Production of an esterase from Fusarium oxysporum catalysing transesterification reactions in organic solvents

Process Biochemistry Vol. 33, No. 7, pp. 729-733, 199~ ¢ 199S Published by Elsevier Science Ltd. All rights reserved Printed in Great Britain I~i)32-t)592/98 $ - - see front mailer

ELSEVIER

PII:

S0032-9592t

98)00039-9

Production of an esterase from Fusarium oxysporum catalysing transesterification reactions in organic solvents P. C h r i s t a k o p o u l o s , " B. Tzalas," D. M a m m a , ~' H. Stamatis, ~' G. N. Liadakis, b C. Tzia, b D. Kekos, ~' F. N. Kolisis ~' and B. J. Macris ~'* "Biosystems Technology Laboratory, Chemical Engineering Department, National Technical University of Athens, 9 Iroon Polytechniou St, 15 700, Zografou Campus, Athens, Greece "Food Science and Technology Laboratory, Chemical Engineering Department, National Technical Universily ol Athens, 9 lroon Polytechniou St, 15 700. Zografou Campus, Athens, Greece ( R e c e i v e d 12 F e b r u a r y 1998: a c c e p t e d ,R M a r c h 1998)

Abstract

The production of an esterase by Fusarium oxysporurn, grown on tomato skins as the sole carbon source, was studied in submerged and solid state cultures. Under optimum growth conditions, enzyme yields as high as 7.3 U/ml of culture medium and 19-4 U/g of carbon source were obtained. The esterasc catalysed the synthesis of esters in organic solvents. Geraniol was transacetylated in hexane by the esterase using triacetyl as an acetyl donor. The geranyl acetate yield was 68%. © [998 Elsevier Science Ltd. All rights reserved Keywords: Fusarium oxyspomm, tomato skins, esterase, transesterification, organic solvents.

because they offer numerous potential advantages compared to aqueous media from a biotechnological perspective [3]. In such a system, hydrolytic cnzymcs can carry out synthetic reactions since the equilibrium of the reaction is shifted sufficiently enough to produce a high yield of the ester product [3]. Gcraniol esters of short-chain fatty acids, especially geranyl acetate, are important flavour and fragrance compounds widely used in the food, beverage, cosmetic and pharmaccutical industry [4,5]. The present work aims at the productkm of extracellular esterasc by Fusarium oAysporum, grown on tomato skins in submerged and solid state cultures and the use of the enzyme for transestcrification reactions in organic solvents.

Introduction

Esterases catalyse the hydrolysis of ester bonds and arc widely distributed not only in animals and plants but also in microorganisms [1]. Although investigations on microbial esterase production is limited, when compared with studies on the equivalent mammalian enzymes some microorganisms have been reported to produce cutin hydrolysing esterases (cutinases) [2]. Microbial enzyme production from wastes is a challenge to both environmental protection and process economy. Wastes from the tomato industry account for one third of the large quantities processed and are mainly obtained in the form of seed and skin residues. The latter are covered with a protective barrier, called cuticle. Its structural component, cutin, is composed of chloroform/methanol-soluble lipids and wax esters, protecting the plant from microbial and insect attack

Materials and methods

[21.

Tomato skins

There is considerable interest in the use of esterases as industrial catalysts in monophasic organic solvents

T o m a t o pomace was obtained from a tomato processing plant (KOPAIS SA, Aliartos, Greece), It was sun-dried (25-30°C, 3 - 4 days) anti ground in a blendcr

*To whom correspondence should be addressed. 729

730

P. Christakopoulas et al.

(Waring Commercial Blender, Dynamics Co., New Hartford, CO). The major part of the skins was removed using a l-mm sieve. The skins remaining on the sieve were used after separation from seeds with a fan blowing an upward air stream.

Microorganism A laboratory strain F3 of Fusarium oxysporum, isolated from cumin [6], was used in the present investigation. The stock culture was maintained on potato-dextrose agar (PDA).

Growth conditions The composition of the mineral medium was (g/litre): NaH2PO4, 10.0; KHzPO4, 1.0; CaCI2, 0.3 and MgSO4.7H20, 0.3, supplemented with 1% (w/v) ammonium phosphate as nitrogen source. The pH of the medium was adjusted to 7-0 by the addition of NaOH. A 20-ml spore suspension (105 spores) was aseptically transferred from the stock culture to a 500-ml Erlenmeyer flask containing 200ml of the above mineral medium supplemented with 0.2 g/litre glucose and 1 g/litre tomato skin. The flasks were incubated at 30°C for 24 h in a rotary shaker operating at 250 rpm and served as a preculture. The growth medium, used in submerged culture, was composed of 100 ml mineral medium, placed in 250 ml Erlenmeyer flasks and supplemented with different quantities of tomato skins (carbon source). Following heat sterilisation (121°C for 20rain), the medium was inoculated with 5% v/v of the preculture and incubated at 30°C in a rotary shaker, operating at 250 rpm. Solid state culture utilised a stationary solid layer in 100 ml Erlenmeyer flasks. The layer was composed of 2.5 g of tomato skin, moistened with different volumes of mineral medium and sterilised at 121°C for 30 min. The flasks were inoculated with 2 ml of the preculture and incubated at 30°C.

Enzyme recovery The submerged culture broth was centrifuged (7000g, 30 min) and the supernatant was used for the enzyme assay. The solid state culture flasks were extracted at 30°C with 25 ml of distilled water by shaking at 150rpm for 60min. The suspended materials and fungal biomass were separated by centrifugation (7000g, 30 rain) and the clarified supernatant was used as the source of the enzyme.

Crude enzyme preparation Following filtration (0.45 Itm) the clarified supernatant was concentrated by ammonium sulphate precipitation

(80%, w/v). The precipitate was separated by centrifugation (5000g, 10min) at 4°C, dissolved in 0-05 M acetate buffer (pH 5.0), and dialysed overnight against a 200-fold volume of deionised water at 4°C. The dialysate was freeze-dried and used for studying the enzyme properties.

Transesterification reaction For the acetylation of geraniol, 0.3 M geraniol (Sigma Chemical Co., St Louis, MO), 0.1 M triacetin (Sigma Chemical Co., St Louis, MO) and 150rag of dry esterase preparation were added to 3 ml of n-hexane. The reaction mixture was incubated at 45°C and magnetically stirred. Samples were withdrawn at various time intervals to determine the concentrations of substrate and product. All substrates and organic solvent were dehydrated with molecular sieves before use. Control experiments were carried out without the enzyme.

Analytical methods Fatty acids were determined as methyl esters by gas chromatography (GC) according to AOAC [7] procedures on a GC 8000 (Fisons Instruments, Milano, Italy). Gas chromatographic conditions were: helium carrier (50 ml/min), 60 m x 0-32 mm fused silica capillary column DB-23 (J and W Scientific, Folsom, CA, USA) packed with 50%-cyanopropyl methylpolysiloxane, running isothermally at 185°C, injector 230°C, flame ionisation detector 250°C. Compounds were identified by peak retention times, and compared with those of methyl ester standards. Moisture, ash and crude fat (ether extract) content as well as the iodine test for starch were conducted according to the AOAC methods [7]. Crude protein content was measured by the Kjeldhal method (Nx6.25). Total sugars were determined by the phenol-sulphuric acid method of Dubois [8] and expressed as glucose equivalents. Insoluble and soluble dietary fibre content was determined by the ProskyAOAC method [9]. The uronic acids content was measured by the method of Scott [10] and expressed as polysaccharide. The cellulose, hemieellulose and lignin content were determined as described previously [11]. Geraniol and geranyl acetate concentrations were determined by gas chromatography, using a Perkin Elmer 8500 chromatography apparatus, equipped with a 3 ft glass packed column, loaded with l%-Dexsil 300 (Supelco) and a FID detector. Nitrogen was used as the carrier gas at a flow rate of 12ml/min, with detector port temperature 250°C. The oven temperature was held constant for l min at 80°C and then increased linearly (10°C/rain) up to 170°C.

Enzyme assay General esterase activity was determined by a spectrophotometric assay with p-nitrophenyl butyrate

Esterase production from Fusarium oxysporum

(PNPB) (Sigma Chemical Co,, St Louis, MO) as substrate [2]. The assay mixture (1 ml) contained 0.5 mM PNPB in 51)mM phosphate buffer pH 6.0. Aliquots of 25-50 ILI of culture filtrate were added in the assay mixture and the change in absorbance at 405 nm was monitored for 1 rain. During this period of time active preparations showed a linear increase in A4osnm at 35°C. The rate of reaction was determined by reference to a standard curve prepared by adding various amounts of p-nitrophenol to the assay mixture. PNB was substituted with other p-nitrophenyl esters (0.5mM) in this assay to determine the effect of p-nitrophenyl ester chain length on activity. For determining the kinetic constants K,,,, and Vm~,~the action of the csterase on PNPB was studied at varying concentrations of the substratc 10-1-5 mM).

731

their suitability as a carbon source for microbial growth, The skins also contained approximately 3.0% (w/w) crude fat with oleic linoleic and palmitic as the predominant fatty acids, accounting for 90% of the total recovered fatty acids (Table 1). It has been reported that the presence of biopolymers containing lipids in the growth medium induces the synthesis of esterases in microorganisms [2, 13, 14].

Submerged culture

Protein concentration was determined by the dye-binding procedure of Bradford [12] using bovine serum albumin as a standard.

Tomato skin concentration had a linear effect on esterase activity up to 7% (w/v) [Fig. I(A)]. Higher concentrations were not feasible for reasons related to viscosity of the growth medium. Time-course experiments of enzyme production, carried out at the optimum tomato skin concentration (7%, w/v), showed optimum activity at the 5th day of growth [Fig. I(B)]. The observed maximum esterase activity was 7-8 U/ml of growth medium. This activity level compared favourably with those reported in the literature for microbial csterases [2, 15, 16].

Results and discussion

Solid state culture

Tomato skin composition

The effect of the moisture content of the growth medium on esterase activity was studied using four different moisture levels. The results appear in Fig. 2(A) and show that a moisture level equal of 70% yielded the highest enzyme activity. When esterase production was examined during the course of growth, optimum esterase activities were obtained at the 5th day of growth [Fig. 2(B)]. As reported elsewhere [17-19] high moisture content enhanced fungal growth and enzyme production in solid state cultures.

Protein estimation

The chemical analysis of dry tomato skins revealed that they were rich in carbohydrates (Table 1), justifying Table 1. Composition of tomato skin Skin component

(% dw)

Moisture Total sugars Crude fat Crude protein Ash Pectin Cellulose Hemicellulose Lignin

7-5 5-0 3"0 13.5 2.9 15.6 27.7 9.1 15'7

Crude fat component

(%)

Lauric Myrictic Palmitic Stearic Oleic Linolcic Not determined

0.5 1.4 15"8 4.8 36.4 38-5 2.7

Enz3,me characterisation Important kinetic characteristics of crude esterase were calculated. The optimum temperature of activity was 70°C and 34% of this activity remained at 80°C. PNPB hydrolysis revealed apparent Km and V,..... values of

8

E7

A

6 5 4 3

5 •~

4 3

2 1 uJ 0

B

7

6

i

!

I

I

i

i

i

1

2

3

4

5

6

7

2 1 0-

v

0

8

Tomato skin concentration (%, w/v)

0

1

2

3

4

5

6

7

Growth time (days)

Fig. 1. Effect of tomato skin concentration (growth time: 5 days) (A) and growth time (tomato skin concentration 7e/~, w/v) (B) on esterase production in submerged culture.

P Christakopoulas et al.

732 A

25

25

.

L~

U) aD

20

-"1

15

A

,y

10 5 0

ul

40

I

I

I

I

50

60

70

80

10~ ~ 90

0

I

I

I

I

I

I

1

2

3

4

5

6

7

Growth time (days)

Moisture c o n t e n t (%)

Fig. 2. Effect of moisture content (growth time: 5 days) (A) and growth time (moisture content: 70%) on esterase production in solid state culture.

Table 2. Substrate specificity of esterase Esterase activity (%)

Substrate

100 24 nil nil

p-Nitrophenyl-butyrate p-Nitrophenyl-acetate p-Nitrophenyl-laurate p-Nitrophenyl-palmitate

I

100

Conclusions

Esterase was produced by the fungus E oxysporum grown on tomato skins, an annual by-product of the tomato canning industry. Both submerged and solid state cultures were successfully used for the production of the enzyme. The esterase was well suited for the production of geranyl acetate by transesterification in organic solvents. This esterase seems to have the potential of becoming an important enzyme for the synthesis and production of industrially significant acetates and esters in organic solvents.

' O Geranyl acetate I

8O

o~ c

60

•

40

o

co

2(1 O

centration increased during the course of reaction time reaching a constant value after 96 h. The initial rate of transesterification was 5-7 mM/h and compared favourably with that reported in the literature for other esterases [5, 25].

References •

0

i

i

i

i

i

20

40

60

80

100

T i m e (hr)

Fig. 3. Esterase transesterification of geraniol by triacetin in n-hexane.

0"7 mmol and 1.28 mmol/min/mg protein, respectively. The effect of acyl ester chain length on esterase activity towards p-nitrophenyl esters was investigated (Table 2). As with other microbial esterases [15, 20-24], the enzyme showed specificity only towards short-acylchain esters, being more active on p-nitrophenylbutyrate.

Transesterification reaction Triacetin was chosen as the acetyl donor for the acetylation of geraniol by the esterase. The results of enzymic transesterification of geraniol by triacetin in hexane are shown in Fig. 3. The geranyl acetate con-

1. Krisch, K., In The Enzymes, ed. P.D. Boyer. Academic Press, New York, 1971, pp. 43-69. 2. Fett, W.F., Gerard, H.C., Moreau, R.A., Osmaan, S.F. and Jones, L.F., Cutinase production by Streptomyces spp. Current Microbiology 1992, 25, 165-171. 3. Dodrick, J.S., Enzymatic catalysis in monophasic organic solvent. Enzyme Microbiology Technology 1989, 11, 194-211. 4. Stamatis, H., Xenakis, A., Provelegiou, M. and Kolisis, F.N., Esterification reactions catalysed by lipases in microemuisions. The role of enzyme localisation in relation to its selectivity. Biotechnology and Bioengineering 1993, 42, 103-110. 5. Karra-Chaabouni, M., Pulvin, S., Touraud, D. and Thomas, D., Enzymatic synthesis of geraniol esters in a solvent-free system by lipases. Biotechnology Letters 1996, 18, 1083-1088. 6. Christakopoulos, P., Macris, B. and Kekos, D., Direct fermentation of cellulose to ethanol by

Fusariurn oxysporum. Enzyme Microbiology Technology 1989, 11, 236-239. 7. AOAC, Official Methods of Analysis of the Associa-

Esterase production from Fusarium oxysporum tion of Official Analytical Chemists. AOAC Inc., USA, 1990. 8. Dubois, M., Gilles, K.A., Hamilton, J., Rebers, P.A. and Smith, F., Colorimetric method for the determination of sugars and related substances. Analytical Chemistry 1956, 28, 35(t-356. 9. Prosky, L., Asp, N.-G., Schweizer, T.F., DeVreis, J.W. and Furda, I., Determination of insoluble, soluble and total dietary fibre in foods products: interlaboratory study. Journal of the Association o1" Official Analytical Chemists 1988, 71, 1017-1023. 10. Scott, R.W,, Colorimetric determination of hexuronic acids in plant materials. Analytical Chemistry 1979, 51,936-941. 11. Aravantinos-Zafiris, G., Oreopoulou, V., Tzia, C. and Thomopoulos, C.D., Fibre fraction from orange peel residues after pectin extraction. Lebensmittel Wissenscht{ft und Technologie 1994, 27, 468-471. 12. Bradford, M.M., A rapid and sensitive method for the quantification of microgram quantities of protein utilising the principle of protein dye binding. Analytical Biochemist~ 1976, 72, 248-254. 13. Lin. T.S. and Kolattukudy, P.E., Induction of a biopolyester hydrolase (cutinase) by low levels of cutin monomers in Fusarium solani sp. pisi. Journal of Bacteriol{zw 1978, 133, 942-951. 14. Lin. T.S. and Kolattukudy, P.E., Isolation and characterization of cuticular polyester (cutin) hydrolyzing enzyme from phytopathogenic fungi. Physiology, and Plant Pathol~w 1980, 17, 1-15. 15. McQueen, D.A. and Schottel, J.I., Purification and characterization of a novel extracellular esterase from pathogenic Streptomyces scabies that is inducible by zinc. Journal of" Bacteriolo~' 1987, 169, 1967-1971. 16. Meghi, K., Ward, O.P. and Araujo, A., Production, purification, and properties of extracellular carboxy[ csterases from Bacillus subtilis NRRL 365. Applied Environmental Microbiolog3' 1990, 56, 3735-3740.

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17. Dechamps, F., Giuliano, C., Asther, M.. Huet, M.C. and Roussos, S., Cellulase production by Trichoderma harzianurn in static and mixed solid state fermentation reactors under nonaseptic conditions. BiotechnoloL,v and Bioengineerin~ 1985, 27, 1385-1388. 18. Abdulah, A.L., Tergendy, R.P. and Murphy, V.G., Optimization of solid substrate fermentation of wheat straw. Biotechnolotw and Bioengineering 1985, 27, 211-27. 19. Roussos, S., Raimbault, M., Saucedo-Castaneda, G., Vinicgra-Gonzalez, G. and Lonsane, B.K., Kinetics and ratios of carboxy-methyl cellulase and flter paper activities of the cellulolytic enzymes produced by Trichoderma harzlanum on different substrates in solid state fermentation. Micologia Neotropieal Aplicada 1991, 4, 19-40. 20. Sobek, H. and Gorisch, tt.. Purification and characterization of a heat-stable estcrase from the thermoacidophilic archaebacterium Sulfolobus acidocaldarius. Biochernistn' Journal 1988, 250, 453-458. 21. Higerd, T.B., Isolation of acetvl esterase mutants of Bacillus subtilis 168. Journal of Bacteriolog3' 1977, 129, 973-977. 22. Matsunaga, A., Koyama, N. and Nosoh, Y., Purification and properties of esterase from Bacillus

stearothermot~hih~s. Archives ~l Biochemistry and Biophysics 1974, 160, 504-513. 23. Nakagawa, A., Tsujita, T. and Okuda, H., Purification and some properties of intracellular esterasc from Pseudomonas .[tuorescens. Journal ~l Biochemi,~tO' 1984, 95, 1047-1054. 24. Toshimitsu, N., Hamada, H. and Kojima, M., Purification and some properties of an esterase from yeast..loumal of Fetvnentation ]~'chnolqq~, 1996, 64, 459-4~'12. 25. Lin, T.S. and Kolattukudy, P.E., Isolation and characterization of cuticular polyester (cutin) hydrolyzing enzyme from phytopathogenic fungi. Physiolo£,y and Plant Pathologq' 198(I, 17, I - 15.