Biochimica et Biophysica Acta (BBA) - Protein Structure and Molecular Enzymology
ReviewLipase protein engineering
Introduction
Lipases are enzymes, which catalyse the hydrolysis or formation of lipids. The lipase discussed in the present review has mainly the triacylglycerol activity and is classified in the EC 3.1.1.3 group. The reaction is shown by the following equation:The lipases are a versatile group of enzymes and often express other activities like, e.g., phospholipase, lysophospholipase, cholesterol esterase, cutinase, amidase and other esterase type of activities [1], [2], [27]. Generally lipases have preference for the substrate type, whether it is a triglyceride or a diglyceride, and therefore have diglyceride and monoglyceride as products, rather than the glycerol and fatty acids alone. The regioselectivity is often rather high for the positions sn1 and sn3 and less frequently the sn2 is degraded (see Fig. 1) [1], [3], [4]. The lipase enzymes consist of a large family showing the same overall structural fold [5], [28], but which has a versatility of loop structures in contact with the substrate, and exhibits versatile substrate specificities. Another definition of lipases often used is lipolytic enzymes, which are capable of hydrolysing lipid substrates, thus including phospholipases, cutinases or enzymes hydrolysing ester substrates of lipid character. Protein engineering of selected other lipases, hydrolases or enzymes and proteins having homologous X-ray structures is also occasionally included in this review. For general protein engineering references including lipases see [30], [35].
Lipases are enzymes with general interest within many industrial applications. Lipases are used within the industry, e.g., detergents, oil and fats, baking, organic synthesis, hard surface cleaning, leather industry and paper industry [1], [2], [6], [7], [8], [9], [41], [58].
Protein engineering of triacylglycerol lipases has been done since the mid-1980s. The first example on protein engineering of a lipase is the work on the Pseudomonas mendocina lipase [10]. The early work was done based only on sequence information. When the first structures became available in the late 1980s the protein engineering interest increased dramatically. A European funded project, as well as many academic and industrial laboratories around the world, focused on solving new lipase structures of the enzymes Rhizomucor miehei lipase [11], Humicola lanuginosa lipase [12] (now Thermomyces lanuginosa lipase), Pseudomonas glumae lipase [13], Fusarium solani pisi cutinase [14], Candida antarctica B lipase [15] and Candida rugosa lipase [16] (or Candida cylindracea lipase), and most of these enzymes became targets for protein engineering. Now more than 12 X-ray structures of lipases are available in the protein database (PDB), of which 10 are microbial. Many of the lipases are solved in both a closed and an open conformation, i.e., with the lid or lids displaced from the active site. The Rhizomucor miehei lipase [11] and Human pancreatic lipase [17] X-ray structures was the first to be published. The Rhizomucor miehei lipase X-ray structure was found to have an active site triad as in the proteases [11]. Later by solving the X-ray structure in a form inhibited by a phosphonate inhibitor [18], which covalently binds to the active site Ser, it was revealed that the lid was displaced from the active site by a hinge bending movement [20], creating an increased hydrophobic surface [21]. Other lipases have also been crystallised with a number of different substrate analogous like the Candida antarctica B lipase [22], Human pancreatic lipase [23], Pseudomonas sp. lipase [24], Fusarium solani pisi cutinase [25] and Candida rugosa lipase [19], [26]. A list of selected solved lipase structures is given in Table 1.
From many of these structures lipases with homologous sequences can be model build, using adequate programs as, e.g., Homology or Modeller from MSI [27], or using threading programs [28]. Protein engineering of other lipases has also been performed, e.g., of the lipoprotein lipases [29] and the phospholipases, especially the phospholipase A2 (PLA2) [30]. Within the mammalian lipases many point mutations have been isolated from various sources including humans. These mutations are responsible for several diseases related to the lipase function [31]. Surface charge variants in Humicola lanuginosa lipase have been addressing identification of epitopes of the wild-type enzyme [43].
The overall structure of the triacylglycerol lipases can be described as a structure with a central β-sheet with the active serine placed in a loop (see Fig. 2), termed the catalytic elbow. Above the serine a hydrophobic cleft are present or formed after activation of the enzyme [32], [33]. The hydrophobic cleft is an elongated pocket suitable for acyl moieties to fit into. The activation, which is often necessary for the lipase enzyme, is a movement of a lid or several lids. This process is a part of the activation of activity, often referred to as interfacial activation [34], which takes place above the critical micellar concentration of the substrate. In Humicola lanuginosa lipase the opening of the lid can be described as a hinge bending motion of a helical lid. For other lipases the activation is more complex, e.g., Human pancreatic lipase [23] and Candida rugosa lipase [36], having both more than one lid or flap. In addition, activation is only present for certain substrates, e.g., Fusarium solani pisi cutinase, has a small structural change associated with long chained acyl substrates [36], whereas there is no or little change of structure upon binding of phosphonate inhibitor for the more esterase like enzyme of Candida antarctica B lipase [22]. The lipases belong to the α/β hydrolase family [5]. The α/β hydrolase family contains enzymes within lipases, esterases, proteases, peroxidases and lyases and others [5], [37]. Not all lipases belong to the same structural families, as, e.g., the PLA2, PLA-D, and some lysophospholipases, having very distant structures compared to the triacylglycerol lipases.
The hydrophobic surface area is placed centrally in the lipid contact zone. This lipid contact zone has been addressed by protein engineering. To locate the lipid contact zone, cysteine residues have been engineered into the surface of PLA2 [38]. Binding of spin labels to the cysteines and measuring the ESR signal perturbation on binding to the lipid surface, indicates a special orientation of binding to the macroscopic substrate surface. The suggested lipid contact zone of the Humicola lanuginosa lipase is shown in Fig. 3. In the Humicola lanuginosa lipase the lipid contact zone contains several big hydrophobic residues like Phe, Trp, Ile, Leu and Tyr. Some of the Phe and Ile residues are suggested to function as lipid anchors, and thus probably penetrates into the hydrophobic part of the lipid surface. The lid is also a central part of the lipid contact zone of most triacylglycerol lipases. The displacement of the lid increases the hydrophobic surface area of the lipases dramatically within the lipid contact zone area. A number of lipase structures with their lid or lids highlighted are shown in Fig. 4.
The lipases can be grouped into subfamilies by sequence homology analysis [39], [40], [41] based on sequence homology, thus dividing them into two main families, the mammalian and the microbial lipase family. Within microbial lipases, several families have now been found, the bacterial lipases, containing the Staphylococcus lipase family, Pseudomonas lipase family, Bacillus lipases and others, and the fungal lipases with the Rhizomucor miehei lipase family, Candida rugosa lipase family [39], [41], [42], and other subfamilies.
Within microbial lipases the fungal family of Rhizomucor miehei family and the bacterial family of Pseudomonas sp., as well as the Fusarium solani pisi cutinase have been devoted most protein engineering interest. See Table 2 for overview of selection of the lipases and protein engineering issues discussed in the following sections. The following topics of lipase protein engineering will be discussed: variants addressing active site, variants addressing activity (including specific activity, substrate type specificity and chain length specificity), variants addressing stability (including temperature stability, protease stability and oxidation stability), variants addressing lid function, variants addressing macroscopic substrate interaction, variants addressing calcium binding, variants addressing detergent use and surfactant compatibility, and variants addressing X-ray structure and dynamics.
Section snippets
Variants addressing active site
Variants in the cutinase type lipase from Pseudomonas mendocina focused on identifying the active site residues and changes in the nearest residues in the sequence. Using this idea the active site Ser and His were suggested, and changes in activity, especially perhydrolysis, was obtained [10]. After obtaining the structures of the first lipases the active site residues were found to be homologous to the triad of serine proteases (Fig. 5). In Humicola lanuginosa lipase and Rhizomucor miehei
Specific activity
Variants made within 6–7 residues from the active site residues in Pseudomonas mendocina cutinase (identified as mentioned above), have in assays been shown to alter the specific activity of perhydrolysis. The ratio of perhydrolysis/trioctanoin hydrolysis was increased by approximately four times [57]. Mutations made in the lipid contact zone of Humicola lanuginosa lipase have been shown to alter the specific activity as well. Normally the specific activity is decreased by mutations, but some
Thermostability
Proline residues exhibit special dihedral angles for the φ and ψ angles in the polypeptide backbone. The cyclic proline residue lowers the entropy of unfolding and thus stabilises the protein. Proline substitutions have been made in Humicola lanuginosa lipase in positions where the φ and ψ dihedral angles are appropriate for a Pro residue [77]. The temperature stability has been increased by 2°C for the mutation G225P in Humicola lanuginosa lipase measured by differential scanning calorimetry
Mutations addressing lid function
The lid(s) or flap(s) of the lipases has always attracted a big interest. Deletions or substitutions in the lid have been attempted in many lipases including Human lipoprotein lipase, Human pancreatic lipase and Humicola lanuginosa lipase, with the purpose of changing the lipase function or changing the conformation of the lid or flap. Covalently trapping of the lid in an open conformation has been attempted resulting a partly open lid, but also a decrease in activity [83]. Transfer of the flap
Mutations addressing macroscopic substrate specificity
Measuring lipase activity for the macroscopic substrate is an issue to address as well as the monosubstrate type. The surface of lipid has importance for the activation of the enzyme (see above) and for the presentation of the substrate to the enzyme. Also the surface pressure is important for the enzyme stability as well as its specificity [4]. In order to analyse binding of the lipase to the real lipid substrate, an inactive variant with only one Trp in the lid has been made. In Humicola
Mutations addressing calcium binding
The lipases from Pseudomonas species has a calcium-binding site, and by homology the Staphylococcus species was suggested to have a calcium site. Staphylococcus hyicus lipase has been made calcium independent by mutations in position 357 and 354 [101]. The mutations were made based on sequence alignment to other lipase sequences, including the Pseudomonas glumae lipase sequence, of which an X-ray structure is present showing a calcium-binding site, and further experimental data on the
Mutations addressing detergent use and surfactant compatibility
In detergents, anionic and non-ionic surfactants are the main components. The anionic compatibility of Fusarium solani pisi cutinase has been addressed by mutations in position 172, 17 and 196 based on surface charge analysis [70]. Substitution resulting in a more positively charged surface, N172K, was found to be less stable than the wild-type cutinase. Introducing negative charged residues in positions 17 and 196 (R17E and R196E), resulted in an enzyme with increased stability against the
Acknowledgements
Thanks to Shamkant Anant Patkar, Andreas Larvig Andersen, Sanne Ormholt Schrøder Glad and Stefan Minning for critical reading of the manuscript.
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