Journal of Biotechnology 66 (1998) 41–50

Biodegradable products by lipase biocatalysis

Yu-Yen Linko a,*, Merja Lamsa b, Xiaoyan Wu a, Esa Uosukainen a, Jukka Seppala a,Pekka Linko a

a Department of Chemical Technology, Helsinki Uni6ersity of Technology, P.O. Box 6100, FIN-02015 HUT, Finlandb Vegetable Oil Laboratory, Raisio Chemicals Ltd., FIN-21200 Raisio, Finland

Received 6 January 1998; received in revised form 5 March 1998; accepted 13 March 1998

Abstract

The interest in the applications of biocatalysis in organic syntheses has rapidly increased. In this context, lipaseshave recently become one of the most studied groups of enzymes. We have demonstrated that lipases can be used asbiocatalyst in the production of useful biodegradable compounds. A number of examples are given. 1-Butyl oleatewas produced by direct esterification of butanol and oleic acid to decrease the viscosity of biodiesel in winter use.Enzymic alcoholysis of vegetable oils without additional organic solvent has been little investigated. We have shownthat a mixture of 2-ethyl-1-hexyl esters can be obtained in a good yield by enzymic transesterification from rapeseedoil fatty acids for use as a solvent. Trimethylolpropane esters were also similarly synthesized as lubricants. Finally,the discovery that lipases can also catalyze ester syntheses and transesterification reactions in organic solvent systemshas opened up the possibility of enzyme catalyzed production of biodegradable polyesters. In direct polyesterificationof 1,4-butanediol and sebacic acid, polyesters with a mass average molar mass of the order of 56000 g mol−1 orhigher, and a maximum molar mass of about 130000 g mol−1 were also obtained by using lipase as biocatalyst.Finally, we have demonstrated that also aromatic polyesters can be synthesized by lipase biocatalysis, a higher than50000 g mol−1 mass average molar mass of poly(1,6-hexanediyl isophthalate) as an example. © 1998 Elsevier ScienceB.V. All rights reserved.

Keywords: Biocatalysis; Biodegradable; Enzyme; Esterification; Lipase, Rapeseed oil

1. Introduction

Lipases (triacylglycerol acylhydrolase, EC3.1.1.3) are enzymes which normally catalyze thehydrolysis of glycerol esters at lipid/water inter-faces. In organic solvent systems, however, lipases

* Corresponding author. Tel.: +358 9 4512540; fax: +3589 462373; e-mail: yu-yen.linko@hut.fi

0168-1656/98/$ - see front matter © 1998 Elsevier Science B.V. All rights reserved.

PII S0168-1656(98)00155-2

Y.-Y. Linko et al. / Journal of Biotechnology 66 (1998) 41–5042

have also been shown to catalyze ester synthesis(Klibanov, 1989), they have been employed in themodification of fats and oils both to providenovel materials of improved characteristics and toupgrade inexpensive raw-materials to valuableproducts by transesterification (West, 1988), andthey have been used in the production ofbiodegradable polyesters (Linko et al., 1995a,c;Linko and Seppala, 1996). Nevertheless, enzymicalcoholysis of vegetable oils without additionalorganic solvent has been little investigated (Ishii etal., 1990; Mittelbach, 1990). Further, the discov-ery that lipases can also catalyze ester synthesesand transesterification reactions in organic solventsystems has opened up the possibility of enzymecatalyzed synthesis of biodegradable polyesters(Wallace and Morrow, 1989; Linko et al., 1994b,1995a,b). Biocompatibility, biodegradability, andenvironmental acceptability of biotechnically pro-duced polyesters are desired properties in agricul-tural and medical applications (Evans and Sikdar,1990).

Different microbial strains produce enzymes ofvarying characteristics such as hydrolytic and syn-thetic activity, stability, etc. The increasedavailability and stability of commercially availablelipases has resulted in markedly increased interestin the potential applications of lipase biocatalysis(Bjorkling et al., 1991; Vulfson, 1994). Thepresent paper discusses the lipase catalyzed syn-thesis of 1-butyl oleate as an additive in biodieselduring winter use, the synthesis of rapeseed oilfatty acid esters, rapeseed oil fatty acid 2-ethyl-1-hexyl ester by enzymic alcoholysis as a solvent incar shampoos and trimethylolpropane (TMP) es-ter as a biodegradable hydraulic oil and lubricant.Finally, the synthesis of high molar massbiodegradable aliphatic and aromatic polyesters isdescribed.

2. Materials and methods

2.1. Enzymes

Candida rugosa (ex. cylindracea) (42.5 U mg−1;water 5.0%), Chromobacterium 6iscosum (13.3 Umg−1; water 5.9%), Rhizomucor miehei (4.0–7.2

U mg−1, 7.4% water) and Pseudomonas fluores-cens (11.9 U mg−1; water 3.1%) lipase powderswere obtained from Biocatalysts (Pontypridd,UK) and used without further treatment. Immo-bilized lipases Lipozyme IM (R. miehei lipaseexpressed in Aspergillus oryzae) and Novozym435 (Candida antarctica lipase expressed in A.oryzae) were obtained from Novo Nordisk A/S(Bagsvaerd, Denmark). One unit of activity wasdefined as that amount of enzyme which catalyzesthe release of 1 mmol of free fatty acid from oliveoil in 1 min at pH 7.0, 37°C.

2.2. Chemicals

1-Butanol was obtained from E. Merck (Darm-stadt, Germany) and oleic acid from EastmanKodak (Rochester, NY). Refined low erucic acidrapeseed oil and synthetic rapeseed oil 2-ethyl-1-hexylester were obtained from Raisio Group (Rai-sio, Finland). The approximate fatty acidcomposition of the oil was 57% oleic acid, 22%linoleic acid, 12% linolenic acid, 4% palmitinicacid, 2% eicosaenoic acid, 1% stearic acid, B1%erucic acid, 1% others. 2-Ethyl-1-hexanol (watersolubility at 25°C :2.5%) and terephthalic acidwere obtained from Fluka Chemie AG (Buchs,Switzerland). Isophthalic acid was from Riedel-deHaen AG (Seelze, Germany). 1,4-Butanedioland diphenyl ether were from Aldrich-ChemieAG (Steinheim, Germany), and 1,6-hexanediolfrom Merck (Darmstadt, Germany). Mono-, di-and triolein standards were from Sigma (St.Louis, MO), and glycerol from May and Baker(Dagemham, UK). All other chemicals used wereof analytical grade with a purity of 99.5% orhigher. Organic solvents were stored over a 4-A(0.4 nm) molecular sieve.

2.3. 1-Butyl oleate synthesis

1-Butyl oleate synthesis was carried out at 37°Cin 8-ml screw-capped flasks containing varyingquantities of different lipases and the substrates atdifferent molar ratios, with or without variousquantities of added water. The contents werestirred with a magnetic stirrer at 200 rpm. At agiven time a suitable quantity of a mixture (1:1,

Y.-Y. Linko et al. / Journal of Biotechnology 66 (1998) 41–50 43

v/v) of diethylether and ethanol was added, andthe free oleic acid was titrated immediately with0.1 M sodium hydroxide. The yield, in %, wascalculated from oleic acid consumed on the ba-sis of the limiting substrate.

2.4. Synthesis of 2-ethyl-1-hexyl esters ofrapeseed oil fatty acids

A preliminary study of the transesterificationbetween 2-ethyl-1-hexanol and rapeseed oil withthe four lipases (10 mg; 3.3%) was carried outin capped 13-ml test tubes under magnetic stir-ring at 200 rpm, using 0.680 mmol (107 m l) of2-ethyl-1-hexanol, 0.227 mmol (:0.2 g) rape-seed oil (molar ratio 3:1) and 3.0% added water.After 72 h lipase was separated by centrifuga-tion for 5 min at 2000 rpm (Martin Christ TypeUJ3, Osterode, Germany), and the supernatantwas pipetted into Eppendorf tubes for storing at−20°C for later analysis. Subsequent transester-ification reactions were carried out for up to 72h using varying substrate molar ratios (ethylhexanol to rapeseed oil from 10 to 0.5), C.rugosa lipase (from 0.3 to 14.6% of substrates),added water (from none to 50%), and tempera-tures (from 37 to 60°C). In semi-pilot scale ex-periments the reaction mixture contained 25 ml(20 g) to 1.0 l (829 g) 2-ethyl-1-hexanol and50–2000 g of rapeseed oil, (substrate molar ra-tio of 2.8), 3.4% (w/w) lipase both with andwithout a carrier, and 1–5% (w/w) added water.

2.5. Trimethylolpropane esters of rapeseed oilfatty acids

2.5.1. Chemical synthesis of rapeseed oil methylester (RMe)

Rapeseed oil methyl ester was synthesizedchemically as follows: rapeseed oil (0.3 mol) wasweighed into a 100 cm3 3-necked flask, equippedwith a thermometer, condenser, stirrer, and sam-ple adapter, and methanol (2.0 mol) was addedunder stirring. The reaction mixture was thenheated to 60°C and 0.5% (w/w) alkaline catalystwas added. After the reaction was completed in4 h as determined by thin layer chromatography(TLC), the reaction mixture was washed by

acidic water. Glycerol formed was separated andthe excess alcohol was distilled off. The RMe(average molar mass 884 g mol−1, meltingrange 56–59°C) content of the product variedfrom 95 to 99%, as determined by HPLC.

2.5.2. Enzymic synthesis of rapeseed oiltrimethylolpropane ester

Transesterification for the synthesis oftrimethylolpropane tri-ester of rapeseed oil fattyacids (average molar mass 922 g mol−1) wascarried out either at ambient pressure in cappedor open 13 cm3 Kimax test tubes or at a re-duced pressure (2.0–13.3 kPa) in 25 cm3 roundbottomed flasks equipped with a vertical con-denser (cooling water temperature 6°C) typicallyas follows: trimethylolpropane (0.607 g, 4.5mmol) was dissolved in 0.7 ml (15%, w/w oftotal mass of the substrates) of water, afterwhich RMe (4.00 g, 13.6 mmol) and solid lipasepreparation (40% w/w) were added. Reactionwas usually carried out at a temperature be-tween 37 and 47°C under a reduced pressure ofabout 5–16 kPa with magnetic stirring at 250rpm (or 150–200 rpm for the round bottomedflasks), and the TMP to RMe ratio was either1:3.5 or 1:4.5. The total sample was extractedtwice with acetone, after which the enzyme pre-cipitate was removed by centrifuging at 3500rpm (RCF 1900×g). The supernatant wastransferred into a 1.5 cm3 Eppendorf tube andstored at −20°C for later analysis.

2.6. Polyester synthesis

Typical lipase catalyzed polymerization reac-tion was carried out as follows: crude lipasepowder was added to a reaction mixture ofequimolar quantities of a dicarboxylic acid or itsderivative and an aliphatic diol in an organicsolvent, usually diphenyl ether. The mixture wasstirred by a magnetic stirrer at 600 rpm, 37–45°C for aliphatic and 60°C for aromaticpolyesters (unless otherwise stated). After thefirst 20 h, the polymerization was completed inabout 20 kPa (0.15 mmHg) vacuum. In order toobtain a high average molar mass polyester itwas very important that the water or alcohol

Y.-Y. Linko et al. / Journal of Biotechnology 66 (1998) 41–5044

formed was removed. When the reaction wascompleted in about 3–6 days, the product wasextracted with chloroform, lipase was filteredoff, chloroform was removed by distillation at40–50°C under a reduced pressure, and a whitesolid polyester was obtained upon precipitationfrom methanol. Blank tests, without lipase, re-sulted in no observable polymerization.

2.7. Analytical methods

Qualitative analyses were carried out by thinlayer chromatography (TLC). Samples were di-luted 1:10 (v/v) with ethanol, and 0.01 ml of thediluted samples were used for TLC analysis.Hexane:diethylether:acetic acid (80:20:1 v/v) wasused as solvent on Kieselguhr 60 F254 plates (E.Merck, Darmstadt, Germany) with one hourrunning time. Slightly dried plates were sprayedwith 0.1% 2%,7%-dichlorofluorescein (Aldrich-Chemie, Steinheim, Germany) in 99.5% ethanol(Alko, Finland) for detecting of the spots at 254and 360 nm.

Rape seed oil conversion (% rapeseed oilused) and ester yield (% of theoretical) was de-termined using reversed phase high performanceliquid chromatography (HPLC) as modifiedfrom El-Hamdy and Perkins (1981) and Forssellet al. (1993) employing a Perkin-Elmer (Nor-walk, CT) four pump module, ISS-100 sampler,and 101 oven, Novapack C18 3.9×150 mmcolumn with 4-mm silica particles, HP 1047A re-fractive index detector, PE 316 integrator andPE 7500 professional computer. The runningsolvent was acetone:acetonitrile (1:1 v/v) at 1.0ml min−1, 37°C, 30 min. Residual 2-ethyl-1-hex-anol could not be determined by the HPLCmethod, because the alcohol overlapped with thesolvent peak. Consequently, any excess 2-ethyl-1-hexanol was determined by TLC as describedabove. Samples were diluted with acetone to10–20 mg ml−1, filtered through a Millex-LCR4

disposable filter of 0.5-mm porosity (Millipore,Bedford, UK), and 0.02 ml of the filtrate wasused for the analysis.

Moisture content of the enzyme preparationswas determined by drying about 4 g samplesovernight at 105°C.

2.8. Molar mass measurement

Mass average molar mass (M( m) was determinedby GPC using three serially connected styragelcolumns (0.01, 0.05, and 1 mm) and an HP 1047 Arefractive index detector (Hewlett-Packard). Elevendifferent polystyrene standards with known molarmasses ranging from 162 to 370000 g mol−1

(Polymer Laboratories, Church Stretton, UK) wereused for constructing a calibration curve. TC*SECsoftware and Turbochrom chromatography system(both from Perkin Elmer, Norwalk, CT) were usedfor data analyses. Samples were dissolved intetrahydrofuran to a concentration of about 0.05g ml−1, and filtered through a 0.5 mm Millex-LCR4disposable filter (Millipore) or 0.45 mm GHPAcrodisc Syringe Filter (Gelman Sciences). For theGPC analysis, 50-m l samples of the filtrate wereinjected. Tetrahydrofuran was used as the mobilephase at a flow rate of 0.8 ml min−1 at ambienttemperature.

2.9. Melting point measurement

Melting point of the polyester was determined bydifferential scanning calorimeter (DSC), using PLThermal Science DCS equipment (ReometricScientific, UK). Nitrogen was used as the sweepinggas, and a 4–8 mg sample was heated twice at arate of 10°C min−1. The scanning temperaturerange was from −20 to 180°C.

3. Results and discussion

3.1. Lipase catalyzed 1-butyl oleate synthesis

Commercial lipases are sold on the basis oftheir hydrolytic activity, and there is no guaranteein regard to their esterification activity. Thereappears to be little correlation between thehydrolytic and ester synthesis activities ofdifferent lipases and, as has been recently shown,even lipases from different fermentation batchesof a similar hydrolytic activity may vary widely intheir esterification activities (Wu and Linko, 1996;Wu et al., 1996a). Consequently, it is necessary toscreen lipases for their desired synthetic activity

Y.-Y. Linko et al. / Journal of Biotechnology 66 (1998) 41–50 45

under various processing conditions. On screening25 commercially available lipases in the absenceof additional organic solvent using the synthesisof butyl oleate as the model system, we found thathigh ester yields were obtained only with lipasesfrom Candida rugosa, Chromobacterium 6iscosum,Rhizomucor miehei, and Pseudomonas fluorescens(Linko et al., 1990). The ester yield was affectedby the initial water content of the system, lipasequantity, and the molar ratio of 1-butanol to oleicacid. Without additional water, only Ch. 6iscosumlipase yielded 98% of 1-butyl oleate in 12 h with a1-butanol excess. The addition of 3.2% waterincreased the initial rate of reaction. With an oleicacid excess only 0.3% (of substrates) of C. rugosaor R. miehei lipases with an initial water contentof 3.2% and 14% was needed to yield 94 and100% ester, respectively. The lipase from C.rugosa was found to give both high ester yieldsand superior cost/benefit ratio in direct biocata-lytic synthesis of n-butyl oleate.3.2. 2-Ethyl-1-hexyl esters from rapeseed oil

One of our aims in rapeseed oil transesterifica-tion with 2-ethyl-1-hexanol was to obtain a maxi-mum rapeseed oil conversion with no or littleresidual substrates. Unlike in the butyl oleatesynthesis, rapeseed oil conversion was always lowwhen an alcohol excess was used. In such a case,the product mixture contained large quantities ofresidual alcohol often with residual oil. On screen-ing, when C. rugosa lipase was employed as thebiocatalyst a 98% conversion of rapeseed oil wasobtained in 24 h with no residual rapeseed oil andlittle byproducts detectable by TLC (Linko et al.,1994a). Relatively high yields were also obtainedwith Ch. Viscosum and P. fluorescence lipases(Table 1). C. rugosa lipase was chosen for furtherstudies because of its superiority in catalyzingboth esterification (Linko et al., 1992) and trans-esterification (Otero et al., 1990; Wu et al., 1996b)reactions, and its commercial availability in largequantities at a relatively low cost. As could beexpected, an increase in lipase quantity markedlyincreased both 1-butyl oleate synthesis and therapeseed oil conversion during the first few hours,but in about 7–12 h the differences were almostleveled off.

The importance of the control of water content(and of water activity) in lipase catalyzed estersyntheses has been often emphasized (Halling,1987; Malcata et al., 1992; Wehtje and Adler-creutz, 1997). Although a minimum quantity ofwater is necessary for enzyme catalysis to takeplace, ester synthesis is favored under restrictedwater availability (low water activity). Ester syn-thesis and hydrolysis are reversible processes, andthe equilibrium may be shifted towards synthesiseither by an excess of one of the substrates or bycontrolling of the water content of the reactionsystem. As with the most lipases used in 1-butyloleate synthesis, the water (:5%) present in thelipase preparations used was insufficient in trans-esterification, and without any added water onlyan about 25% conversion was reached in 7 h.With a minimum of about 1.0% of added water,an about 50% conversion was reached already inone hour, and a nearly complete conversion in 5h. Further, best results were obtained with analcohol to rapeseed oil molar ratio of 2.8:1. Therewas little difference in conversion within the tem-perature range of 37–55°C, and an about 90%conversion in 2–3 h and a nearly complete con-version in 7 h was obtained in all cases. Thisagrees well with the results of Mittelbach (1990),according to whom the optimal temperature ofCandida sp. lipase catalyzed sunflower oil alcohol-ysis is between 45 and 50°C. At 60°C lipase wasclearly inactivated under the experimentalconditions.

Most previous papers on lipase catalyzed ester-ification and transesterification have involved im-

Table 1The production of 2-ethyl-1-hexanol rapeseed oil fatty acidester by different lipasesa

Lipase Conversion (%)

24 h 48 h

98Candida rugosa 98Chromobacterium 6iscosum 9796

99Pseudomans fluorescens 968745Rhizomucor miehei

a 2-Ethyl-1-hexanol to rapeseed oil molar ratio 3.0, 3.3% li-pase, and 3% added water.

Y.-Y. Linko et al. / Journal of Biotechnology 66 (1998) 41–5046

mobilized lipase (Eigtved, 1989; Leitgeb andKnez, 1990; Forssell et al., 1993). Also in thepresent case a carrier was thought to help inkeeping the enzyme more evenly dispersed. Noimprovement was found with the addition of glassbeads, polyurethane foam or Amberlite XAD-2resin (Lamsa et al., 1994), but with AmberliteXAD-7 resin as a carrier an about 90% (approxi-mately 97% of theoretical) rapeseed oil (2 kg)conversion was achieved in 8 h with 1 l (829 g) of2-ethyl-1-hexanol, 3% added water, 100 g of li-pase, 300 g of XAD-7 resin, 170 rpm, and 37°C.This was equal to 22.4 g of ester per g lipase or112 g ester per 100 g rapeseed oil.

3.3. Transesterification of trimethylolpropane andrapeseed oil methylester

We have previously demonstrated that immobi-lized R. miehei lipase Lipozyme IM 20 can beused as biocatalyst in the synthesis of TMP estersof rapeseed oil fatty acids (Linko et al., 1997).The highest total conversions of about 95% toTMP esters were obtained only in 24 h (47°C, 5.3kPa, 13% water) with the lipase immobilized onDuolite ES-561 (40%, w/w, biocatalyst). Thehighest yield of about 70% TMP tri-ester wasreached in 78 h. With the commercial immobilizedR. miehei lipase Lipozyme IM 20 conversion toTMP tri-ester was about 75% in 24 h (58°C, 5.3kPa, no added water), while total conversions toTMP esters of as high as 92.5% were obtained.

In the present work, crude C. rugosa lipasepowder was used as the biocatalyst without anyadditional organic solvent. The absence of solventallows higher substrate and product concentra-tions, simplifies down stream processing, and im-proves safety (Ergan et al., 1991; Ison et al.,1988). In preliminary trials using capped testtubes with a number of commercially availablecrude lipases at 37°C, C. rugosa lipase appearedagain to be superior to other lipases tested, al-though all conversions obtained were low. Whenthe temperature was increased to 47°C and therelative lipase quantity to 20% (of substrates) atthe substrate molar ratio of 3.5, the rapeseed oilconversion was increased to about 95% in 68 h.However, with substrate molar ratios of less than

3.5 or higher than 4.5 considerable quantities ofunreacted rapeseed oil methyl ester remained inthe mixture. Osada et al. (1987) synthesized polyolesters using tetrahydrofuran as a solvent and Rhi-zopus delemar lipase as biocatalyst at ambientpressure, but yields were relatively low, for exam-ple only 40% in the case of pentaerythritol mono-caproate in 72 h at 30°C. In the present work, anincrease in temperature up to 58°C did not im-prove conversion, apparently owing to methanolformed during the transesterification. Under areduced pressure of 2.0 kPa the yield increased toabout 68% in a round bottomed flask equippedwith a vertical condenser under otherwise identi-cal conditions

Interestingly, high conversions were obtainedwith as much as 15% of added water. When thetemperature was increased to 47°C with 15%added water, an about 60% conversion to TMPtri-ester was reached already in 24 h and 73% in68 h, with no byproducts nor any residual RMe.Under such conditions, the total conversion toTMP esters increased to about 75% in 24 h, witha maximum yield of 98%. Although generallylipase catalyzed esterification reaction is slowerthan hydrolysis, according to Monot et al. (1990)TMP tri-caprylate is very difficult to hydrolyzeenzymically. They employed a two- to three-foldmolar excess of acid in order to drive the reactiontowards completion. This may, at least in part,explain the relatively high transesterification rateand yield even at high water concentration. How-ever, from a practical application point of view asmall amount of the di-ester is of a much lesserproblem than a large excess of unreacted acid orRMe.

3.4. Synthesis of biodegradable aliphaticpolyesters

Powdered ‘dry’ R. miehei lipase was used inmost experiments for the synthesis of aliphaticpolyesters owing to the favorable structure of it’sactive center (Linko and Seppala, 1996; Jaaske-lainen et al., 1996, 1997). The type of organicsolvent appeared to be a very important factor inthe lipase catalyzed aliphatic polyester synthesis(Wang et al., 1996). Diphenyl ether was found to

Y.-Y. Linko et al. / Journal of Biotechnology 66 (1998) 41–50 47

Table 2Effect of chain length of dicarboxylic acid and diol on mass average molar mass of aliphatic polyestersa

C3-diol C4-diol C5-diol C6-diolC2-diol

980C4-diacid (succinic acid) 374052902140M( w 1150

28.4DP 6.8 7.3 12.4 18.770 430C6-diacid (adipic acid) 37 2706420M( w 38 97026 830

174.2DP 37.3 114.3 194.8 308.943 48048 590C8-diacid (octanedioic acid) M( w 55 0202090 51 730

227.3 169.8213.1DP 10.4 241.746 130 62 480C10-diacid (sebacic acid) 53 680M( w 9760 35 550

219.9198.8180.2DP 42.8 146.916 22021 450C12-diacid (dodecanedioic acid) M( w 28 3208960 26 070

52.095.0DP 35.0 96.6 75.5

a 1.5 mmol dicarboxylic acid with 1.5 mmol diol at 37°C, 7 days with programmed vacuum catalyzed by 0.25 g M. miehei in 2.25ml diphenyl ether. Yields varied from 85 to 93%.

be a superior solvent, and was used in mostexperiments. There seemed to be no correlationbetween the log P value of the solvent and thesynthetic activity of lipase.

With equimolar quantities of the activatedbis(2,2,2-trifluoroethyl) sebacate and 1,4-butane-diol, an aliphatic polyester of a mass averagemolar mass (M( m) of 46130 g mol−1 (DP]183)or higher was obtained in vacuum at 37°C in 3days in diphenyl ether. The M( m of the polyesterincreased with an increase in the substrate concen-tration of up to about 0.83 M. With hexanedioicacid and hexandiol an M( m of up to 77400 gmol−1 [DP=340, Pd (M( m/M( n)=4.4, and a melt-ing temperature of 66.8°C as determined by DSC]was obtained, with a maximum molar mass of131190 g mol−1 (DP=520) (Table 2). This isbelieved to be the highest M( m reported by lipasecatalyzed polyesterification between a diacid anda diol. 13C NMR studies showed that the partiallypurified white solid obtained was a linearpoly(1,4-butyl sebacate). In general, this meansthat we obtained at least a four-fold increase inthe M( m with the polyesterification of underiva-tized diacids and diols in a vacuum in comparison

to the polyesterification at atmospheric pressure,the molar masses obtained in a vacuum were ofthe same order of magnitude than those we hadobtained earlier in the polytransesterification ofan activated, derivatized sebacic acid and 1,4-bu-tanediol (Linko et al., 1995b).

3.5. Synthesis of aromatic polyesters

Although of several different commerciallyavailable crude lipases R. miehei lipase resulted inthe highest mass average molar mass in the syn-thesis of aliphatic polyesters, surprisingly this li-pase was unable to catalyze the synthesis of thearomatic polyester of poly(1,6-hexanediyl isoph-thalate) beyond oligomer stage. In this case, onlythe immobilized Novozym 435 lipase was foundto be an efficient biocatalyst under the conditionsused. Even with Novozyme 435 no polyester wasobserved when terephthalic acid (1,4-benzene di-carboxylic acid) was used as the diacid, while ahigh molar mass was obtained by using isoph-thalic acid (1,3-benzene dicarboxylic acid) as sub-strate. This suggested that the relative position ofthe two functional carboxylic groups on a benzene

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Table 3Effect of different substrates and enzymes for aromatic polyester synthesis

Substrates Molar mass (g mol−1)

45°C 60°CDiacid 70°CDiol

Novozyme 435n.d. n.d.Terephtalic acid n.d.1,4-butanediol

n.d.n.d.n.d.1,6-hexanediolTerephtalic acid2300 n.d.Isophtalic acid 1,4-butanediol 1100

38 000 55 000Isophtalic acid 1,6-hexanediol 2000

Lipozyme IM600n.d. —1,6-hexanediolIsophthalic acid

n.d., Not detectable.

ring was a critical factor for the lipase-catalyzedpolyesterification (Table 3). This result was in anagreement with that described recently by Mezoulet al. (1996), who attempted to prepare aromaticpolyesters from dimethyl phthalate, terephthalateor isophthalate and 1,6-hexanediol in toluene, us-ing Novozym 435 lipase as the biocatalyst. Noproduct was found with dimethyl phthalate.Dimethyl terephthalate gave only a molar mass ofabout 2800 g mol−1, whereas dimethyl isophtha-late resulted in a high molar mass of 31700 gmol−1.

In the present work, the type of diol was alsofound to be important in order to obtain a highmolar mass aromatic polyester. With isophthalicacid and 1,4-butanediol as substrates and 0.25 g(8.1%, w/w of reaction mixture) of Novozym 435as the biocatalyst, the mass average molar mass ofthe polyester obtained was only 1100 g mol−1 at45°C and about 2300 g mol−1 at 60°C, but with1,6-hexanediol a mass average molar mass of38000 g mol−1 was obtained at 60°C and as highas about 55000 g mol−1 at 70°C under otherwisesimilar reaction conditions. The effect of tempera-ture on the synthesis of poly(1,6-hexanediyl isoph-thalate) has been shown to be nearly linear, withan increase in M( m from 45 to 70°C

Considerably higher quantity of Novozym 435lipase and a longer reaction time was required forthe aromatic polyester synthesis than had beenpreviously shown to be necessary for the synthesisof aliphatic polyesters (Linko et al., 1995a; Wu etal., 1996c). Vacuum was found to be very impor-

tant also in the lipase catalyzed synthesis of aro-matic polyesters in order to obtain a high massaverage molar mass. For example, with vacuum,the highest obtained mass average molar mass ofthe poly(1,6-hexanediyl isophthalate) was about50000 g mol−1, while without vacuum a molarmass of only about 16000 g mol−1 was reached.

The melting point of the obtained poly(1,6-hex-anediyl isophthalate) ranged from 60 to 90°Cprobably due to the variations in the molar massdistribution of the polyester.

4. Conclusions

We have demonstrated above the feasibility ofthe production of a variety of biodegradable es-ters and polyesters by using lipase as the biocata-lyst. The best results depended critically on theright choice of the process conditions and of thelipase used. In the esterification of oleic acid with1-butanol, high 1-butyl oleate ester yields wereobtained with crude lipases from C. rugosa, Ch.6iscosum, R. miehei, and P. fluorescens with opti-mal addition of water. In transesterification of 2kg rapeseed oil with 2-ethyl-1-hexanol, 97% con-version of theoretical was obtained using C.rugosa lipase powder as the biocatalyst underoptimal conditions. In the more complex case oftrimethylpropane esters, up to 75% conversion totri-esters was obtained in 24 h under a reducedpressure, with the commercial immobilized R.miehei lipase Lipozyme IM 20 as biocatalyst, and

Y.-Y. Linko et al. / Journal of Biotechnology 66 (1998) 41–50 49

with no added water, while the total conversion toTMP esters exceeded 90%. Similar high yieldswere also obtained by using crude C. rugosa lipasepowder. We have also shown that biodegradablelipase catalyzed polyesterification both ofaliphatic and aromatic substrates is possible. Un-der vacuum, linear polyesters of up to of anM( m=77400 g mol−1 with a maximum molarmass of the order of 130000 g mol−1 of poly(1,4-butyl sebacate), and of an M( m=50000 ofpoly(1,6-hexanediyl isophthalate) were obtainedwith powdered ‘dry’ R. miehei and commercialimmobilized C. antarctica Novozym 435 lipases,respectively.

Acknowledgements

The authors are grateful to the Academy ofFinland for financial support.

References

Bjorkling, F., Godtfredsen, S.E., Kirk, O., 1991. The futureimpact of industrial lipases. Trends Biotechnol. 9, 360–363.

Eigtved, P., 1989. Immobilized Mucor miehei lipase for trans-esterification. US patent no. 4,798,793.

El-Hamdy, A.H., Perkins, E.G., 1981. High-performance re-versed-phase chromatogrpahy of natural triglyceride mix-tures: critical pair separation. J. Am. Oil Chem. Soc. 58,867–872.

Ergan, F., Trani, M., Andre, G., 1991. Use of lipases inmultiphasic systems solely composed of substrates. J. Am.Oil Chem. Soc. 68, 412–417.

Evans, J.D., Sikdar, S.K., 1990. Biodegradable plastics: anidea whose time has come? Chemtech 20, 38–42.

Forssell, P., Parovuori, P., Linko, P., Poutanen, K., 1993.Enzymatic transesterification of rapeseed oil and lauricacid in a continuous reactor. J. Am. Oil Chem. Soc. 70,1105–1109.

Halling, P.J., 1987. Water activity in biphasic reaction systems.In: Laane, C., Tramper, J., Lilly, M.D. (Eds.), Biocatalysisin Organic Media. Elsevier, Amsterdam, pp. 125–132.

Ishii, T., Mori, T., Chen, J., Itoh, Y., Shimura, S., Kirimura,K., Usami, S., 1990. Ester synthesis by a crude lipase ofRhizopous oligosporus in aquaeous system. J. Ferment.Bioeng. 70, 188–189.

Ison, A.P., Dunnill, P., Lilly, M.D., 1988. Effect of solventconcentration on enzyme catalysed interesterification offats. Enzyme Microb. Technol. 10, 47–51.

Jaaskelainen, S., Wu, X.Y., Linko, S., Wang, Y., Linko,Y.-Y., Teleman, O., Linko, P., 1996. Production, charac-terization and molecular modeling of lipases for esterifica-tion. Ann. NY Acad. Sci. 799, 129–138.

Jaaskelainen, S., Linko, S., Raaska, T., Laaksonen, L., Linko,Y.-Y., 1997. Molecular modelling of lipase catalyzedpolyester synthesis. J. Biotechnol. 52, 267–275.

Klibanov, A.M., 1989. Enzymatic catalysis in anhydrous or-ganic solvents. Trends Biochem. Sci. 14, 141–144.

Leitgeb, M., Knez, Z., 1990. The influence of water on thesynthesis of n-butyl oleate by immobilized Mucor mieheilipase. J. Am. Oil Chem. Soc. 67, 775–778.

Linko, Y.-Y., Seppala, J., 1996. Producing high molecularweight biodegradable polyesters. Chemtech 26 (8), 25–31.

Linko, Y.-Y., Koivisto, U.-M., Kautola, H., 1990. Optimiza-tion of enzymatic production of oleic acid. Ann. NY Acad.Sci. 613, 691–696.

Linko, Y.-Y., Rantanen, O., Yu, H.-C., Linko, P., 1992.Factors affecting lipase catalyzed n-butyl oleate synthesis.In: Tramper, J., Vermue, M.H., Beeftink, H.H., vonStockar, U. (Eds.), Biocatalysis in Non-Conventional Me-dia, Progress in Biotechnology 8. Elsevier, Amsterdam, pp.601–608.

Linko, Y.-Y., Lamsa, M., Huhtala, A., Linko, P., 1994a.Lipase catalyzed transesterification of rapeseed oil and2-ethyl-1-hexanol. J. Am. Oil Chem. Soc. 71, 1411–1414.

Linko, Y.-Y., Wang, Z., Seppala, J., 1994b. Lipase-catalyzedsynthesis of poly(1,4-butanediol succinate) in organic sol-vent. Biocatalysis 8, 269–282.

Linko, Y.-Y., Wang, Z.-L., Seppala, J., 1995a. Lipase cata-lyzed linear aliphatic polyester synthesis in organic solvent.Enzyme Microb. Technol. 17, 506–511.

Linko, Y.-Y., Wang, Z.-L., Seppala, J., 1995b. Lipase-cata-lyzed synthesis of poly(1,4-butyl sebacate) from sebacicacid or its derivatives with 1,4-butanediol. J. Biotechnol.40, 133–138.

Linko, Y.-Y., Lamsa, M., Huhtala, A., Rantanen, O., 1995c.Lipase biocatalysis in the production of esters. J. Am. OilChem. Soc. 72, 1293–1299.

Linko, Y.-Y., Tervakangas, T., Lamsa, M., Linko, P., 1997.Production of trimethylolpropane esters of rapeseed oilfatty acids by immobilized lipase. Biotechnol. Technol. 11,889–892.

Lamsa, M., Huhtala, A., Linko, Y.-Y., Linko, P., 1994.2-Ethyl-1-hexanol fatty acid esters from rapeseed oil bytransesterification. Biotechnol. Technol. 8, 451–456.

Malcata, F.X., Reyes, H.R., Garcia, H.S., Hill, C.G. Jr.,Amundson, C.H., 1992. Kinetics and mechanisms of reac-tions catalysed by immobilised lipases. Enzyme Microb.Technol. 14, 426–446.

Mezoul, G., Lalot, T., Brigodiot, M., Marechal, E., 1996.Enzyme-catalyzed syntheses of poly(1,6-hexanediyl isoph-thalate) and poly(1,6-hexanediyl terephthalate) in organicsolvent. Polym. Bull. 36, 541–548.

Mittelbach, M., 1990. Lipase catalyzed alcoholysis ofsunflower oil. J. Am. Oil Chem. Soc. 67, 168–170.

Y.-Y. Linko et al. / Journal of Biotechnology 66 (1998) 41–5050

Monot, F., Benoit, Y., Ballerini, D., Vandecasteele, J.-P.,1990. Enzymatic synthesis of neopentylpolyol esters inorganic media. Appl. Biochem. Biotechnol. 24/25, 375–386.

Osada, F., Kobayashi, M., Asaoka, S., Kitazato, H., 1987.Preparation of polyol esters by lipase-catalyzed esterifica-tion. Japan Kokai Tokkyo Koho JP 62 296 884.

Otero, C., Pastor, E., Ballesteros, A., 1990. Synthesis ofmonobutyrylglycerol by transesterification with solubleand immobilized lipases. Appl. Biochem. Biotechnol. 26,35–44.

Vulfson, E.N., 1994. Industrial applications of lipases. In:Woolley, P., Petersen, S.B. (Eds.), Lipases: Their Struc-tures, Biochemistry and Applications. Cambridge Univer-sity Press, Cambridge, pp. 271–287.

Wallace, J.S., Morrow, C.J., 1989. Biocatalytic synthesis ofpolymers: synthesis of an optically active, epoxy-substi-tuted polyester by lipase-catalyzed polymerization. J.Polym. Sci. A, Polym. Chem. 27, 2553–2567.

Wang, Z.-L., Hiltunen, K., Orava, P., Seppala, J., Linko,

Y.-Y., 1996. Lipase-catalyzed polyester synthesis. J.Macromol. Sci. A, Pure Appl. Chem. A33, 599–612.

Wehtje, E., Adlercreutz, P., 1997. Lipases have similar wateractivity profiles in different reactions. Biotechnol. Lett. 11,537–540.

West, S.I., 1988. Enzymes in the food processing industry.Chemistry in Britain 12, 1220–1222.

Wu, X.Y., Linko, Y.-Y., 1996. Variations among four com-mercial lipases from Chromobacterium 6iscosum in estersynthesis. Process Biochem. 31, 765–772.

Wu, X.Y., Jaaskelainen, S., Linko, Y.-Y., 1996a. Purificationand partial characterization of Rhizomucor miehei lipasefor ester synthesis. Appl. Biochem. Biotechnol. 59, 145–158.

Wu, X.Y., Jaaskelainen, S., Linko, Y.-Y., 1996b. An investiga-tion of various lipases catalyzing hydrolysis, esterficationand transesterification. Enzyme Microb. Technol. 19, 226–232.

Wu, X.Y., Seppala, J., Linko, Y.-Y., 1996c. Lipase-catalyzedpolyester synthesis. Biotechnol. Technol. 10, 793–798.

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