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References
[1] Arpigny JL, Jaeger K-E. Bacterial lipolytic enzymes: classification and properties.
Biochem J 1999;343:177–83.
[2] Iwasaki Y, Yamane Ts. Enzymatic synthesis of structured lipids. J Mol Catal B:
Enzym 2000;10:129–40.
[3] Chen Sh, Qian L, Shi B. Purification and properties of enantioselective
lipase from a newly isolated Bacillus cereus C71. Process Biochem 2007;42:
988–94.
[4] Moreira M, Nascimento M. Chemo-enzymatic epoxidation of (+)-3-carene.
Catal Commun 2007;8:2043–7.
[5] Mateo C, Palomo J, Fernandez-Lorente G, Guisan J, Fernandez-Lafuente R.
Improvement of enzyme activity, stability and selectivity via immobilization
techniques. Enzyme Microb Technol 2007;40:1451–63.
[6] Bora L, Kalita M. Production of thermostable alkaline lipase on vegetable
oils from a thermophilic Bacillus sp. DH4, characterization and its poten-
tial applications as detergent additive. J Chem Technol Biotechnol 2008;83:
688–93.
[7] Li J, Jiang Zh, Wu H, Liang Y, Zhang Y, Liu J. Enzyme–polysaccharide inter-
action and its influence on enzyme activity and stability. Carbohyd Polym
2010;82(1,2):160–6.
[8] Martın J, Nus M, Gago J, Sanchez-Montero J. Selective esterification of phthalic
acids in two ionic liquids at high temperatures using a thermostable lipase
of Bacillus thermocatenulatus: a comparative study. J Mol Catal B: Enzym
2008;52–53:162–7.
Fig. 8. Multiple use of the biocatalysts in alcoholysis of vinyl acetate with isoamyl
alcohol.
(ꢀ) nano-SnO2-CRL; (ꢂ) PP-CRL.
Reaction conditions: isoamyl alcohol (1.1 mL, 10 mM), vinyl acetate (0.92 mL,
10 mM), and 0.1 g immobilized lipase were mixed. The reaction mixture was stirred
at 50 ◦C for 24 h.
when nano-SnO2-CRL and PP-CRL were used as catalysts, respec-
tively. The raise of the temperature with 10 ◦C, however, resulted in
an increase in the yield with 2–7% only for 24 h. Meanwhile, the pro-
longed heating damaged the tertiary structure and had detrimental
effect on the catalytic activity of the enzyme (heat denaturation).
[9] Ulbert O, Belafi-Bako K, Tonova K, Gubicza L. Thermal stability enhance-
ment of Candida rugosa lipase using ionic liquids. Biocatal Biotransfor
2005;23(3–4):177–83.
[10] Hernandez-Fernandez F, de los Rios A, Tomas-Alonso F, Gomez D, Villora G. Sta-
bility of hydrolase enzymes in ionic liquids. Can J Chem Eng 2009;87(6):910–4.
[11] Gao S, Wang Y, Diao X, Luo G, Dai Y. Effect of pore diameter and cross-linking
method on the immobilization efficiency of Candida rugosa lipase in SBA-15.
Bioresource Technol 2010;101:3830–7.
[12] Shakeri M, Kawakami K. Effect of the structural chemical composition of meso-
porous materials on the adsorption and activation of the Rhizopus oryzae
lipase-catalyzed trans-esterification reaction in organic solvent. Catal Commun
2008;10:165–8.
[13] Reis P, Holmberg K, Watzke H, Leser M, Miller R. Lipases at interfaces: a review.
Adv Colloid Interface Sci 2009;14:7–148, 237–250.
[14] Dyal A, Loos K, Noto M, Chang S, Spagnoli Ch, Shafi K, Ulman A, Cowman M,
Gross R. Activity of Candida rugosa lipase immobilized on (␥Fe2O3 magnetic
nanoparticles. J Am Chem Soc 2003;125:1684–5.
[15] Tang Zh-X, Qian J-Q, Shi Lu-E. Characterizations of immobilized neutral lipase
on chitosan nano-particles. Mater Lett 2007;61:37–40.
[16] Lei L, Bai L, Li Y, Yi L, Yang Y, Xia Ch. Study on immobilization of lipase
onto magnetic microspheres with epoxy groups. J Magn Mater 2009;321:
252–8.
[17] Chen YZ, Yang CT, Ching CB, Xu R. Immobilization of lipases on hydrophobi-
lized zirconia nanoparticles: highly enantioselective and reusable biocatalysts.
Langmuir 2008;24(16):8877–84.
[18] Dimitrov M, Tsoncheva T, Shao Sh, Kohn R. Novel preparation of nanosized
mesoporous SnO2 powders: physicochemical and catalytic properties. Appl
Catal B: Environ 2010;94:158–65.
3.5.4. Multiple use of nano-SnO2-CRL and PP-CRL
The repeated use of enzymes is a major concern for large-scale
application. We assessed the ability of nano-SnO2-CRL and PP-CRL
to catalyze several consecutive 24 h-cycles of alcoholysis of vinyl
acetate with isoamyl alcohol at 50 ◦C (Fig. 8). Both preparations
showed a sharp decline in activity after the second run. Kanwar et al.
reported similar behaviour after the 4th cycle (46%) for hydrogel-
bounded B. coagulans lipase applied in the same reaction [39]. The
limited reusability of both biocatalysts is probably due to an inhi-
bition by substrates or products (mainly by isoamyl alcohol), long
reaction time and prolonged heating.
4. Conclusion
To our knowledge this is the first report on the application
of tin dioxide as enzyme carrier. A lipase from C. rugosa was
successfully immobilized on the novel nanostructured material
(nano-SnO2-CRL). Probably due to the large size of CRL molecules
(5 nm × 4.2 nm × 3.3 nm) (pdb 1TRH), the enzyme is distributed
mainly on the surface of the carrier which results in low loading
capacity of the tin dioxide. Further pore size optimization or sur-
face modification of the material is required in order to increase the
loading capacity of the tin dioxide. The inorganic material stabilizes
the protein, and nano-SnO2-CRL has shown a significant specific
activity, thermal- and pH-stability. The novel catalyst was success-
fully applied in the isoamyl acetate production (banana flavor ester)
and the results are comparable with the best results found in the
literature. Nano-SnO2-CRL was significantly tolerant toward the
reaction medium and can be applied in synthetic reactions in pres-
ence of organic solvents. The results obtained here show the good
potential of nanostructured tin dioxide as a carrier for enzymes that
should be explored circumstantially. Further investigations includ-
ing variations in tin dioxide pore characteristics and/or the size of
the lipase molecules are in progress.
[19] Guncheva M, Zhiryakova D. High-yield synthesis of wax esters catalysed by
modified Candida rugosa lipase. Biotechnol Lett 2008;30:509–12.
[20] Lowry O, Rosebrough N, Farr A, Randall R. Protein measurement with the folin
phenol reagent. J Biol Chem 1951;193:265–75.
[21] Kwon D, Rhee J. A simple and rapid colorimetric method for determination of
free fatty acids for lipase assay. J Am Oil Chem Soc 1986;63(1):89–92.
[22] Halling P. Salt hydrates for water activity control with biocatalysts in organic
media. Biotechnol Tech 1992;6(3):271–6.
[23] Hammond C. The Basics of crystallography and diffraction. New York: Oxford
Science Publications; 2001. p. 180–3 [chapter 9].
[24] Sing K, Everett D, Haul R, Moscou L, Pierotti R, Rouquerol J, Siemieniewska T.
Pure Appl Chem 1985;57:603–19.
[25] Reis P, Holmberg K, Watzke H, Leser M, Miller R. Lipases at interfaces: a review.
Adv Colloid Interface Sci 2009;147–148:237–50.
[26] Mingarro I, Abad C, Braco L. Interfacial activation based molecular bioimprint-
ing of lipolytic enzymes. Proc Natl Acad Sci USA 1995;92:3308–12.
[27] Lewis J. Colloidal processing of ceramics. J Am Ceram Soc 2000;83(10):2341–59.
[28] Chong A, Zhao X. Functionalized nanoporous silicas for the immobilization of
penicillin acylase. Appl Surf Sci 2004;237:398–404.
[29] Bosley J, Peilow A. Immobilization of lipases on porous polypropylene:
reduction in esterification efficiency at low loading.
1997;74(2):107–11.
J Am Oil Chem Soc
[30] Orrego C, Valencia J, Zapata C. Candida rugosa lipase supported on high crys-
tallinity chitosan as biocatalyst for the synthesis of 1-butyl oleate. Catal Lett
2009;129:312–22.
[31] Persson M, Wehje E, Adlercreutz A. Immobilisation of lipases by adsorption and
deposition: high protein loading gives lower water activity optimum. Biotech-
nol Lett 2000;22:1571–7.
[32] Othman S, Basri M, Hussein M, Rahman M, Rahman R, Salleh A, Jasmani
H. Production of highly enantioselective (−)-methyl butyrate using Candida
rugosa lipase immobilized on epoxy-activated supports. Food Chem 2008;43:
7–443.
Acknowledgement
The authors thank the National Science Fund of Bulgaria (project
DMU 02/20) for the financial support.