- Research article
- Open Access
Enhancement of stability of a lipase by subjecting to three phase partitioning (TPP): structures of native and TPP-treated lipase from Thermomyces lanuginosa
© Kumar et al. 2015
- Received: 3 July 2015
- Accepted: 15 September 2015
- Published: 24 September 2015
The lipase enzyme converts long chain acyltriglycerides into di- and monoglycerides, glycerol and fatty acids. The catalytic site in lipase is situated deep inside the molecule. It is connected through a tunnel to the surface of the molecule. In the unbound state under aqueous conditions, the tunnel remains closed. The tunnel can be opened when the enzyme is exposed to a lipid bilayer or a detergent or many hydrophobic/hydrophilic surfaces.
In the present study, the lipase was subjected to three-phase partitioning (TPP) which consisted of mixing in tert-butanol and ammonium sulphate to the solution of lipase in the aqueous buffer. The enzyme formed an interfacial precipitate between the tert-butanol rich and water rich phases. The stability of the enzyme subjected to TPP was found to be higher (Tm of 80 °C) than the untreated enzyme (Tm of 77 °C). The activity of the enzyme subjected to TPP (3.3 U/mg) was nearly half of that of the untreated one (5.8 U/mg). However, the activity of the treated enzyme was higher (17.8 U/mg) than the untreated one (8.6 U/mg) when a detergent was incorporated in the assay buffer.
The structure determination showed that the substrate binding site in the treated enzyme was more tightly closed than that of the untreated protein.
- Crystal structures
- Three-phase partitioning
- New binding site
Enzymes offer a more sustainable option for catalyzing chemical processes . Hence, enhancing their stability and activity is a worthwhile goal. The present work describes efforts to understand how a simple process called three phase partitioning (TPP) resulted in altering both stability and activity of a lipase from Thermomyces lanuginosus (TLL). Lipases (triacylglycerol hydrolases; EC 220.127.116.11) have been extensively used in both aqueous media as well as non aqueous media [1–8]. One of the main reasons of their large-scale usage has been their broad specificity [8, 9]. Lately, this range has been further enlarged by the reports on the lipase catalysed promiscuous reactions [10–13]. The catalytic promiscuity refers to enzymes belonging to a particular class in the enzyme classification (EC) system catalyzing reactions of the type which are generally catalyzed by another class of enzymes. Lipases, classified as hydrolases, have been shown to catalyze many C–C bond formation reactions [10, 11, 14, 15]. It is believed that in such cases, substrates interact with the active site in a manner different from that observed by natural substrates .
The lipase from Thermomyces lanuginosus (TLL) is a glycosylated hydrolase which consists of 269 amino acid residues with a molecular weight of about 29 kDa. It has an optimum pH of 11–12 . The structure of TLL was reported earlier in the native state which showed that the substrate binding cleft was in the closed form . The structure determined in the presence of a detergent showed open conformation . TLL is among the most frequently used lipases in biotechnology  and its applications include its use in catalysing C–C bond formation because of its promiscuous activity .
In the present work, we report the protein structures based upon X-ray crystallographic studies of TLL including (1) native untreated form (TLL), (2) TLL after being subjected to three-phase partitioning (TPP-TLL). These structures are discussed alongwith the results from the biochemical studies. It was found that TPP of TLL, contrary to what was expected, resulted in the introduction of rigidity around the active site.
Activity of TLL and TPP-TLL
Three phase partitioning of Thermomyces lanuginosus lipase (TLL) at 25 °C
Specific activity (U/mg protein)
Assay in the absence of Triton X-100
Assay in the presence of Triton X-100
The distances (Å) between atoms of residues from opposite sides of the substrate binding cleft indicating the very closed, closed and open states of substrate binding channel
TPP-TLL (PDB id: 4FLF)
TLL (PDB id:4ZGB)
Detergent-treated (PDB id: 1GT6)
86 ILE CD1
87 GLU CG
95 PHE O
However, when assayed in the presence of detergent triton X-100, the TPP-TLL showed an increase in the specific activity from 3.3 U/mg protein to 17.8 U/mg protein (Table 1). On the other hand, the native TLL showed an increase in the specific activity from 5.8 U/mg proteins to 8.6 U/mg proteins. This shows that the presence of detergent has a more marked effect on the specific activity of the TPP-TLL. In this case, the opening of the lid not only resulted in the more active “open” conformation (which is a well known phenomenon) but also presumably reversed the introduction of more rigidity in the active site by TPP treatment.
The lipase-detergent interactions have been exploited in multiple ways . The most well known among these is via “interfacial activation”. This leads to the movement of the molecular lid (present in several lipases including TLL) away from the active site access and makes the active site more accessible. This “lid opening” has also been achieved by immobilization [37, 38] and bioimprinting [39, 40].
Structure of the untreated native lipase
Structure of TLL subjected to TPP
Crystal structure of TPP-TLL was determined at 2.15 Å resolution. The overall structure treated form is similar to that of untreated native protein with an r.m.s. shift of 0.7 Å for the Cα atoms. The exposure to the interface between the aqueous layer and the upper tert-butanol rich layer was expected to cause a conformational change in the cleft. However, the structure did not indicate the opening of the lid. This is easy to understand since it is well known that interfacial activation of the lipases requires interaction of the lipase in free solution with an interface [43, 44]. In TPP treatment, the enzyme precipitates out of the solution and cannot interact with the tert-butanol rich layer (unlike the enzyme in free solution). In fact, rather than opening the lid, the TPP treatment resulted in making the active site less accessible due to enhanced molecular rigidity in that region. As a result, the lid further moved closer to the segment on the opposite side thus shortening the lengths of van der Waals contacts (Table 2). The TPP treatment, involving complex interactions of the SO4 2− anions (kosmotropy, conformation tightening and electrostatic forces) simultaneously with interactions with tert-butanol induces a conformational change. Such conformational changes upon TPP treatment have been reported [22, 27, 28, 30, 45]. In the present structure of the TPP-TLL, as seen from Fig. 4c, the two walls moved closer to each other thus optimizing the van der Waals contacts between the two sides of the substrate binding cleft. In this case, the distances of hydrophobic contacts decreased at least by 10 %. This suggests that after the TPP treatment, the protein structure becomes more tight and hence more stable than the native state as observed in untreated TLL.
Enhancing the thermal stability of enzymes continues to be a useful goal in biocatalysis [46–48]. Many techniques including chemical crosslinking [49–51], immobilization on solid or soluble supports [52–54], protein engineering and directed evolution [55–58] have been employed for this purpose. In the present case, however, it was an incidental result of these studies wherein the aim was to improve both the natural and promiscuous activities of the lipase by subjecting it to TPP as seen with other enzymes [22, 23, 27–30, 45]. This is the first example where the TPP treatment seems to have resulted in the increase in the Tm. In cases reported so far, TPP treatment has resulted in a decrease of thermal stability . The increase in the Tm of TLL upon TPP-treatment correlates well with the reduction in the activity. In this regard, it is noteworthy that the substrate binding site became more tight because the distances of the van der Waals contacts between the two sides of the substrate binding channel became considerably shorter than those observed in the untreated protein. This would have made it more difficult for the cleft to open when the substrate approached it. However, in the presence of detergent, the TLL subjected to TPP showed a higher activity because of the more favourable stereochemistry of the active site residues in the treated enzyme. The TLL subjected to TPP was less active and more stable which is good for its shelf life. On the other hand, in the presence of detergent, the TLL subjected to TPP showed activity more than two times higher than the untreated enzyme. An interesting aspect is that the TLL subjected to TPP in the absence of a detergent showed a decrease in the hydrolytic activity which involves hydrolysis of an analogue of a natural substrate, that is, p-nitrophenylpalmitate. However, it shows higher activity during catalysis of the promiscuous reaction, that is, aldol condensation. Broos  has provided convincing data which shows that changes in flexibility of the enzyme structure may have opposite consequences in terms of selectivity for natural and unnatural substrates. The promiscuous substrates are extreme examples of “unnatural substrates”.
It is noteworthy that the TLL subjected to TPP showed 5-fold increase in the initial rates for the aldol condensation (in the presence of 40 % water in acetone as the medium which was also one of the substrates). This reaction was carried out in the absence of detergents, so this involved ‘closed lid’ structure of the enzyme. Also, one should not overlook the fact that one of the substrates in the promiscuous reaction was acetone. The presence of acetone in the medium was expected to influence the structure of the enzyme. The information about the change or its extent could not unfortunately be obtained by X-ray studies. The efforts to obtain the crystallization of TLL in the presence of acetone did not succeed.
The lipase from Thermomyces lanuginosus was a kind gift from Novozyme A/S (Bagsvaerd, Denmark). p-Nitrophenyl palmitate (pNPP) was procured from Sigma-Aldrich Company (St. Louis, USA). All other chemicals used were of analytical grade.
Three phase partitioning (TPP) of lipases
The protocol reported earlier was followed for carrying out the TPP treatment of TLL . The solutions of TLL (2 mL, 20 mg/mL in 10 mM sodium phosphate buffer, pH 7.0 were saturated with varying concentrations of ammonium sulphate (w/v). This step was followed by the addition of 2 mL tert-butanol. The solutions were vortexed and allowed to stand at 25 °C for 1 h. Three phases (i.e., upper layer of tert-butanol, interfacial precipitate of protein and lower aqueous layer) were formed. The solutions were then centrifuged at 2000×g at 25 °C for 10 min. The lower aqueous and upper organic layer were separated using a pasteur pipette. The interfacial precipitates obtained were dissolved in 1 mL of 10 mM sodium phosphate buffer pH 7.0 and dialysed against the same buffer for 24 h with frequent changes of buffer. This was checked for lipase activity using 4-nitrophenyl palmitate (pNPP) as a substrate  and for amounts of protein using Bradford method . The percentage activity and protein in the precipitates were calculated by taking the starting amount of activity and protein as 100 %. This preparation is referred to as TPP-TLL.
Assay for lipase activity
The hydrolytic activity of lipase was monitored by measuring the rate of hydrolysis of 4-nitrophenyl palmitate (pNPP) spectrophotometrically at 410 nm by following the procedure described earlier . The reaction mixtures consisted of 1.8 mL of buffer (10 mM sodium phosphate, pH 7.0 containing 150 mM NaCl [and with or without 0.5 % (v/v) triton X-100]), 0.2 mL of enzyme solution in 10 mM sodium phosphate buffer, pH 7.0 and 20 µl of 50 mM substrate pNPP in acetonitrile. The mixtures were incubated for 30 min at 37 °C and after this the samples were kept in a domestic microwave oven along with a beaker containing a volume of water sufficient to make the total volume of the liquid in the cavity as 100 mL (the additional water absorbs a significant amount of microwave energy, this was done to avoid overheating of the sample) and irradiated for 30 s and read at 410 nm. One unit (U) of enzyme activity is defined as the amount of the enzyme that liberates 1 µmol of 4-nitrophenol per min at pH 7.0 and 37 °C.
Protein concentrations were determined according to the procedure described by Bradford . Protein solutions (0.5 mL) were incubated with 4.5 mL of the dye reagent at 25 °C for 10 min and the absorbance of the solutions were read at 595 nm and bovine serum albumin was used as the standard protein.
Measurement of the melting temperature (Tm)
The melting temperature of TLL and TPP-TLL were determined by using circular dichroism studies. Thermal denaturation curves were determined directly by monitoring the ellipticity changes at 222 nm. The samples with a concentration of 0.2 mg mL−1 were used. The temperature of sample solution was raised linearly by 1 °C min−1 from 50 to 100 °C. The heating curves were corrected for an instrumental baseline obtained by heating the buffer (10 mM sodium phosphate, pH 7.0) alone. The melting temperature (Tm) was calculated from the first-order derivatives of the ellipticity-temperature plot.
The samples of both (1) non-treated lipase (TLL) and (2) the lipase subjected to three phase partitioning (TPP-TLL) were dissolved in solution containing 0.1 M HEPES buffer, 1 M NaCl pH 7.5 at a concentration of 30 mg/mL to make solutions (1) and (2), respectively. The drops of 10 µl of solutions (1) and (2) were prepared for the hanging drop vapour diffusion method. The protein drops were equilibrated against 1.6 M ammonium sulphate. The rectangular shaped crystals measuring up to dimensions of 0.4 × 0.3 × 0.3 mm3 were obtained after 3 weeks from the drops of the solutions. The crystals from the two samples were washed with reservoir buffer before placing them into a fresh solution containing reservoir buffer and 25 % glycerol as a cryoprotectant for data collection at low temperatures.
Data collection and processing
Crystallographic data collection and refinement statistics
Resolution range (Å)
Unit cell parameters
a = b (Å)
Number of molecules in the asymmetric unit
Total number of measured reflections
Number of unique reflections
R.m.s.d in bond lengths (Å)
R.m.s.d in bond angles (°)
R.m.s.d in torsion angles (°)
Ramachandran plot analysis
Most favoured (%)
Additionally allowed (%)
Generously allowed region (%)
Disallowed region (%)
B factors (Å2)
Wilson B factor
Average B-factor for main chain atoms
Average B-factor for the side chain atoms and water oxygen atoms
Mean B-factor for all atoms
Number of protein atoms
Number of water oxygen atoms
Number of carbohydrate (NAG) residues
Structure determinations and refinements
The structure of TLL was determined with molecular replacement method  using the coordinates of lipase in its unbound state (PDB code: 1DT3) . This produced a clear solution with two peaks. Initially, the structure was refined for 20 cycles using REFMAC 5.5 . The coordinates of partially refined structure of TLL were used as the starting model for refining the TPP-TLL structure. The initial calculations for 25 cycles were carried out as the rigid body refinement. The electron density maps with (2Fo–Fc) and (Fo–Fc) coefficients were calculated. The models were improved by manual model building with programs O  and Coot  using graphics workstations. After another 25 cycles of refinements, when the values of Rcryst/Rfree factors dropped to less than 0.27/0.30, Fourier maps with (2Fo–Fc) and (Fo–Fc) coefficients were calculated for both the structures. These maps were used for determining the positions of water oxygen atoms in both structures. The coordinates of 245 water oxygen atoms were obtained in the TLL structure while the coordinates of 288 water oxygen were determined in the TPP-TLL structure. These coordinates were included in the final cycles of refinements. The refinements finally converged to values of 0.221/0.270 and 0.227/0.268 for the Rcryst/Rfree factors of the untreated and treated structures. The final refinement statistics are included in Table 3. The refined atomic coordinates of structures of TLL and TPP-TLL have been deposited in the protein data bank with accession codes of 4ZGB and 4FLF respectively.
In the case of TLL, TPP treatment introduces a rigidity rather than flexibility around the active site region.
Upon opening up the lid by interfacial activation, TLL subjected to TPP assumes a structure which was more along the expected lines. As in the case of proteinase K , it might be more flexible in the state with an open lid than the native structure. This might have led to the significant increase in the enzyme activity.
Three phase partitioning has slowly emerged as a simple approach to subtly alter conformational flexibility. Even subtle changes in conformational flexibility are important for both natural and promiscuous reactions of enzymes and influence even their enantioselectivity .
The present work shows that TLL subjected to TPP behaves differently while it exists as a “closed lid” structure and as an “open lid” structure. In the “closed lid structure”, TLL subjected to TPP has enhanced rigidity while the opening of the lid by well known interfacial activation removes this rigidity. The results are also useful in the context of understanding the role which conformational flexibility plays in catalytic promiscuity.
The group at AIIMS (MK, MS, PK, SS and TPS) were involved in the crystallization of the enzyme and its complexes, interpretation of the X-ray data and discussion with the IITD group. JM carried out the experimental work consisting the activity assay, Tm measurements etc. MNG suggested probing TLL activity with or without TPP treatment and correlating the results of the X-ray studies with the biochemical implications. All authors read and approved the final manuscript.
This work was supported by Grants from the Department of Science and Technology (DST-SERB), Govt. of India (Grant No. SR/SO/BB-68/2010) to the IITD. The AIIMS group also thanks Department of Science and Technology (DST), New Delhi and Department of Biotechnology (DBT), New Delhi for financial support. TPS also thanks the Indian National Science Academy for the grant of position of INSA-Senior Scientist. JM thanks CSIR (Govt. of India) for the senior research fellowship.
Compliance with ethical guidelines
Competing interests The authors declare that they have no competing interests.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Fessner WD, Anthonsen T (2008) Modern biocatalysis: stereoselective and environmentally friendly reactions. Wiley, New YorkView ArticleGoogle Scholar
- Gupta MN (1992) Enzyme function in organic solvents. Eur J Biochem 203:25–32View ArticleGoogle Scholar
- Carrea G, Riva S (2000) Properties and synthetic applications of enzymes in organic solvents. Angew Chem Int Ed Engl 39:2226–2254View ArticleGoogle Scholar
- Gupta MN (ed) (2000) Methods in non-aqueous enzymology. Birkhäuser-Verlag, BaselGoogle Scholar
- Halling PJ (2000) Biocatalysis in low-water media: understanding effects of reaction conditions. Curr Opin Chem Biol 4:74–80View ArticleGoogle Scholar
- Hudson EP, Eppler RK, Clark DS (2005) Biocatalysis in semi-aqueous and nearly anhydrous conditions. Curr Opin Biotechnol 16:637–643View ArticleGoogle Scholar
- Mattiasson B, Adlercreutz P (1991) Tailoring the microenvironment of enzymes in water-poor systems. Trends Biotechnol 9:394–398View ArticleGoogle Scholar
- Adlercreutz P (2013) Immobilisation and application of lipases in organic media. Chem Soc Rev 42:6406–6436View ArticleGoogle Scholar
- Kapoor M, Gupta MN (2012) Lipase promiscuity and its biochemical applications. Process Biochem 47:555–569View ArticleGoogle Scholar
- Khersonsky O, Tawfik DS (2010) Enzyme promiscuity: a mechanistic and evolutionary perspective. Annu Rev Biochem 79:471–505View ArticleGoogle Scholar
- Busto E, Gotor-Fernández V, Gotor V (2010) Hydrolases: catalytically promiscuous enzymes for non-conventional reactions in organic synthesis. Chem Soc Rev 39:4504–4523View ArticleGoogle Scholar
- Arora B, Pandey PS, Gupta MN (2014) Lipase catalyzed Cannizzaro-type reaction with substituted benzaldehydes in water. Tetrahedron Lett 55:3920–3922View ArticleGoogle Scholar
- Malhotra D, Mukherjee J, Gupta MN (2015) Sustainability of biocatalytic processes. In: Letcher T, Scott JL, Patterson DA (eds) Chemical processes for a sustainable future. Royal Society of Chemistry, CambridgeGoogle Scholar
- Hult K, Berglund P (2007) Enzyme promiscuity: mechanism and applications. Trends Biotechnol 25:231–238View ArticleGoogle Scholar
- Arora B, Mukherjee J, Gupta MN (2014) Enzyme promiscuity: using the dark side of enzyme specificity in white biotechnology. Sustain Chem Process 2:25View ArticleGoogle Scholar
- Li N, Zong M-H, Ma D (2009) Unexpected reversal of the regioselectivity in Thermomyces lanuginosus lipase-catalyzed acylation of floxuridine. Biotechnol Lett 31:1241–1244View ArticleGoogle Scholar
- Derewenda U, Swenson L, Wei Y, Green R, Kobos PM, Joerger R, Haas MJ, Derewenda ZS (1994) Conformational lability of lipases observed in the absence of an oil-water interface: crystallographic studies of enzymes from the fungi Humicola lanuginosa and Rhizopus delemar. J Lipid Res 35:524–534Google Scholar
- Brzozowski AM, Savage H, Verma CS, Turkenburg JP, Lawson DM, Svendsen A, Patkar S (2000) Structural origins of the interfacial activation in Thermomyces (Humicola) lanuginosa lipase. Biochemistry 39:15071–15082View ArticleGoogle Scholar
- Fernandez-Lafuente R (2010) Lipase from Thermomyces lanuginosus: uses and prospects as an industrial biocatalyst. J Mol Catal B 62:197–212View ArticleGoogle Scholar
- Cai JF, Guan Z, He YH (2011) The lipase-catalyzed asymmetric C–C Michael addition. J Mol Catal B Enzym 68:240–244View ArticleGoogle Scholar
- Lovrein RE, Goldensoph C, Anderson P, Odegard B (1987) Three phase partitioning (TPP) via t-butanol: enzyme separation from crudes. In: Burgess R (ed) Protein purification, micro to macro. Marcel Dekker Inc., New YorkGoogle Scholar
- Dennison C, Lovrien R (1997) Three phase partitioning: concentration and purification of proteins. Protein Expr Purif 11:149–161View ArticleGoogle Scholar
- Roy I, Gupta MN (2004) α-Chymotrypsin shows higher activity in water as well as organic solvents after three phase partitioning. Biocatal Biotransform 22:261–268View ArticleGoogle Scholar
- Narayan AV, Madhusudhan MC, Raghavarao KS (2008) Extraction and purification of Ipomoea peroxidase employing three-phase partitioning. Appl Biochem Biotechnol 151:263–272View ArticleGoogle Scholar
- Roy I, Sharma A, Gupta MN (2004) Obtaining higher transesterification rates with subtilisin Carlsberg in nonaqueous media. Bioorg Med Chem Lett 14:887–889View ArticleGoogle Scholar
- Raghava S, Barua B, Singh PK, Das M, Madan L, Bhattacharyya S, Bajaj K, Gopal B, Varadarajan R, Gupta MN (2008) Refolding and simultaneous purification by three-phase partitioning of recombinant proteins from inclusion bodies. Protein Sci 17:1987–1997View ArticleGoogle Scholar
- Lovrein RE, Conroy MJ, Richardson TI, Gregory RB (1995) Protein solvent interactions. Marcel Dekker Inc., New YorkGoogle Scholar
- Singh RK, Gourinath S, Sharma S, Roy I, Gupta MN, Betzel C, Srinivasan A, Singh TP (2001) Enhancement of enzyme activity through three-phase partitioning: crystal structure of a modified serine proteinase at 1.5 A resolution. Protein Eng 14:307–313View ArticleGoogle Scholar
- Rather GM, Mukherjee J, Halling PJ, Gupta MN (2012) Activation of alpha chymotrypsin by three phase partitioning is accompanied by aggregation. PLoS One 7:e49241View ArticleGoogle Scholar
- Rather GM, Gupta MN (2013) Three phase partitioning leads to subtle structural changes in proteins. Int J Biol Macromol 60:134–140View ArticleGoogle Scholar
- Rather GM, Gupta MN (2013) Refolding of urea denatured ovalbumin with three phase partitioning generates many conformational variants. Int J Biol Macromol 60:301–308View ArticleGoogle Scholar
- Singh N, Jabeen T, Sharma S, Roy I, Gupta MN, Bilgrami S, Somvanshi RK, Dey S, Perbandt M, Betzel C, Srinivasan A, Singh TP (2005) Detection of native peptides as potent inhibitors of enzymes. Crystal structure of the complex formed between treated bovine alpha-chymotrypsin and an autocatalytically produced fragment, IIe-Val-Asn-Gly-Glu-Glu-Ala-Val-Pro-Gly-Ser-Trp-Pro-Trp, at 2.2 angstroms resolution. FEBS J 272:562–572View ArticleGoogle Scholar
- Mukherjee J, Mishra P, Gupta MN (2015) Urea treated subtilisin as a biocatalyst for transformations in organic solvents. Tetrahedron Lett 56:1976–1981View ArticleGoogle Scholar
- Jain P, Jain S, Gupta MN (2005) A microwave-assisted microassay for lipases. Anal Bioanal Chem 381:1480–1482View ArticleGoogle Scholar
- Mogensen JE, Sehgal P, Otzen DE (2005) Activation, inhibition, and destabilization of Thermomyces lanuginosus lipase by detergents. Biochemistry 44:1719–1730View ArticleGoogle Scholar
- Shah S, Gupta MN (2007) Obtaining high transesterification activity for subtilisin in ionic liquids. Biochim Biophys Acta 1770:94–98View ArticleGoogle Scholar
- Bastida A, Sabuquillo P, Armisen P, Fernández-Lafuente R, Huguet J, Guisán JM (1998) A single step purification, immobilization, and hyperactivation of lipases via interfacial adsorption on strongly hydrophobic supports. Biotechnol Bioeng 58:486–493View ArticleGoogle Scholar
- Fernandez-Lafuente R, Armisén P, Sabuquillo P, Fernández-Lorente G, Guisán JM (1998) Immobilization of lipases by selective adsorption on hydrophobic supports. Chem Phys Lipids 93:185–197View ArticleGoogle Scholar
- Yilmaz E (2002) Improving the application of microbial lipase by bio-imprinting at substrate-interfaces. World J Microbiol Biotechnol 18:37–40View ArticleGoogle Scholar
- Fishman A, Cogan U (2003) Bio-imprinting of lipases with fatty acids. J Mol Catal B 22:193–202View ArticleGoogle Scholar
- Yapoudjian S, Ivanova MG, Brzozowski AM, Patkar SA, Vind J, Svendsen A, Verger R (2002) Binding of Thermomyces (Humicola) lanuginosa lipase to the mixed micelles of cis-parinaric acid/NaTDC. Eur J Biochem 269:1613–1621View ArticleGoogle Scholar
- de Diego T, Lozano P, Gmouh S, Vaultier M, Iborra JL (2005) Understanding structure-stability relationships of Candida antartica lipase B in ionic liquids. Biomacromolecules 6:1457–1464View ArticleGoogle Scholar
- Desnuelle P (1961) Pancreatic lipase. Adv Enzymol 23:129–161Google Scholar
- Brockman HL, Law JH, Kézdy FJ (1973) Catalysis by adsorbed enzymes. The hydrolysis of tripropionin by pancreatic lipase adsorbed to siliconized glass beads. J Biol Chem 248:4965–4970Google Scholar
- Sharma A, Roy I, Gupta MN (2004) Affinity precipitation and macroaffinity ligand facilitated three-phase partitioning for refolding and simultaneous purification of urea-denatured pectinase. Biotechnol Prog 20:1255–1258View ArticleGoogle Scholar
- Santoro MM, Liu Y, Khan SM, Hou LX, Bolen DW (1992) Increased thermal stability of proteins in the presence of naturally occurring osmolytes. Biochemistry 31:5278–5283View ArticleGoogle Scholar
- Gupta MN (ed) (1993) Thermostability of enzymes. Springer, HeidelbergGoogle Scholar
- Kuznetsova IM, Turoverov KK, Uversky VN (2014) What macromolecular crowding can do to a protein. Int J Mol Sci 15:23090–23140View ArticleGoogle Scholar
- Khare SK, Gupta MN (1988) A preparation of E. coli beta galactosidase. Appl Biochem Biotechnol 16:1–15View ArticleGoogle Scholar
- Tyagi R, Gupta MN (1998) Chemical modification and chemical crosslinking of protein/enzyme stabilization. Biochem (Moscow) 63:334–344Google Scholar
- Yamazaki T, Tsugawa W, Sode K (1999) Increased thermal stability of glucose dehydrogenase by crosslinking chemical modification. Biotechnol Lett 21:199–202View ArticleGoogle Scholar
- Cao L (2005) Carrier bound immobilized enzymes: principles, application and design. Wiley-VCH Verlag GmbH and Co., WeinheimView ArticleGoogle Scholar
- Minteer SD (ed) (2011) Enzyme stabilization and immobilization: methods and protocols. Humana Press, New YorkGoogle Scholar
- Guisan JM (ed) (2013) Immobilization of enzymes and cells. Humana Press, New YorkGoogle Scholar
- Carey PR (ed) (1996) Protein engineering and design. Academic, New YorkGoogle Scholar
- Nosoh Y, Sekiguchi T (1993) Protein engineering for thermostabilization. In: Gupta MN (ed) Thermostabilization of enzymes. Springer, Heidelberg, pp 182–204Google Scholar
- Arnold FH, Georgiou G (2003) Directed enzyme evolution: screening and selection methods. Humana Press, TotowaView ArticleGoogle Scholar
- Arnold FH, Georgiou G (2003) Directed evolution library construction: methods and protocol. Humana Press, TotowaView ArticleGoogle Scholar
- Broos J (2002) Impact of the enzyme flexibility on the enzyme enantio-selectivity in organic media towards specific and non-specific substrates. Biocatal Biotransform 20:291–295View ArticleGoogle Scholar
- Mondal K, Roy I, Gupta MN (2006) Affinity-based strategies for protein purification. Anal Chem 78:3499–3504View ArticleGoogle Scholar
- Bradford MM (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72:248–254View ArticleGoogle Scholar
- Otwinowski Z, Minor W (1997) Denzo and scalepack. Meth Enzymol 276:307–326View ArticleGoogle Scholar
- Vagin A, Teplyakov A (2010) Molecular replacement with MOLREP. Acta Crystallogr D Biol Crystallogr 66:22–25View ArticleGoogle Scholar
- Murshudov GN, Skubák P, Lebedev AA, Pannu NS, Steiner RA, Nicholls RA, Winn MD, Long F, Vagin AA (2011) REFMAC5 for the refinement of macromolecular crystal structures. Acta Crystallogr D Biol Crystallogr 67:355–367View ArticleGoogle Scholar
- Jones TA, Zou JY, Cowan SW, Kjeldgaard M (1991) Improved methods for building protein models in electron density maps and the location of errors in these models. Acta Crystallogr A 47:110–119View ArticleGoogle Scholar
- Emsley P, Cowtan K (2004) Coot: model-building tools for molecular graphics. Acta Crystallogr D Biol Crystallogr 60:2126–2132View ArticleGoogle Scholar
- Mukherjee J, Gupta MN (2015) Increasing importance of protein flexibility in designing biocatalytic processes. Biotechnol Rep 6:119–123View ArticleGoogle Scholar
- DeLano WL (2005) The case for open-source software in drug discovery. Drug Discov Today 10:213–217View ArticleGoogle Scholar