombinant T1 lipase is a thermophilic alkalophilic that exhibits stability at an optimum temperature is 70 ℃ and an optimum pH of 9 (Rahman et al. 2010 ). T1 lipase exhibits several substrate specificities but with variable fat and oil preferences (Leow et al. 2007 ). Indeed, it showed a high preference toward the C12 chain of fatty acids and exhibited 1,3-regio-selectivity (Ishak et al. 2021 ; Qin et al. 2021 ). A previous study showed that T1 lipase altered the T1 lipase regiospeccifity by targeting the bulky lid and oxyanion hole residues (Albayati et al. 2023 ). To explore the role of bulky lid domain residues in altering lipase specificity, a recent study was conducted to explore the role of bulky residues in the lid domain of T1 lipase on specificity and thermostability. The prediction of convenient small-residue amino acid substitutions that would enhance the lipase specificity simultaneously not affecting the stability was conducted by utilizing deep learning software. Site-directed mutagenesis was performed along with biochemical, biophysical, and in silico characterization. Results and discussion Screening for substitution Phe180 amino acid residue of wt-T1 lipase According to a previous study conducted on T1 lipase, which implied changing T1 lipase regiospecificity and fatty acid selectivity by targeting two of the lid domain (F180/F181) and one of the oxyanion hole (F16) bulky conserved residues (Albayati et al. 2023 ). This study targeted the lid domain residue Phe180, which may hinder the entrance of the bulky substrate. Therefore, this residue was substituted with a small amino acid to reduce steric hindrance; however, the substituted residue should not affect T1 lipase stability. To detect convenient amino acid residues with small side chains, screening was performed using Dynamut2. This screening showed that Cys was the best option because it exhibited a stabilizing effect by showing the (ΔΔ G Stability ) 0.74 kcal/mol, which may result in an increase in the stability change upon substitution. Mutated lipase expression and purification The lid domain residue Phe 180 was targeted by rational design to construct a Cys variant (F180 C) using a QuikChange Lightning Mutagenesis kit (Stratagene, Agilent Technologies, Santa Clara, CA, USA) and a specific mutagenesis codon primer at the targeted site. The alignment of the mutated lipase sequence indicated the existence of nucleotide changes at the targeted residue positions ( Fig. S1 ) . Affinity chromatography (Ni-Sepharose High Performance, GE Healthcare, Chicago, IL, USA) was utilized for the purification of mutagenized lipase in this study and wild-type lipase, in which the recovery rates of purification for wt-T1 and its variant F180 C were 54.8% and 55.1%, respectively, for respective protein yields of 305.2 and 372.4 mg/L (Table S2 ). The targeted lipases were successfully purified to homogeneity, and SDS–polyacrylamide gel electrophoresis was utilized to determine the existence of a single band at approximately 44 kDa (Fig. S2 ). The residue was targeted because many studies have reported the importance of flap domain residues on lipase thermostability (Mohammadi et al. 2016 ; Veno et al. 2019 ; Cui et al. 2022 ; Yu et al. 2012 ). In addition, the enhancement of enzyme specificity by substituting the lid domain has been reported in many studies (Akoh et al. 2004 ; Khan et al. 2017a ; Mohamed et al. 2018 ). In a study, the activity enhancement was up to 2.6-fold of that of Bacillus thermocatenulatus lipase when F181 was exchanged with alanine. This activity improvement was attributed to the reduced lid domain steric hindrance (Karkhane et al. 2009 ). In addition, targeting the lid domain of Pseudomonas sp. CR611 Lip I.3 lipase by exchanging serine 154 with threonine and glycine 152 with leucine introduced a five-fold increase in activity and a two-fold enhancement (Panizza et al. 2015 ). Moreover, a study revealed that targeting the lid domain hydrophobicity of P. expansum lipase by substituting Y225 F/S232 A residues increased the enzyme activity by approximately 35% and affected interfacial activation (Tang et al. 2015 ). Determination of the optimal temperature and pH of the wt-T1 and T1/F180 C variant lipases The mutated T1/F180 C lipase exhibited optimal activity at a temperature of 75 °C compared with the wt-T1 lipase, which exhibited optimal activity at 70 °C (Fig. 1 ). The Kwon and Rhee colorimetric method was utilized to investigate the optimal pH of the wt-T1 and T1/F180 C variants using various buffers ranging from pH 4 to 12 (Kwon and Rhee 1986 ). The lipase variant was pre-incubated at pH 4 to 12 for 30 min and 70 °C. The high activity displayed by the F180 C variant was between pH values 6.0–10.0, and at 70 ℃. The optimum activity of the T1/F180 C variant was determined at the optimum wt-T1 glycine NaOH buffer at pH 9 and at 70 ℃ ( Fig. 2 ) . A study reported enhanced Staphylococcus epidermidis AT2 (rT-M386) lipase thermostability from 25 to 45 °C by substituting the lid domain residue Gly 210 with Cys. The stability improvement was attributed to the construction of stronger noncovalent interactions by the Cys residue in the protein structure (Veno et al. 2019 ). Furthermore, engineering disulfide bonds is another reason for improving thermostability and activity, which is offered by substituting a targeted residue with Cys (Wiedemann et al. 2020 ). The l ‐Isoleucine hydroxylase enzyme was also subjected to a site-directed mutagenesis approach and substituted T181 with Cys, and this substitution resulted in enhancing the enzyme thermostability to 10.27‐fold of half‐life and 3.56‐fold of the specific enzyme activity (Qiao et al. 2020 ). Regarding the effect of the mutation on pH, substituting four lid residues of T1 lipase did not shift the optimum pH value (Tang et al. 2017 ). In addition, the enzymatic conformational stability is affected by the surface-charged residues, which is demonstrated via electrostatic interactions and hydrogen bonds with water molecules in the surrounding environment (Pedersen et al. 2019 ; Wu et al. 2020 ). Fig. 1. Open in a new tab Optimal temperatures for T1/F180 C and wt-T1 lipases. Lipase activity was identified utilizing the colorimetric Kwon and Rhee method, and the optimal activity was recorded as 100% for each enzyme in the temperature range from 40 to 90 °C. The standard protein concentration was 1 mg/ml for each sample. The samples were analyzed in triplicate for this assay. SD is indicated as error bars. When the error bar cannot be seen, the deviation is less than the size of the symbol Fig. 2. Open in a new tab Optimal pH values of T1/F180 C and wt-T1 lipases. a wt-T1 lipase. b T1/F180 C. Lipase activity was identified using the Kwon and Rhee method, and the optimal activity was recorded as 100% for each enzyme. The standard protein concentration was 1 mg/ml for each sample. All samples were analyzed in triplicate. SD is indicated as error bars. When the error bar cannot be seen, the deviation is less than the size of the symbol The impacts of metal ions on the activity of the wt-T1 and T1/F180 C variant lipase The impacts of the metal ions on the structural modifications and catalysis are known to be large. Therefore, to analyze the effects of the altered structure caused by the mutation on the lipase function, the metal ions were utilized to measure the lipase activity. The analyzed data showed various catalytic behaviors in the tested variant, T1/F180 C, compared to wt-T1 lipase after treatment with metal ions such as Ca 2+ , Na +2 , Mg 2+ , Fe 3+ , Cu 2+ , Ni 2+ , and Zn 2+ . The results demonstrated that calcium ions efficiently stimulated the activity of the mutagenized T1/F180 C lipase compared with the wt-T1 lipase (Fig. 3 ). The increase in T1/F180 C activity reached 137% and 95% compared with 115% and 88% for wt-T1 lipase after exposure to 1 and 5 mM calcium ions, respectively. The Na +2 effect was observed to have a slight effect on the activity of the T1/F180 C variant compared with that of wt-T1 lipase. Other metal ions, such as Fe 3+ ,Cu 2+ , Ni 2+ , and Zn 2+ at concentrations of 1 mM and 5 mM were observed to have a negative impact on the activity of T1/F180 C more than the wild type. Mg 2+ affected the activity in T1/F180 C as it displayed approximately 48% and 46% compared to 65% and 77% of wt-T1 lipase after treatment with 1 mM and 5 mM, respectively. Similarly, for the Fe 3+ ion, it showed varied activity ratio of T1/F180 C was 30% compared to 45% for wt-T1 at 1 mM and 20% of T1/F180 C compared to 12% of wt-T1 lipase activity at 5 mM. Ni 2+ affected the activity of T1/F180 C by 45% compared to 89% for wt-T1 lipase at 1 mM and 76% compared to 89% at 5 mM compared to wt-T1 lipase. The remaining Cu 2+ and Zn 2+ ions showed similar effects on the activity of the negative impacts of T1/F180 C compared with wt-T1 lipase. To maintain the structural stability of enzymes under extreme conditions, metal ions are essential (Kim et al. 2000 ; Hertadi and Widhyastuti 2015 ; Tayyab et al. 2011 ). For some lipases, such as Geobacillus sp. and Staphylococcus sp, calcium ions are required for maintaining their stability and activities (Simons et al. 1999 ; Tayyab et al. 2011 ). Calcium is considered a crucial ion for maintaining the enzyme active conformation because it protects the enzymes against thermal denaturation (Yamada et al. 2016 ; Johri et al. 2012 ). Similarly, sodium ions have also been reported to have a positive effect on the lipase stability and activity (Quilles Junior et al. 2016 ; Alabdalall et al. 2021 ). However, the incubation of the enzyme with Ni 2+ , Fe 3+ , Cu 2+ , and Zn 2+ ions inhibited the enzyme activity. A similar finding reported previously indicated that Ni 2+ , Fe 3+ , and Cu 2+ ions potentially inhibit the activity of T1 lipase in Geobacillus zalihae (Ishak et al. 2020 ). In addition, another study was conducted on Bacillus thermoamylovorans (Lip501r), in which Zn 2+ inhibited the lipase activity. In the same study, lipase activity was stimulated in the presence of calcium ions. Meanwhile, the addition of Fe 3+ reduced the enzyme activity in most variants, except in T118 N, which increased its activity in the presence of a small amount of Fe 3+ and decreased its activity with increasing the concentration of the same ion (Yamada et al. 2016 ). The main mechanism underlying the modulation of enzyme activity via induction and reduction mechanisms influenced by metal ions is dependent on the type of ion, specifically whether it engages in competitive interaction with the enzyme. For example, if the ion does not compete with the enzyme, enzymatic activity is induced. In contemporary times, if an enzyme participates in competitive interactions, its activity is reduced (Ghori et al. 2011 ). Fig. 3. Open in a new tab Impact of metal ions on the catalytic activities of the T1/F180 C and wt-T1 lipases. Lipase activity was identified using the Kwon and Rhee method. The untreated lipase activity in the absence of metal ions was used to compare the relative activities of the two lipases. The 100% activity represents the untreated enzyme for each lipase that is not depicted in the graph. SD is indicated as error bars. When the error bar cannot be seen, the deviation is less than the size of the symbol The impacts of organic solvents on the lipase activity of wt-T1 and T1/F180 C lipases The data analysis related to the stability of the T1/F180 C variant in different organic solvents revealed that it exhibited different tolerances compared with wt-T1 lipase (Fig. 4 ). The residual lipase activity of the untreated lipase was used as a control to calculate and compare the lipase activity in the organic solvents. The results demonstrated that the relative lipase activity of the T1/F180 C variant improved in the presence of dimethyl sulfoxide (DMSO) to 86%, compared with 67% for wt-T1 lipase. In addition, T1/F180 C exhibited increased residual lipase activity in the existence of benzene 75% compared to only 41% of wt-T1 lipase. Furthermore, the T1/F180 C variant showed slightly increasing activity in other organic solvents such as toluene, hexane, octane, and methanol, as it exhibited approximately 66%, 54%, 55%, and 42% activity compared with 47%, 39%, 47%, and 18% of wt-T1 lipase, respectively. However, other organic solvents such as acetonitrile, 1-propanol, and xylene have been found to negatively affect the activity of both lipases, with a slight difference in T1/F180 C activity compared with that of wt-T1 lipases. Moreover, all solvents showed downregulated lipase activity in both lipases, except for DMSO and benzene. The addition of 1-propanol and acetonitrile caused lipase denaturation resulting from the lipase structure; thus, the ability of the solvents to enhance the hydrolysis activity decreased in 1-propanol and acetonitrile. The residual activities of T1/F180 C after treatment with DMSO, benzene, toluene, n-hexane, octane, methanol, and xylene at 25% v/v concentrations reached approximately more than 50–80% which refers to high stability in organic solvents. Notably, the low log P value in the solvents could be the main reason for the reduced enzymatic activity, whereas the elevated log P value non-polar (non-polar) refers to the enhancement of enzymatic activity. In the present study, the T1/F180 C variant lipase exhibited improved stability in the presence of organic solvents. The variant and wt-T1 lipases were approximately stable in DMSO and benzene. However, it showed weak hydrolysis activity when subjected to acetonitrile, xylene, and 1-propanol. The residual activities of T1/F180 C have been reported to exceed 60% when subjected to many organic solvents such as DMSO, benzene, xylene, n-hexane, n-heptane, and toluene. Generally, amino acid alteration induces enzyme structural modifications, such as the establishment of novel hydrogen bonds and salt bridges, enhancement of hydrophobic core packing, and shifting of the side chain isoelectric point (pI). A previous study revealed enhanced metalloprotease PT121 stability toward acetonitrile and acetone after subjecting the enzyme to a directed evolution approach. The stability explanation assumed that the reason for the enzyme stability was the construction of an extensive non-covalent bond network, which improved the structural compactness (Zhu et al. 2020 ). In addition, T1 lipase derived from Geobacillus zalihae was subjected to a directed evolution strategy and subsequent site-directed mutagenesis, which resulted in a group of variants, Ala83 Asp/Met121Ile/Lys251Glu/Gly327Ser and Ala83 Asp/Met121Ile/Ser195 Asn/Thr272 Met, which exhibited higher stability in the organic solvent (methanol). The main explanation for the enhanced stability was the shortening of theradial distribution function distances, which enabled the construction of novel hydrophobic interactions and hydrogen bonds (Maiangwa et al. 2021 ). In another study, the substitution of a couple of arginine (Arg237 and Arg241) and a single residue of aspartate (Asp248) with which constructed salt bridges close to the active site resulted in the loss of the structure of couple variants (Arg237 Ala/Asp248 Ala) activity completely in 40% organic solvent within 1 h compared with the wild type, which maintained approximately 80% of the activity in organic solvents at a 40% ( v / v ) concentration. The considerable activity decline was assumed to be due to the loss of the two pairs of salt bridges constructed by (R237 A/D248 A), which is close to the active site. These double salt bridges are assumed to have an essential impact on preserving the correct catalytic conformation, and their loss has been shown to damage structural stability (Dachuri et al. 2022 ). Additionally, in the engineering of Geobacillus stearothermophilus T6 solvent tunnels for introducing a number of aromatic residues (Leu184Phe/Ala187Phe/Leu360Phe), which enhanced methanol stability, the assumed factor for enhancing stability was mutation that enabled the formation of novel π – π interactions (Gihaz et al. 2018 ). The enhanced stability of T1/F180 C in organic solvents was due to the increased rigidity of the lipase structure, as most newly constructed hydrogen bonds existed on the surface of the enzyme. In addition, the novel constructed aromatic interactions ( S / π interactions) were also located at the N- and C-terminal and could enhance the structure in solvent environments. These interactions facilitate the tight retention of water molecules on the enzyme surface, thereby reducing water stripping. Consequently, amino acids modulate the enzyme activity in solvent environments. Fig. 4. Open in a new tab Impacts of organic solvents on the wt-T1 l and T1/F180 C lipases catalytic activities. Lipase activity was identified using the Kwon and Rhee method.The activities of untreated lipase without organic solvents were used to compare the relative activities. The value of the activity was taken as 100% of the untreated enzyme. SD is indicated as error bars. When the error bar cannot be seen, the deviation is less than the size of the symbol The wt-T1 and T1/F180 C substrate specificity against p-nitrophenylester and various natural oils For identifying substrate preferences, different carbon lengths of p -nitrophenyl, including (C2–C18), were utilized in this study. Based on the obtained results, T/F180 C exhibited higher activity toward long-chain carbon p NP-C10–C18 with a preference toward p NP-dodecanoate (C14) compared to wt-T1 lipase, which has a preference toward C10–C18 with a high preference toward C12. The results indicate that T/F180 C lipase exhibits low activities toward short-chain carbon p NP-acetate (C2) and p NP-butyrate (C4) and a high preference toward long-chain carbons for hydrolysis ( Fig. 5 ) . A previous study reported that modifying the T1 lipase lid area via site-directed mutagenesis modified substrate specificity toward long-chain fatty acids (Tang et al. 2017 ). In addition, partial swapping of the Thermomyces lanuginosus lipase (TLL) lid domain with an esterase lid domain has been shown to significantly influence the lid region of amino acids. It altered the specificity of TLL lipase by exhibiting water-soluble substrate activity (Willems et al. 2018 ). Furthermore, altering the lid hinge residues modified lipase specificity toward longer chain lengths (Tang et al. 2015 ; Santarossa et al. 2005 ). Additionally, substituting the hydrophobic lid of R. chinensis lipase was replaced with that of an Aspergillus niger ferulic acid esterase has successfully altered lipase specificity toward water-soluble substrates and functioned as esterases (Yu et al. 2014 ). The altered T1 lipase specificity might be assumed to be due to the small size of the substituted cysteine amino acid compared with phenylalanine, as this residue is located on the lid and opposite the binding site, which might confer more space for fatty acids of larger length. Fig. 5. Open in a new tab The substrate specificities of the wt-T1 and T1/F180 C against various pNP-ester carbon lengths. The test was performed at 70 ℃ for 10 min. The p-nitrophenol product was monitored spectrophotometrically at 410 nm and was determined according to the pNP standard. All samples were triplicated for this assay, and SD is indicated by an error bar The substrate specificity toward various natural oils of wt-T1 and T1/F180 C lipases were also studied by comparing the hydrolytic activities of natural oils with those of olive oil. The hydrolysis reaction efficiency of lipases was varied according to the olive oil examined. The variant T1/F180 C lipase displayed a high preference for corn oil, canola oil, coconut oil, and rice bran oil, with relative activity levels of 219%, 160%, 141%, and 114%, respectively, compared with 133%, 120%, 70%, and 87% for wt-T1 -T1, respectively. However, T1/F180 C showed a lower preference toward soy bean and sunflower, with relative activity levels of 91% and 106% compared to 125% and 151% for wt-T1 (Fig. 6 ). A previous study reported that swapping the lid domain of Lip MRD via site-directed mutagenesis with Rhizopus delemar lipase lid domain residues altered the lipase specificity toward bulky fatty acid triglycerides. A previous study reported that altering W89 is assumed to be the key to increasing lipase activity by rendering more space for the binding site (Fernandez-Lopez et al. 2023 ). The binding site volume was also measured by YASARA software, as enclosed by the molecular surface of T1/F180 C was 883.81 Å 3 compared to 693.60 Å 3 of the wt-T1 Fig. 7 A, which refers to the space conferred by the binding site after substituting the Phe180 residue into Cys, which may reduce the steric hindrance effect. In addition, the distance between the (Phe180) of the wt-T1 lid domain and the catalytic residue (Ser113) was measured using YASARA, and the results are presented (8.11 Å). It is obvious that the substituted residue (Cys) increased these distances by (1.2 Å) as it showed approximately (9.31 Å) distances between the lid domain (Cys180) and the catalytic residue (Ser 113) Fig. 7 B. The T1 lipase Geobacillus zalihae , selectivity against p -nitrophenyl ester was modified from (C10-C12) to (C14-C18) and regioselectivity was modified from sn- 1,3 to only sn- 3 after substituting the lid domain residues (Phe180Gly and Phe181Ser) and the oxyanion residue Phe 16 Trp, as these mutations resulted in altering the binding site conformation and space, which blocked sn-1 and orientated the substrate to bind to only sn- 3 (Albayati et al. 2023 ). In addition, another similar study revealed that substituting the BTL2 lipase (96% sequence identity with wt -T1) lid domain with two consecutive bulky residues known as F181 and F182 enhanced the lipase specificity toward long fatty acids. During interfacial activation, these residues move away from the highly hydrophobic binding site and are exposed from the highly hydrophobic binding site via lipase transitions from closed to open conformation. This phenomenon is assumed to enable interaction with the lipid interface. This specificity modulation is proposed to belong to the expansion cavity space conferred by these residues because they construct the upper part of the binding site, thus enabling the accommodation of bulky fatty acids for hydrolysis by reducing steric hindrance (Carrasco-López et al. 2009 ). Fig. 6. Open in a new tab Natural oil hydrolysis specificities of the wt-T1 and the T1/F180 C variant. The test was performed at 70 ℃ for 30 min using the Kwon and Rhee method. The reaction of 1 μmol substrate per minute is considered one unit ( U ). The hydrolytic activity of lipase in the olive oil was determined at 100%. All samples were analyzed in triplicate and SD is indicated by an error bar Fig. 7. Open in a new tab A The surface of the wt-T1 and T1/F180 C lipases binding site structure. (a) wt-T1 lipase, (b) T1/F180 C variant. The catalytic residues are highlignted yellow and labeled white, and the light gray surface represents the binding site and binding pocket . The lid domain residue is yellow and labeled black. B Measured distances between the lid domains of wt-T1 and T1/F180 C lipases and their catalytic residue Ser 113. (a) wt-T1 lipase. (b) T/F180 C variant. The catalytic and lid domain residues are shown in yellow and black, respectively, and the lid domain is highlighted in blue Kinetic analysis of the wt-T1 and T1/F180 C variants To conduct kinetic studies on wt-T1 and its variant, the pNP-C12 substrate was utilized, which was performed according to previous and current studies of the wt-T1 substrate preference (Leow et al. 2007 ). This substrate was utilized to determine the reaction rate in the optimal T1 lipase buffer (Glycine NaOH pH 9.0) at an incubation temperature 70 ℃. The incubation time of the reaction mixture ranged from 0 to 12 min with 3 min intervals. The plotted Michaelis–Menten curve initial reaction rate was V 0 as a function of substrate concentration, and the inverted values of the curve were used to plot Lineweaver–Burk (Fig. S3 ). The substrate concentration and the values maximum velocity ( V max ) were detected on a linear plot at half V max ( K m ) corresponding to the catalytic lipase's efficiency and turnover rate ( k cat ). The rate of the enzyme product released from a specific amount of substrate is represented by the catalytic efficiency ( k cat / K m )to estimate lipase efficiency against the specific substrate. Table. S3 demonstrates that T1/F180 C variants displayed higher catalytic efficiencies than the wt-T1 lipase. The unique catalytic efficiency enhancement of F180 C might be due to the smaller side chain displayed by this residue compared to the bulky side chain residue Phe. A study conducted on BTL2 lipase revealed that the F180 residue on the flap domain was oriented toward the binding site, and it was assumed that activation of BTL2 was initiated by the interaction of F181, which is situated opposite to F180 of T1 lipase (Carrasco-López et al. 2009 ) and it covers the active site so it may occupy some space. The alteration of bulky residues with smaller side chains may enlarge the binding cavity space by conferring some space and reducing steric hindrance, which facilitates the access of the substrate. Moreover, this residue (F180) is assumed to be exposed to the solvent during the open conformation, thereby it is suggested that this amino acid residue is crucial for the interaction of the enzyme–substrate at the phase of the water–lipid interface (Tang et al. 2017 ). Additionally, it has been reported that the lipase catalytic efficiency can be enhanced by substituting the residues of the enzymes in the organic solvent environment (Yaacob et al. 2019 ). Secondary structure analysis of the wt-T1 and T1/F180 C variant lipases To understand the impact of this mutation on the secondary structure of T1 lipases, variants T1/F180 C and wt-T1 lipases were analyzed via circular dichroism far-UV spectrometry. The wt-T1 and its variant T1/F180 C circular dichroism (CD) spectra region (190–240 nm) were superimposed to compare the secondary structures using a Jasco J-815 CD spectrophotometer. The wt-T1 and T1/F180 C variant estimated secondary structures were estimated from the CD spectra at 20 ℃ and the content was analyzed as a proportion of β-sheet, α-helix, random coil, and turn by utilizing the software provided alongside the apparatus, as delineated in (Table S4 ). The analysis of secondary structures showed a considerable change in the T1/F180 C secondary structure compared to wt-T1 lipase. The variant of F180 C exhibited a 19.5% in α- helix fractions, which is slightly lower than the wt-T1 ratio, which accounted for 20%. However, β- sheet fragment showed an approximately 2% increase in T1/F180 C, exhibiting a 14.9% increase compared with the 12.9% increase in wt-T1 lipase. The β-turn fragment, displayed a 1.5% decrease in the number of the T1/F180 C variants, which approximately accounted for 23.9% of the lipase variants, compared with only 25.6% of the wt-T1 variants. The increases also included the random coil fragment as it increased for 0.2% of the T1/F180 C variant, which displayed 41.6% compared with 41.4 of wt-T1 lipase. Based on these superimposed structures, it is noteworthy that the F180 C variant has experienced a considerable structural change compared to the wt-T1 variant (Fig. 8 ). However, the analysis of the secondary structure by PDB sum revealed a decrease in the beta turn fragment, as it accounted for approximately 36 for T1/F180 C compared with 37 of wt-T1 lipase and this support the analyzed structure by the circular dichroism (CD). In addition, the number of gamma turns also decreased from 7 in wT-T1 to only 6 in the T1/F180 C variant (Fig. S4 a, b). Both lipases are composed of two band ranges that end with negative bands at 240 nm and start with positive bands at 193 nm. The R0-14 open conformation structure showed higher thermostability than the closed conformation structure, and this thermostability increment was assumed to be due to the α-helices fragment ration increasing. The NTU 03 lipase thermostability was modified after the forming α-helices, which improved the structural rigidity and thermostability (Shih and Pan 2011 ). The increment and decrease in the secondary structure ratio induced the folding and unfolding process, which resulted in helical and sheet content structural variations. A previous study revealed that high temperatures induce significant structural alterations (Kumar and Deshpande 2021 ). Furthermore, heating has been reported to induce the milk protein β-sheet to β-turn structure transformation and different temperature ranges have induced continuous transformation of the whole ratios of the secondary structure (Ye et al. 2017 ). In contrast, the thermostability of Penicillium cyclopium lipase was improved by conserving the β-sheets ratio (Liu et al. 2017b ). Fig. 8. Open in a new tab Secondary structure conformational changes released circular dichroism (CD) spectra analysis of T1/F180 C variant and wt-T1 lipases . The assay were conducted at 20 ℃, and pH 7.4 of 10 mM sodium phosphate buffer along with 0.1 mg/mL concentration of protein Analysis of the thermal denaturation point of the wt-T1 and T1/F180 C variant lipases Protein denaturation occurs as a consequence of protein aggregation and precipitation resulting from the exposure of the occluded hydrophobic amino acids to an incompatible environment containing solvents and water (Ahmad and Rao 2009 ). The stability of wt-T1 and its variant T1/F180 C was studied by conducting protein denaturation point (melting temperature) analyses, and this analysis was conducted by measuring the melting temperature ( T m ) within the range of 20–95 ℃. The wt-T1 and its variant melting temperature curves are depicted in Fig. S5 . Table S4 lists the thermal denaturation points of wt-T1 and T1/F180 C and their helical and sheet contents. The alteration of α-helix 6 which constitute the component of the flap domain, led to an elevation of the melting temperature ( T m ) to 78.2 of T1/F180 C compared to 73.5 °C of wt-T1 lipase. Substituting hydrophobic residues with hydrophilic residues in the lid area decreases the T m of T1 lipase (Tang et al. 2016 ). This decrease in Tm indicates that protein structural rigidity and packing were diminished by a reduction in the hydrophobic interactions between the flab and the enzyme binding site. In addition, the substitution of residues on the αC, β3, and β5 fragments of B. subtilis affects the hydrogen bond network, resulting in a decrease in melting temperature and thermostability, as a consequence of disturbing protein integrity (Zhang and Ding 2017 ). Moreover, a study reported that substituting the cold-active Pseudomonas AMS8 lipase flab residues T52Y and G55Y, which represent aromatic hydrophobic-polar amino acids, enhanced lipase stability, and it is assumed that these amino acid residues had a crucial role in the restriction of lid mobility and contributed to constructing an intramolecular H bond and rendering the structure highly stable (Yaacob et al. 2019 ). In silico structural investigation Atomic interaction analysis The T1/F180 C variant structure model was derived via homology modeling utilizing YASARA (Yet Another Scientific Artificial Reality Application) software, T1 lipase (PDB ID: 2DSN) as a template, and PyMOL software to predict and plot atomic interaction (Krieger et al. 2002 ; Matsumura et al. 2008 ). Based on the predicted structure of the T1/F180 C lipase variant, F180 was localized on the surface of the lid domain α-helix 6 ( Fig. 9 ) . The substitution of Phe 180 into Cys deleted specific hydrogen bonds and allowed the construction of novel bonds. However, the novel bonds were constructed around the entire surface; thus, the total number of bounds was 58 for wt-T1 and 51 for T1/F180 C (Table S5 ). Fig. 9. Open in a new tab Structure of wt-T1 lipase (PDB 2DSN) containing targeted amino acids. The figure was created using the Yasara software (Krieger et al. 2002 ). The blue helical structures represent the lid domain. The binding site is indicated by a green surface. The mutant residue is colored yellow The novel constructed hydrogen bonds representing Asp36 and Glu 38, Lys 229 to Glu 226, Lys 229 to Asp 178, Glu 284 to Asn 355, Glu 270 to Asn 355, Tyr 242 to Lys 251, Asp 311 to Arg 303, Asp 357 to Asn 288, and Tyr 302 to Asn 304 (Fig. S6 a–f), the whole-structure hydrogen bond distances increased from 0.1 to 0.3 Å in maximum range. The deletion and construction of the novel bonds also included salt bridges and aromatic bonds. Regarding the salt bridges, T1/F180 C formed two novel bonds and deconstructed five bonds. However, two constructed bonds, Arg 303 to Asp 311, and Asp 340 to Lys 344, were located on the C-terminal of the protein (Table S6 ), (Fig. S7 a, b). Substitution of Phe 180 to Cys also resulted in the deletion of three aromatic bonds and constructing novel two (S/π bond) are Phe 375 to Phe 370 and Tyr 29 to Trp 19, which are located on the N-terminal and C-terminal (Table S7 , Fig. S8 a–c). The analysis of the structures showed differences in the total atomic interactions, with changes in the bond distances and the locations of the novel constructed bonds. Based on the structure analysis, the enhanced thermostability variant T1/F180 C exhibited a lower number of atom interactions; however, the newly constructed bonds located on the N-terminal and C-terminal might be the reason for the enhanced lipase stability against high temperatures.The correlation between thermostability and molecular interactions was previously identified via the construction of extra interactions in lipase derived from Geobacillus stearothermophilus , which led to increased lipase thermostability (Dror et al. 2014 ). Atomic interactions such as H bonds and salt bridges play a pivotal role in protein folding and are considered essential for maintaining the catalytic structure, and the regulation of unfolding mechanisms occurs in these structures (Choo 2011 ). In addition, hydrophobic and aromatic interactions, along with molecular interactions, can rigidify and strengthen the protein structures. Protein structure rigidity is essential for enhancing protein stability in harsh environments, such as those involving organic solvents and high temperatures. Many researchers have reported the enhancement of protein stability by engineering atomic interactions such as ion-pair interactions and hydrogen bonds through amino acid exchange (Wu et al. 2015 ; Lan et al. 2015 ). Substituting asparagine into threonine (T118 N) resulted in the destabilization of the protein structure, melting temperature, and stability due to a decrease in the molecular interactions number (Charbonneau and Beauregard 2013 ). In addition, ion pairs play pivotal roles in stabilizing single-α helix (SAH) domains in solvents. This study revealed that these-helix ion pairs exhibit dynamic behavior, rapidly constructing and deconstructing bonds, and this process occurs alternatively within the helix different partner residues (Batchelor et al. 2019 ). A previous study reported that lipase I from Penicillium cyclopium (PCL) was subjected to a directed evolution approach that enhanced its thermostability to within 5 °C. Analysis of the thermostable lipase structure revealed that hydrophilic and hydrophobic interactions along with proline substitution and helix propensity stabilize the structure (Liu et al. 2017a ). The aromatic residues play an essential role in protein structures and are significantly involved in protein stability. These interactions are essential for vital processes such as protein folding, molecular recognition, and ligand binding (Kumar et al. 2023b ). MD simulation of wt-T1 and T1/F180 C lipases MD simulations were performed in triplicate (Table S8 ) to investigate the effects of temperature on the dynamics of the enzyme, and the simulation outcome was analyzed using the root mean square deviation (RMSD), root mean square fluctuation (RMSF), radius of gyration (Rg), and solvent accessible surface area (SASA) values. The RMSD values, which were calculated for 100-ns trajectories on the C atoms of wt-T1 and T1/F180 C lipases were analyzed at 70, 75 and 80 °C (Fig. S9 ). At the beginning of the trajectory ( t = 0 ns) at 70, 75 and 80 °C, RMSD displayed a value of about 0, and 0.001 Å for both wt-T1 and T1/F180 C lipases, respectively. This indicates the C-backbone movement of both lipase structures during the thermalization and equilibration periods. During the simulation at 70 °C, the RMSD value of wt-T1 and T1/F180 C lipases fluctuated within the range of 0.008 to 0.176 Å but T1/F180 C lipase structure stabilized at a range of 0.008 to 0.163 Å until the end of the simulation ( t = 100 ns). At 4 ns, the wt-T1 RMSD increased rapidly to a value of 0.159 Å, while the RMSD of T1/F180 C was only 0.137 Å at the same trajectory. The structure of T1/F180 C exhibited the highest RMSD of 0.161 at the time of 25 ns compared to the wt-T1 RMSD of 0.137 Å. Moreover, at a time of 68 ns, the RMSD of wt-T1 increased to 0.160 compared to 0.137 Å of T1/F180 C. In addition, the RMSD of wt-T1 increased to 0.170 and 0.176 compared to 0.157, and 0.161 Å of T1/F180 C at 89 and 96 ns. Overall, the RMSD of T1/F180 C had a lower RMSD range than that of wt-T1 for 100 ns at 70 °C, which might be due to the more stabilized structure of T1/F180 C compared to wt-T1. Regarding the simulation result at 75 °C, the structure of the mutant T1/F180 C showed greater stability, as it displayed lower RMSD values than wt-T1. Both structures fluctuated across the simulation period of 100 ns, but the fluctuation range of wt-T1 was between 0.097 and 0.226 Å, compared to the fluctuation range of the mutant T1/F180 C, which was between 0.086 and 0.160 Å (Fig. S9 ). During the simulation, the structure of wt-T1 increased to 160, 167, and 0.220 Å, at 2, 22, and 48 ns compared with the mutant structure of T1/F180 C, which increased to 0.142, 0.146, and 0.151 Å respectively. Furthermore, at the end of the simulation at 98 ns, the structure of the mutant T1/F180 C showed a lower value of 0.111 compared to 0.151 Å of wt-T1 lipase. During the simulation at 80 °C, at 5 ns the structure of wt-T1 spiked to 0.126 compared to 0.168 Å of the mutant structure T1/F180 C, and this was the lower fluctuation point revealed by wt-T1. Thereafter, the structure fluctuated within the range of 0.169–0.231 Å compared to 0.130 to 0.179 Å of the mutant structure T1/F180 C. Therefore, these computational findings exhibit a positive correlation between the mutant structure and the experimental results, indicating that the variant is comparatively more stable and exhibits high catalytic performance at high temperatures. Local dynamic changes around the mutation site were measured to examine the flexibility of the residues, which is known as the RMSF of the C backbone against each residue. The RMSF curve exhibited a similar pattern to the RMSD of the C backbone per residue for most of the 100 ns at the three conducted simulation temperatures of 70, 75 and 80 °C (Fig. S10 ). In agreement, at 70 °C simulation, no obvious change in the RMSF curves between wt-T1 and T1/F180 C was observed. The largest fluctuation amplitude occurred near the N-terminal and the highest fluctuation occurred close to the C-terminal. Moreover, most of the fluctuation occurred near the surface and apart from the catalytic site. In the simulation at 70 °C, the fluctuation range remained within 0.1 to 0.2 Å except for many peaks that exceeded this range. The highest peak observed for wt-T1 was 0.514, compared to 0.332 Å of the T1/F180 C. This peak represents β – turn and involve the residues 272–284. The second peak was observed near the N-terminal and represented α -helix and residues involved 32–50. Overall, the fluctuation patterns for both structures were similar, with slight deviations. The simulation outcomes for both structures at 75 °C, revealed lower fluctuation range than in the previous simulation at 70 °C. The highest spike range of wt-T1 lipase was 0.3 Å, compared to the range of the highest peak of 0.36 Å of the mutant T1/F180 C. The represented figure revealed two peaks both of them on the surface, and the first one represents coil turn and involves the 98 to 104 for both structures, and the peak value for the wt-T1 was 0.307 compared to 0.302 Å of the T1/F180 C mutant. The second peak was next to the first peak and represent the α -helix and involved 128–146 residues. The peak value was 0.235 compared to the 0.369 Å of the T1/F180 C mutant. The simulation patterns of both structures were similar and did not exhibit any notable deviations. The simulation results at 80 °C revealed similar fluctuation patterns for both simulated structures. Both the wt-T1 and T1/F180 C structures revealed three peaks, the first representing the coil turn, which involved residues 100–107; the value of wt-T1 was 0.424 compared to 0.318 Å of the T1/F180 C mutant. The following peak was next to the first peak and represent the α -helix and the value of wt-T1 was 0.342 compared to 0.318 Å of the T1/F180 C structure. The last spiked peak represented a combined structure of the coil turn and α- helix and involved residues 226–240 and the value of the peak of w-T1 was 0.287 compared to 0.227 Å of T1/F180 C. Overall, all spiked peaks were located on the surface, and both simulated structures displayed similar fluctuation patterns. Regarding the structural compactness of the two lipases at three different temperatures, the Rg plot revealed an important difference between wt-T1 and T1/F180 C at the simulation temperature (Fig. S11 ). At all three temperatures, the Rg values of wt-T1 were higher than those of its T1/F180 C variant. At 70 °C, the radius of wt-T1 expanded to a range of 1.759 Å compared to the 1.749 Rg value of T1/F180 C. Meanwhile, at 75 °C, the highest Rg value for wt-T1 was 1.76 compared to the Rg value, which decreased to 1.73 Å. Furthermore, at 80 °C, the Rg value of the wt-T1 initiated with a low value of 1.68 Å, and ended with a higher value of 1.744 Å. In contrast, the T1/F180 C variant started with 1.74 Å and ended with 1.68 Å. The reduction in the Rg value indicates that T1/F180 C is structurally more compact than its wild-type. The solvent accessible surface area (SASA) of the wt-T1 and T1/F180 C lipases were computed using an analytical algorithm for 100-ns trajectories in the water system at 70, 75 and 80 °C (Fig. S12 ). In general, similar patterns of SASA values within a close range (150–166 Å 2 ) were recorded for both lipases during the simulation at different temperatures. However, a slight increase in SASA to a value of 170 Å 2 was noticed at 75 °C for T1/F180 C, suggesting that the folding mechanism in the mutant lipase structure occurred more efficiently than in the wt-T1 structure with 160 Å 2 . However, both simulated lipases exhibited a close range of SASA value reached 165 Å 2 , at 80 °C. The elevated simulation temperatures did not significantly affect the SASA values of the lipase structures. One explanation could be that the accessible surface encloses the volume (Å 3 ) of each lipase is comparable. When proteins exhibiting greater flexibility and specific structural modifications become more susceptible to unfolding and denaturation. Upon unfolding, the proteins experience significant solvation and become more flexible and less compact (Karshikoff et al. 2015 ). MD simulations can demonstrate the dynamics of lipase at different temperatures. At 75 °C and 80 °C, variant T1/F180 C exhibited a lower RMSD value, whereas wt-T1 exhibited a higher RMSD value throughout the simulation period, especially at 80 °C. The flexibility of the residues around the modified Phe 180 was lower and near to described as slight fluctuations at the three simulated temperatures, except for a few peaks near the N-terminal and C-terminal, suggesting a more rigid lid domain after modification. A previous study reported that both the flexibility and rigidity of protein molecules are necessary to run a function; however, integrated rigidity and compactness must compensate for higher thermal fluctuations (Karshikoff et al. 2015 ). The enhanced compactness of the T1/F180 C structure, as demonstrated by its low Rg value, can be attributed to the enhanced stabilization of the loops and secondary structural elements via intramolecular interactions. Furthermore, most rigid thermostable proteins remain in their native folded state at elevated temperatures (Qiu et al. 2015 ). The surface area of a lipase is known as the solvent accessible surface area (SASA), and it is used to calculate the free energy transformation from an aqueous or polar solvent to a nonpolar solvent within a lipid environment (Adamczak et al. 2004 ). A previous study reported that thermostable enzymes typically exhibit reduced accessible areas to facilitate adaptation at elevated temperatures (Gianese et al. 2001 ). A significant decrease in the SASA value typically indicates a good folding process, and the hydrophobicity of atoms generally refers to their accessible surface area (Momen-Roknabadi et al. 2008 ). In addition, the substitution of phenylalanine with cysteine attenuated the interaction between the protein surface and the solvents. A previous study assumed that the entry of water molecules into protein structures destroys the hydrophobic forces. Because of this entrance, the solvent-accessible surface area expands in the hydrophobic region, and with the existence of pressure, the protein unfolding process begins (Okumura 2012 ). Thus, protein surfaces with extensive hydrophobic interactions must maintain their catalytic conformation against solvent intrusion. Methodology Rational design of mutant lipase in the lid domain Amino acid residues at positions 180 in the lid domain of T1 lipase were screened for convenient residue substitution using DynaMut2 , which predicts the effect of mutation on the protein dynamics and stability ( https://biosig.lab.uq.edu.au/dynamut2/ ) (Rodrigues et al. 2021 ). We found that Cys is a suitable candidate for residue substitution based on the signs of protein stability free energy change (ΔΔ G Stability ). This residue also has a molecular structure with small side chains that eliminates stearic hindrance and improves lipase specificity. Engineering T1 lipase thermostability in the lid domain by rational design This strategy was utilized to enhance T1 lipase thermostability by mutating the bulky side chain amino acid F180 on the lid domain, which is located on the α-helix 6 fragment, via a rational design (site directed mutagensis). Residue F180 was substituted with the Cys amino acid variant (F180 C) by utilizing the FASTA file corresponding to the wt-T1 lipase (PDB ID 2DSN), which was obtained from the PDB website and employed to engineer the mutagenic codon primer. The engineered mutagenic codon primer was used with a QuikChange Lightning Site-Directed Mutagenesis Kit (Agilent, USA) (Table S1 ). XL10-Gold ultracompetent cells were used to transform the PCR products. Subsequently, the plasmid was extracted utilizing the MYgen plasmid miniprep kit (Gene Xpress PLT, Selangor, Malaysia), followed by sequencing to validate and confirm the mutation sites'existence (Fig. S1 ). The expression host Escherichia coli strain BL21(DE3) was utilized for positive clones sub-transformation. . Lipase expression and purification Bacteria harboring the mutated plasmid E. coli strain BL21(DE3) were inoculated into 10 mL of Luria Bertani (LB) broth and added 100 g/mL ampicillin. The bacteria were grown at 37 °C and shaken at 150 rpm overnight. The microbial cultures were then subsequently transferred into 200 mL of freshly prepared similar medium until reaching an optical density at 600 nm (OD 600) of 0.6–0.7, at which point induction was performed by the addition of 0.025 mM of isoprophyl-l-d-thiogalactopyranoside (IPTG). The grown cells were harvested after 12 h of culture at 37 °C by centrifugation at the speed of 10,000 χg for 30 min at 4 °C. The cell pellets were resuspended in 20 mL of 20 mM sodium phosphate, 0.5 M NaCl, and 0.5 mM imidazole, pH 7.4. The resuspended pellets were sonicated and then centrifuged at 10,000 χg at 4 °C for 30 min, and the supernatant was purified by nickel sepharose affinity chromatography. To verify and determine the purity and homogeneity of the protein, approximately 0.1 g of each eluted protein was loaded into a 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) (Fig. S2 ). Determination of protein concentration and lipase assay The Bradford method was used to calculate the protein concentrations using the Bradford reagent (AMRESCO®, Avantor, Ohio, OH, USA) (Bradford 1976 ). By utilizing the colorimetric assay according to the modified Kwon–Rhee method, a lipase biochemical assay was executed (Kwon and Rhee 1986 ). Olive oil (Bertolli, Deoleo S.A., Córdoba, Spain) was emulsified with a buffer solution as a substrate emulsion at a ratio of 1:1. The lipase concentration used during the assay was standardized to 1 mg/mL. The assay mixture in the final preparation contained 990 µL of the buffer (50 mM Glycine–NaOH pH 9.0), 10 µL of the enzyme, 20 µL of 20 mM calcium chloride (CaCl 2 ), and 2.5 mL of the substrate emulsion. The incubation period of the prepared mixture was 30 min with 200 rpm shaking at 70 °C. Subsequently, the reaction was terminated after 30 min of incubation by adding 1 mL of 6 N HCl, and this was followed by the addition of 5 mL of isooctane. To separate the buffer containing the released product (oleic acid) and oil, the reaction mixture was vortexed and left at room temperature for 30 min. The released enzyme product was separated in the upper layer of the reaction mixture and mixed with 1 mL of copper pyridine solution pH 6.1 The final mixture was placed in another vial to calculate the enzyme activity by measuring the absorbance at 715 nm. Activity was calculated as one unit ( U ) of enzyme catalyzed by one mol of substrate per minute during the reaction. Optimum temperature study of the lipase variant The lipase variant activity was tested using olive oil as a substrate at 40 °C–90 °C with an agitation rate of 150 rpm. A CD spectropolarimeter J-815 (Japan Spectroscopic Company (JASCO), Tokyo, Japan) was used to determine the melting temperature ( T m ). The enzyme concentration was fixed at 0.1 mg/mL in 10 mM sodium phosphate buffer at pH 7.4. The warming period was set from 20 °C to 95 °C with a heating rate of 1 °C/minat a wavelength of 222 nm, and before each run, a one-cm optical path length was used. The melting temperature analysis was based on the Greenfield protocol (Greenfield 2006 ). The stage at which 50% of the protein sample was denatured was defined as the melting temperature ( T m ). Effect of pH on the lipase activity and stability The optimum stability of the lipase variant was identified by measuring the hydrolytic activity following the standard lipase assay after pre-incubating the enzymes in different pH buffers for 30 min at 60 °C. Wide-pH range was implemented by utilizing various buffers: 50 mM sodium acetate buffer, pH 4.0–6.0; 50 mM sodium phosphate buffer, pH 6.0–7.0; 50 mM Tris–HCl, pH7.0–9.0; 50 mMGlycine-NaOH buffer, pH 9.0–11.0), and 50 mM disodium phosphate, pH 11–12. Effects of metal ions on lipase stability The presence of metal ions was used for studying the lipase stability and was performed by measuring the lipase activity after separate pre-incubation with zinc chloride (ZnCl 2 ), calcium chloride dehydrate (CaCl 2 2H 2 O), iron(III) chloride (FeCl 3 ), magnesium chloride (MgCl 2 ), sodium chloride (NaCl), copper(II) chloride (CuCl 2 ), and nickel(II) sulfate hexahydrate (NiSO 4 6H 2 O), The samples’ stability was identified by following the standard lipase assay after pre-incubation in 1 and 5 mM of metal ions at 60 °C for 30 min. The relative activities of the treated enzymes were measured and calculated compared with those of the untreated enzymes (100%). Effect of organic solvent on lipase stability Various organic solvents (25% v/v) were used for pre-incubation of the purified lipase for 30 min at 60 °C under 150 rpm of shaking to test the lipase stability. The boiling points greater than 60 °C and log P values (values in parenthesis) were used as the bases of the organic solvent selection. The organic solvents included ethanol (0.2), DMSO (1.4), acetonitrile (0.4), methanol (0.7), 1-propanol (0.3), xylene (3.1), benzene (2.0), toluene (2.5), n-heptane (4.0), octanol (2.9), and n-hexane (3.5). A standard lipase assay was used to measure the residual activity using olive oil (emulsified, 1:1 v/v in buffer) at 70 °C. The remaining activity relative to the control (without solvent) was used to measure the lipase stability. Substrate specificity and hydrolysis activity of lipases in various natural oils The substrate specificity of wt-T1 and T1/F180 C variant lipases was detected utilizing p -nitrophenyl esters ( p NP) with different chain lengths, including p NP-acetate, which includes (C2), p NP-butyrate (C4), p NP-decanoate (C10), p NP-dodecanoate (C12), p NP-myristate (C14), p NP-palmitate (C16), and p NP- stearate (C18). The reaction mixture contained 0.188 mL of 50 mM Glycine NaOH (pH 9), supplemented with 1% (w/v) gum Arabic, 0.01 mL of 5 mM p NP-ester with different alkyl chain lengths, and 0.002 mL of 1 mg/mL purified enzyme. The reaction was initiated by adding the enzyme and terminated by adding 10 µL of 1% SDS after 10 min. The hydrolysis of the p -nitrophenol product was monitored spectrophotometrically at 410 nm and was determined according to the p NP standard. The amount of enzyme releasing 1 µmol of p -nitrophenol per min was measured as one unit of enzyme activity ( U ) under the experimental conditions. The hydrolytic activities of wt-T1 and its variant T1/F180 C toward different natural oils were measured by colorimetric assays according to the modified Kwon and Rhee method described previously (Kwon and Rhee 1986 ). Lipase substrate preference was measured using natural oils such as coconut oil (C12:0), olive oil (C18:1), rice bran oil (C18:1), soy oil (C18:1), corn oil (C18:2), sunflower oil (C18:2), and canola oil (C18:1). The reaction mixture was prepared by emulsifying each oil and buffer at a 1:1 ratio. A mixture of a solution containing 2.5 mL of the substrate,10 μL of the enzyme, 20 μL of 20 mM calcium chloride (CaCl 2 ), 2.5 mL of the substrate, and 990 μL of buffer was assayed at 70 °C for 30 min under 200 rpm agitation. Lipase activity was determined by measuring the optical density at 715 nm. The rate of enzyme catalyzing the reaction of 1 μmol substrate per minute is considered one unit ( U ). The hydrolytic activity of lipase in the olive oil was determined at 100%. Kinetic analysis of the wild-type (wt-T1) lipase and its variant The lipase hydrolytic activity toward p NP-dodecanoate (C12:0), which is the preferred substrate of the wild-type T1 lipase, was implemented for the kinetic study with slight modification. Various concentrations of the substrate ( p NP-dodecanoate (C12:0) was used, ranging from 0.1 to 7 mM of 100% 2-propanol. An enzyme concentration of 0.005 mL of 0.01 mg/mL was used to start a reaction mixture composed of 0.190 mL of 50 mM glycine NaOH with pH 9, 0.005 mL of 0.1 mM p NP-dodecanoate (C12:0), and supplemented with 1% (w/v) Arabic gum. A 1% SDS was used to terminate the reaction after incubation at the optimum temperature (70 °C) of lipases with 3 min intervals for a time ranging from 3 to 12 min. The experimental procedure was repeated at various concentrations of the substrate p NP-dodecanoate (C12:0) in triplicate and at different incubation times as previously described. The amount of lipase released from the lipase hydrolytic activity was measured at 410 nm and calculated according to the p NP standard. One unit ( U ) is the quantity of lipase that produces 1 µmol of p -nitrophenol in 1 min. Homology modeling and structural prediction The wt-T1 and T1/F180 C variants were remodeled using the PDB structure of T1 lipase (PDB ID: 2DSN) as a template. The T1/F180 C structures were predicted using an automated homology model developed by Yet Another Scientific Artificial Reality Application (YASARA) (Krieger et al. 2002 ). The energy minimization of the predicted model was performed by 10.000 steps using the steepest-descent minimization algorithm in YASARA. The volume of the binding site of wild-type T1 and variant T1/F180 C was measured as enclosed by the molecular surface using YASARA software. The residues at the binding sites were manually identified and plotted using the YASARA software. The prediction of secondary structure was performed using the web server PDBsum https://www.ebi.ac.uk/thornton-srv/databases/pdbsum/ . The prediction of the hydrogen bonds, salt bridges, and aromatic interactions was conducted using the PyMOL software https://www.pymol.org/ . Molecular dynamic simulations Molecular dynamics (MD) simulations were executed using the Gromacs software (Case Hma et al. 2021 ) and an amber ff19SB force field was assigned to the protein. These simulations evaluated the stability and flexibility of the wt-T1 and T1/F180 C lipase variants. Each molecule was solvated in a cubic water box measuring 90.0 × 90.0 × 90.0 nm, with water molecules modeled using the TIP-3p water model. The sodium ions were added to neutralize the system by replacing water molecules to ensure charge neutrality. Before running the MD simulations, an energy minimization step was performed using the steepest descent algorithm, which involved 635 steps for the wt-T1 lipase variants and 638 steps for the T1/F180 C lipase variants, to remove steric clashes and optimize the geometries. Following energy minimization, the system underwent a two-phase equilibration: NVT (constant Number of particles, Volume, and Temperature) and NPT (constant number of particles, pressure, and temperature), each for 5 ns (ns). During the NVT phase, a position-restraint MD simulation was applied with a constant force of 1000 kJ mol −1 nm −2 on the protein heavy atoms to maintain system stability and adjust internal energies and atomic fluctuations, ensuring constant temperature. Post-equilibration, the systems were prepared for a 100-ns production run at temperatures of 343.15, 348.15, and 353.15 K and a pressure of 1 atmosphere. Various parameters were analyzed to understand the dynamic behavior and stability of each molecule: the root mean square deviation (RMSD) was calculated to measure the deviation of each structure from the starting conformation throughout the simulation. The radius of gyration (Rg) determined the compactness and overall size of the complexes during the simulation. The root mean square fluctuation (RMSF) provides information about the flexibility and fluctuation of individual residues within a molecule. The solvent-accessible surface area (SASA) was calculated to assess the exposure of the lipase variant to the surrounding solvent molecules. Furthermore, the hydrogen bond profile was analyzed to study the formation and stability of the hydrogen bonds in lipase. These analyses were performed using built-in tools in Gromacs, which is widely used for molecular dynamics simulations. By examining these parameters, we gained insights into the dynamic behavior, stability, and intermolecular interactions of the lipase variants throughout the 100-ns production run at various temperatures. Conclusion In this study, the targeted specificity of lipase was improved simultaneously with thermostability via site-directed mutagenesis using a web server that analyzes and visualizes the effect of a single mutation on protein dynamics and stability. The lipase was successfully purified to homogeneity using single-step affinity chromatography, along with the wt-T1 lipase, for comparison. The biochemical and biophysical studies on wt-T1 and its T1/F180 C variant revealed that the T1/F180 C variant successfully enhanced lipase specificity toward long fatty acid chains coupled with thermostability enhancement. Variant structure analysis revealed novel atomic interactions on the surface of the lipase, and these interactions were substituted with those previously observed in the wt-T1 structure. These atomic interactions include hydrogen bonds, salt bridges, and aromatic ( S / π ) interactions, which are associated with single point mutations. Thus, a single mutation can induce the exchange of existing atomic interactions with novel interactions that play an important roles in the conformation, specificity, and thermostability of lipases. The T1/F180 C variant induces the formation of novel hydrogen bonds and salt bridges on the lipase surface and aromatic interactions near the N- and C-terminal. These novel molecular interactions enhanced the structural stability of the variant, which was represented by an increase in the activity of DSMO, benzene, octane, hexane, toluene, and methanol compared with that of wt-T1 lipase. Moreover, this variant exhibited a decrease in the activity of xylene, 1-propanol, and acetonitrile. The MD simulation results showed that the T1/F180 C variant was more stable than the wt-T1 lipase, which supports the biochemical and biophysical results. Regarding lipase specificity, the T1/F180 C variant successfully reduced steric hindrance by exhibiting enhanced selectivity toward C14 fatty acids, corn oil, canola oil, and coconut oil. Therefore, our study provides a better understanding of the relationship between the structure and function of lipases, which may have advantages for biotechnological and industrial applications. Statical analysis The analysis of quantitative data in current study was conducted in GraphPad Prisim 9 (GraphPad software Inc., CA, USA).The data were represented as the mean ± SD. The statistical comparisions were initialy carried out by one-way ANOVA, followed by T-test, in which P -values of < 0.05 were consided statically significant. Supplementary Information Below is the link to the electronic supplementary material. Supplementary file1 (DOCX 17000 KB) (16.6MB, docx) Supplementary file2 (DOCX 39 KB) (38.6KB, docx) Author contributions Conceptualization (Samah Hashim Albayati, Raja Noor Zaliha Raja Abd Rahman); data curation (Samah Hashim Albayati, Muna Hilal Shabeeb); formal analysis (Samah Hashim Albayati, Oluwasola Michael Akinola); funding acquisition (Raja Noor Zaliha Raja Abd Rahman); investigation (Samah Hashim Albayati, Muna Hilal Shabeeb); methodology (Samah Hashim Albayati, Adam Leow Thean Chor,Mohd Shukuri bin Mohamad Ali, Fairolniza binti Mohd Shariff, Noor Dina binti Muhd Noor); project administration (Samah Hashim Albayati, Raja Noor Zaliha Raja Abd Rahman); resources and software (Oluwasola Michael Akinola, Adam Leow Thean Chor); supervision (Samah Hashim Albayati, Raja Noor Zaliha Raja Abd Rahman, Adam Leow Thean Chor, Mohd Shukuri bin Mohamad Ali); validation (Samah Hashim Albayati, Oluwasola Michael Akinola, Raja Noor Zaliha Raja Abd Rahman); Visualization (Samah Hashim Albayati, Nima Ghahremani Nezhad); roles/writing—original draft (Samah Hashim Albayati, Muna Hilal Shabeeb, Nima Ghahremani Nezhad); and writing—review and editing (Samah Hashim Albayati, Muna Hilal Shabeeb, Nima Ghahremani Nezhad, Raja Noor Zaliha Raja Abd Rahman). Funding This research was supported financially by the Ministry of Education of Malaysia. (FRGS/1/2019/STGO5/UPM/01/1/01-01-19-2178 FR). Availability of data and materials Data are available on request. Declarations Conflict of interest The authors declare that there is no conflict of interest regarding the publication of this article. References Adamczak R, Porollo A, Meller J (2004) Accurate prediction of solvent accessibility using neural networks–based regression. Proteins 56(4):753–767. 10.1002/prot.20176 [ DOI ] [ PubMed ] [ Google Scholar ] Ahmad S, Rao NM (2009) Thermally denatured state determines refolding in lipase: mutational analysis. Protein Sci 18(6):1183–1196. 10.1002/pro.126 [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] Akoh CC, Lee G-C, Shaw J-FJL (2004) Protein engineering and applications of Candida rugosa lipase isoforms. 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