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Applied Biochemistry and Biotechnology

, Volume 189, Issue 3, pp 903–918 | Cite as

Screening and Immobilization of Interfacial Esterases from Marine Invertebrates as Promising Biocatalyst Derivatives

  • Alberto del Monte-MartínezEmail author
  • Jorge González-Bacerio
  • Carlos M. Varela
  • Fernando Vega-Villasante
  • Rogelio Lalana-Rueda
  • Héctor Nolasco
  • Joaquín Díaz
  • José M. Guisán
Article

Abstract

Interfacial esterases are useful enzymes in bioconversion and racemic mixture resolution processes. Marine invertebrates are few explored potential sources of these proteins. In this work, aqueous extracts of 41 species of marine invertebrates were screened for esterase, lipase, and phospholipase A activities, being all positive. Five extracts (Stichodactyla helianthus, Condylactis gigantea, Stylocheilus longicauda, Zoanthus pulchellus, and Plexaura homomalla) were selected for their activity values and immobilized on Octyl-Sepharose CL 4B support by interfacial adsorption. The selectivity of this immobilization method for interfacial esterases was evidenced by immobilization percentages ≥ 94% in almost all cases for lipase and phospholipase A activities. Six pharmaceutical-relevant esters (phenylethyl butyrate, ethyl-2-hydroxy-4-phenyl-butanoate, 2-oxyranylmethyl acetate (glycidol acetate), 7-aminocephalosporanic acid, methyl-prostaglandin F, and methyl-6-metoxy-α-methyl-2-naphtalen-acetate -naproxen methyl ester-) were bioconverted by at least three of these biocatalysts, with the lowest conversion percentage of 24%. In addition, three biocatalysts were used in the racemic mixture resolution of three previous compounds. The S. helianthus–derived biocatalyst showed the highest enantiomeric ratios for glycidol acetate (2.67, (S)-selective) and naproxen methyl ester (8.32, (R)-selective), and the immobilized extract of S. longicauda was the most resolutive toward the ethyl-2-hydroxy-4-phenyl-butanoate (8.13, (S)-selective). These results indicate the relevance of such marine interfacial esterases as immobilized biocatalysts for the pharmaceutical industry.

Keywords

Enzymatic bioconversion Interfacial adsorption Interfacial esterases Marine invertebrates Pharmaceutical-relevant esters Racemic mixture resolution 

Notes

Acknowledgments

We are grateful to ICP-CSIC, Madrid, Spain, and CQF-MINSAP, La Habana, Cuba, for supplying several model compounds used in this analysis.

Funding Information

The authors wish to thank INFORMATICA ddmm., Bergamo, Italy, for kindly providing financial support to A. del Monte. Thanks to Havana University, Cuba; ICP-CSIC, Madrid, Spain and CONACyT, México for scientific grants.

Compliance with Ethical Standards

Conflict of Interest

The authors declare that they have no conflict of interest.

References

  1. 1.
    Goujard, L., Villeneuve, P., Barea, B., Lecomte, J., Pina, M., Claude, S., Le Petit, J., & Ferré, E. (2009). A spectrophotometric transesterification-based assay for lipases in organic solvent. Analytical Biochemistry, 385(1), 161–167.CrossRefGoogle Scholar
  2. 2.
    Nunes de Lima, L., Aragon, C. C., Mateo, C., Palomo, J. M., Giordano, R. L. C., Tardioli, P. W., Guisán, J. M., & Fernández-Lorente, G. (2013). Immobilization and stabilization of a bimolecular aggregate of the lipase from Pseudomonas fluorescens by multipoint covalent attachment. Process Biochemistry, 48(1), 118–123.CrossRefGoogle Scholar
  3. 3.
    Silva, T. C., Pereira, D. M., Valentão, P. and Andrade, P. B. (2017). Phospholipase A2 inhibitors of marine origin, in Natural products targeting clinically relevant enzymes (Andrade, P. B., Valentão, P., Pereira D. M., eds.), first edition, Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim, Germany, pp. 69–92.Google Scholar
  4. 4.
    Zarevucka, M., & Wimmer, Z. (2008). Plant products for pharmacology: application of enzymes in their transformations. International Journal of Molecular Sciences, 9(12), 2447–2473.CrossRefGoogle Scholar
  5. 5.
    Rigoldi, F., Donini, S., Redaelli, A., Parisini, E., & Gautieri, A. (2018). Review: Engineering of thermostable enzymes for industrial applications. Applied Bioengineering, 2(1), 011501.  https://doi.org/10.1063/1.4997367.CrossRefGoogle Scholar
  6. 6.
    Jakab, G., Manrique, A., Zimmerli, L., Métraux, J.-P., & Mauch-Mani, B. (2003). Molecular characterization of a novel lipase-like pathogen-inducible gene family of Arabidopsis. Plant Physiology, 132(4), 2230–2239.CrossRefGoogle Scholar
  7. 7.
    Ruiz, C., Falcocchio, S., Pastor, F. I. J., Saso, L., & Diaz, P. (2007). Helicobacter pylori EstV: identification, cloning, and characterization of the first lipase isolated from an epsilon-proteobacterium. Applied and Environmental Microbiology, 73(8), 2423–2431.CrossRefGoogle Scholar
  8. 8.
    Park, J., Cho, S.-J., & Choi, S.-J. (2008). Purification and characterization of hepatic lipase from Todarodes pacificus. BMB Reports, 41(3), 254–258.CrossRefGoogle Scholar
  9. 9.
    Sobrinho, J. C., Kayano, A. M., Simões-Silva, R., Alfonso, J. J., Gomez, A. F., Gomez, M. C. V., Zanchi, F. B., Moura, L. A., Souza, V. R., Fuly, A. L., de Oliveira, E., da Silva, S. L., Almeida, J. R., Zuliani, J. P., & Soares, A. M. (2018). Anti-platelet aggregation activity of two novel acidic Asp49-phospholipases A2 from Bothrops brazili snake venom. International Journal of Biological Macromolecules, 107, 1014–1022.CrossRefGoogle Scholar
  10. 10.
    Nevalainen, T. J., Peuravuori, H. J., Quinn, R. J., Llewellyn, L. E., Benzie, J. A. H., Fenner, P. J., & Winkel, K. D. (2004). Phospholipase A2 in Cnidaria. Comparative Biochemistry and Physiology - Part B, 139(4), 731–735.CrossRefGoogle Scholar
  11. 11.
    Perera, E., Moyano, F. J., Díaz, M., Perdomo-Morales, R., Montero-Alejo, V., Alonso, E., Carrillo, O., & Galich, G. S. (2008). Polymorphism and partial characterization of digestive enzymes in the spiny lobster Panulirus argus. Comparative Biochemistry and Physiology - Part B, 150(3), 247–254.CrossRefGoogle Scholar
  12. 12.
    Brena, B., González-Pombo, P. and Batista-Viera, F. (2013). Immobilization of enzymes: A literature survey, in Immobilization of enzymes and cells (Guisán, J. M., ed.), third edition, Methods in Molecular Biology, vol. 1051, Springer Science+Business Media, New York, pp. 15–31.Google Scholar
  13. 13.
    Barbosa, O., Torres, R., Ortiz, C., Berenguer-Murcia, A., Rodrigues, R. C., & Fernandez-Lafuente, R. (2013). Heterofunctional supports in enzyme immobilization: from traditional immobilization protocols to opportunities in tuning enzyme properties. Biomacromolecules, 14(8), 2433–2462.CrossRefGoogle Scholar
  14. 14.
    Bradford, M. M. (1976). A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry, 86, 248–254.CrossRefGoogle Scholar
  15. 15.
    Sabuquillo, P., Reina, J., Fernandez-Lorente, G., Guisan, J. M., & Fernandez-Lafuente, R. (1998). ‘Interfacial affinity chromatography’ of lipases: separation of different fractions by selective adsorption on supports activated with hydrophobic groups. Biochimica et Biophysica Acta, 1388(2), 337–348.CrossRefGoogle Scholar
  16. 16.
    Baker, J. E., Fabrick, J. A., & Zhu, K. Y. (1998). Characterization of esterases in malathion-resistant and susceptible strains of the pteromalid parasitoid Anisopteromalus calandrae. Insect Biochemistry Molecular, 28(12), 1039–1050.CrossRefGoogle Scholar
  17. 17.
    Bastida, A., Sabuquillo, P., Armisen, P., Fernández-Lafuente, R., Huguet, J., & Guisán, J. M. (1998). A single step purification, immobilization and hyperactivation of lipases via interfacial adsorption on strongly hydrophobic supports. Biotechnology and Bioengineering, 58(5), 486–493.CrossRefGoogle Scholar
  18. 18.
    Taylor, R. F. (1991). Protein immobilization. Fundamentals and applications. New York: Marcel Dekker.Google Scholar
  19. 19.
    Jain, D. K., Jain, N., Charde, R., & Jain, N. (2011). The RP-HPLC method for simultaneous estimation of esomeprazole and naproxen in binary combination. Pharmacy Methods, 2(3), 167–172.CrossRefGoogle Scholar
  20. 20.
    Patel, T. L., Patel, B. C., Kadam, A. A., Tipre, D. R., & Dave, S. R. (2015). Application of novel consortium TSR for treatment of industrial dye manufacturing effluent with concurrent removal of ADMI, COD, heavy metals and toxicity. Water Science and Technology, 71(9), 1293–1300.CrossRefGoogle Scholar
  21. 21.
    Fernández-Lorente, G., Terreni, M., Mateo, C., Bastida, A., Fernández-Lafuente, R., Dalmases, P., Huguet, J., & Guisán, J. M. (2001). Modulation of lipase properties in macro-aqueous systems by controlled enzyme immobilization: enantioselective hydrolysis of a chiral ester by immobilized Pseudomonas lipase. Enzyme and Microbial Technology, 28(4-5), 389–396.CrossRefGoogle Scholar
  22. 22.
    Lozano, P., De Diego, T., Carrié, D., Vaultier, M., & Iborra, J. L. (2004). Synthesis of glycidyl esters catalyzed by lipases in ionic liquids and supercritical carbon dioxide. Journal of Molecular Catalysis A, 214(1), 113–119.CrossRefGoogle Scholar
  23. 23.
    Sahin, O., Erdemir, S., Uyanik, A., & Yilmaz, M. (2009). Enantioselective hydrolysis of (R/S)-naproxen methyl ester with sol-gel encapsulated lipase in presence of calix[n]arene derivatives. Applied Catalysis A, 369(1-2), 36–41.CrossRefGoogle Scholar
  24. 24.
    Langer, M., Gabor, E. M., Liebeton, K., Meurer, G., Niehaus, F., Schulze, R., Eck, J., & Lorenz, P. (2006). Metagenomics: an inexhaustible access to nature’s diversity. Biotechnology Journal, 1(7-8), 815–821.CrossRefGoogle Scholar
  25. 25.
    Trincone, A. (2011). Marine biocatalysts: enzymatic features and applications. Marine Drugs, 9(4), 478–499.CrossRefGoogle Scholar
  26. 26.
    Knotz, S., Boersma, M., & Saborowski, R. (2006). Microassays for a set of enzymes in individual small marine copepods. Comparative Biochemistry and Physiology Part A, 145(3), 406–411.CrossRefGoogle Scholar
  27. 27.
    Rachamim, T., Morgenstern, D., Aharonovich, D., Brekhman, V., Lotan, T., & Sher, D. (2014). The dynamically evolving nematocyst content of an anthozoan, a scyphozoan, and a hydrozoan. Molecular Biology and Evolution, 32, 740–753.CrossRefGoogle Scholar
  28. 28.
    de Caro, J. D., Rouimi, P. and Rovery, M. (1986). Hydrolysis of p-nitrophenyl acetate by the peptide chain fragment (336-449) of porcine pancreatic lipase. European Journal of Biochemistry, 158, 601–607, 3.CrossRefGoogle Scholar
  29. 29.
    Andrade, L. H., & Barcellos, T. (2009). Lipase-catalyzed highly enantioselective kinetic resolution of boron-containing chiral alcohols. Organic Letters, 11(14), 3052–3055.CrossRefGoogle Scholar
  30. 30.
    Jústiz, O. H., Fernández-Lafuente, R., Guisán, J. M., Negri, P., Pagani, G., Pregnolato, M., & Terreni, M. (1997). One-pot chemoenzymatic synthesis of 3′-functionalized cephalosporines (cefazolin) by three consecutive biotransformations in fully aqueous medium. The Journal of Organic Chemistry, 62(26), 9099–9106.CrossRefGoogle Scholar
  31. 31.
    Pregnolato, M., Terreni, M., de Fuentes, I. E., Alcántara, A. R., Sabuquillo, P., Fernández-Lafuente, R., & Guisan, J. M. (2001). Enantioselective enzymatic hydrolysis of racemic glycidyl esters by using immobilized porcine pancreas lipase with improved catalytic properties. Journal of Molecular Catalysis B: Enzymatic, 11(4-6), 757–763.CrossRefGoogle Scholar
  32. 32.
    Tan, T., Chen, B., & Ye, H. (2006). Enzymatic synthesis of 2-ethylhexyl palmitate by lipase immobilized on fabric membranes in the batch reactor. Biochemical Engineering Journal, 29(1-2), 41–45.CrossRefGoogle Scholar
  33. 33.
    Nawani, N., Singh, R., & Kaur, J. (2006). Immobilization and stability studies of a lipase from thermophilic Bacillus sp: the effect of process parameters on immobilization of enzyme. Electronic Journal of Biotechnology, 9, 559–565.CrossRefGoogle Scholar
  34. 34.
    Gottemukkala, V. V., Saripella, K. K., Kadari, A. K., & Neau, S. H. (2008). Effect of methyl branching of C8H18 alkanes and water activity on lipase catalyzed enantioselective esterification of ibuprofen. Electronic Journal of Biotechnology, 11, 1–12.CrossRefGoogle Scholar
  35. 35.
    Palomo, J. M., Segura, R. L., Fernández-Lorente, G., Guisán, J. M., & Fernández-Lafuente, R. (2004). Enzymatic resolution of (±)-glycidyl butyrate in aqueous media. Strong modulation of the properties of the lipase from Rhizopus oryzae via immobilization techniques. Tetrahedron: Asymmetry, 15(7), 1157–1161.CrossRefGoogle Scholar
  36. 36.
    Segura, R. L., Palomo, J. M., Mateo, C., Cortes, A., Terreni, M., Fernandez-Lafuente, R., & Guisan, J. M. (2004). Different properties of the lipases contained in porcine pancreatic lipase extracts as enantioselective biocatalysts. Biotechnology Progress, 20(3), 825–829.CrossRefGoogle Scholar
  37. 37.
    Yilmaz, E., Sezgin, M., & Yilmaz, M. (2010). Enantioselective hydrolysis of racemic naproxen methyl ester with sol-gel encapsulated lipase in the presence of sporopollenin. Journal of Molecular Catalysis B: Enzymatic, 62(2), 162–168.CrossRefGoogle Scholar
  38. 38.
    Cabrera, Z., Fernandez-Lorente, G., Fernandez-Lafuente, R., Palomo, J. M., & Guisan, J. M. (2009). Novozym 435 displays very different selectivity compared to lipase from Candida antarctica B adsorbed on other hydrophobic supports. Journal of Molecular Catalysis B: Enzymatic, 57(1-4), 171–176.CrossRefGoogle Scholar
  39. 39.
    Palomo, J. M., Fernández-Lorente, G., Mateo, C., Ortiz, C., Fernández-Lafuente, R., & Guisán, J. M. (2002). Modulation of the enantioselectivity of lipases via controlled immobilization and medium engineering: hydrolytic resolution of Mandelic acid esters. Enzyme and Microbial Technology, 31(6), 775–783.CrossRefGoogle Scholar
  40. 40.
    Wang, P. Y., Tsai, S. W., & Chen, T. L. (2008). Improvements of enzyme activity and enantioselectivity via combined substrate engineering and covalent immobilization. Biotechnology and Bioengineering, 101(3), 460–469.CrossRefGoogle Scholar
  41. 41.
    Ong, A. L., Kamaruddin, A. H., Bhatia, S., & Aboul-Enein, H. Y. (2008). Enantioseparation of (R,S)-ketoprofen using Candida antarctica lipase B in an enzymatic membrane reactor. Journal of Separation Science, 31(13), 2476–2485.CrossRefGoogle Scholar

Copyright information

© Springer Science+Business Media, LLC, part of Springer Nature 2019

Authors and Affiliations

  • Alberto del Monte-Martínez
    • 1
    Email author
  • Jorge González-Bacerio
    • 1
    • 2
  • Carlos M. Varela
    • 3
    • 4
  • Fernando Vega-Villasante
    • 5
  • Rogelio Lalana-Rueda
    • 6
  • Héctor Nolasco
    • 7
  • Joaquín Díaz
    • 1
  • José M. Guisán
    • 8
  1. 1.Centro de Estudio de Proteínas, Facultad de BiologíaUniversidad de La HabanaHavanaCuba
  2. 2.Departamento de Bioquímica, Facultad de BiologíaUniversidad de La HabanaHavanaCuba
  3. 3.Rosenstiel School of Marine and Atmospheric Science (RSMAS)University of MiamiMiamiUSA
  4. 4.Florida International UniversityMiamiUSA
  5. 5.Centro Universitario de La CostaUniversidad de GuadalajaraPuerto VallartaMexico
  6. 6.Centro de Investigaciones MarinasUniversidad de La HabanaHavanaCuba
  7. 7.Centro de Investigaciones Biológicas del NoroesteConsejo Nacional de Ciencia y Tecnología (CONACyT)La PazMexico
  8. 8.Departamento de Biocatálisis, Instituto de Catálisis y PetroleoquímicaConsejo Superior de Investigaciones Científicas (CSIC) Campus CantoblancoMadridSpain

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