Effect of different carbon sources on lipase production byCandida rugosa<

E. Dalmau*,a, J.L. Montesinosa, M. Lottib, C. Casasa

aDepartament d’Enginyeria Quı´mica, Escola Te`cnica Superior d’Enginyeria, Universitat Auto`noma de Barcelona, 08193 Bellaterra, Barcelona, SpainbDipartimento di Fisiologia e Biochimica Generali, Sezione di Biochimica Comparata, Universita’ degli Studi di Milano, Milano, Italy

Received 23 March 1999; received in revised form 4 August 1999; accepted 26 August 1999

Abstract

Different carbon sources affecting growth and lipase production inCandida rugosawere studied by using batch cultures on definedmedium. Carbohydrates and acids non-related to fats did not induce lipase production. The highest yields of enzyme were obtained withlipids or fatty acids as carbon sources. Tween 80 stimulated lipase biosynthesis and secretion outside the cell. Combinations of two typesof substrates, carbohydrates and fatty acids, did not improve lipase production, and in some cases, their consumption was produced in asequential pattern. Glucose presented a repressing effect on lipase production. Moreover, glucose was found to be effective in stimulatinglipase secretion by cells with a high level of cell-bound lipase activity because of their previous growth in oleic acid. © 2000 ElsevierScience Inc. All rights reserved.

Keywords:Lipase production; Carbon source; Physiological regulation;Candida rugosa

1. Introduction

Lipases (glycerol ester hydrolases EC 3.1.1.3) are en-zymes that are extremely versatile because of the largenumber of reactions, not necessarily esterification reactions,that they can catalyse. Lipases are produced by a wide-spread number of microorganisms, including bacteria [1],fungi, and yeasts [2]. In recent years, research on microbiallipases has increased because of their practical applicationsin industry, as the hydrolysis of fats, production of fattyacids and food additives, synthesis of esters and peptides,resolution of racemic mixtures, or additives in detergents[3,4]. However, factors controlling lipase synthesis andtransport have been investigated only in a few cases. InPseudomonassp. for example, the production of lipase hasbeen shown to be strongly induced by triglycerides anddetergents and not repressed by glucose or glycerol. On the

other hand, long chain fatty acids, such as oleic acid,strongly inhibit lipase production [5–7].

In fungi, although lipidic substrates and fatty acids gen-erally act as inducers, in many species, such asAspergillus[8,9] or Rhizopus[10,11], lipases are produced constitu-tively. The yeastCanadida rugosais an important lipaseproducer. Although biochemical, structural, and catalyticproperties ofC. rugosalipase have been widely documented[12–14], less information about the factors and conditionsthat control its biosynthesis and secretion is available. Alsobecause of the presence of multiple lipase isoenzymes en-coded by a family of related genes data reported in theliterature are difficult to interpret [15]. Thus, althoughChang et al. [16] reported lipase production in rich mediumin the presence of glucose, Valero et al. [17] showed thatlipase production was sensitive to glucose repression. Morerecently, it was suggested that the expression of lipase-encoding genes might indeed be subjected to differentmechanisms of control [18]. Accordingly, the compositionof the culture medium, in particular the incorporation ofdifferent lipidic substances, can result in the production ofdifferent isoenzymes. In a similar way, it has been recentlydemonstrated that lipase- and esterase-specific activity ra-tios of C. rugosalipase can be modified by acting on the

<This work was supported by NATO Collaborative Research Grant9602248, Spanish CYCIT (QUI97-0506-C03) and by EC (Project BIO-4-96-0005). E.D. Also acknowledges award of a grant from CIRIT (Gener-alitat de Catalunya).

* Corresponding author. Tel.:134-9358-1-10-18; fax:134-9358-1-20-13.

E-mail address:dalmau@uab-eq.uab.es (E. Dalmau).

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0141-0229/00/$ – see front matter © 2000 Elsevier Science Inc. All rights reserved.PII: S0141-0229(00)00156-3

operational conditions used during growth, which is consis-tent with a modified isoenzymes composition [19].

As a consequence, the choice of the culture conditions,and in particular of the medium composition, is important toproduction ofC. rugosa lipase preparations with definedand reproducible properties. In this context, it is necessaryto obtain an exhaustive description of the effect of a widerange of substrates on the yield and quality of the enzyme.In previous work the influence of several fatty acids onlipase expression was investigated [20]. In this paper, wesystematically studied the effect of several carbon sourcespreviously not considered as substrates for the growth ofC.rugosaduring lipase production. Results on stimulation oflipase secretion induced by glucose are also presented.

2. Materials and methods

2.1. Microorganism, medium, and growth conditions

C. rugosa(ATCC 14830) was maintained on peptone-malt extract agar plates at 4°C. Basal medium, preparedwith distilled water, contained per liter 15 g KH2PO4, 5.5 gK2HPO4, 5 g (NH4)2SO4, 0.5 g MgSO4 z 7H2O, 0.1 g NaCl,and 0.1 g CaCl2, 1 ml of micronutrients solutions, and 1 mlof vitamin solution. The micronutrients solution containedper liter 500 mg of H3BO3, 40 mg of CuSO4, 100 mg of KI,200 mg of FeCl3, 400 mg of MnSO4, 200 mg of Na2MoO4,400 mg of ZnSO4. The vitamin solution contained per liter2 mg biotin, 400 mg of calcium pantothenate, 2 mg of folicacid, 2000 mg of inositol, 400 mg of niacin, 200 mg ofp-amino benzoic acid, and 400 mg of pyridoxine hydrochlo-ride, 200 mg of rivoflavin, and 400 mg of thiamine hydro-choloride. Different carbon sources were added at a finalconcentration of 2 g/1. The pH of the medium was 6.2before autoclaving. Basal medium was steam sterilized, andthe micronutrients and the vitamins were sterilized by mi-crofiltration (0.22mm).

The microorganism was grown in 1 liter flasks contain-ing 100 ml of medium. For inoculation, cells that werepreviously cultured in agar plates, were resuspended in0.9% NaCl solution and 5 ml added in every flask. Theflasks were incubated in a reciprocal shaking-bath at 30°C,150 rpm, for 24 or 48 h, depending on the carbon sourceused. Experiments in bioreactor were carried out in a BiofloIII (New Brunswick Scientific, Edison, NJ, USA). The cul-ture volume was 1 liter and the operational conditions were30°C, agitation 500 rpm, aeration 0.1 1/min, and pH5 6.2controlled with NH4OH 2M.

2.2. Biomass

Biomass was determined by dry weight. Samples werefiltered (0.45 mm), washed with a mixture of dioxane-propionic acid (1:1), and then washed with 20 ml of distilledwater. The filters were dried at 95°C to constant weight.

2.3. Enzyme Analysis

2.3.1. Turbidimetric assay of extracellular lipase activityThis assay was based on modified Monotest Lipase using

triolein as a substrate (Boehringer Kit 159697, Mannheim,Germany). The reagent was mixed with 5 ml of Tris-HCl200 mM buffer (pH 7.4). A sample (250ml), previouslyfiltered through a 0.45mm filter, was added to the reagentsolution (750ml) in a thermostatically controlled cuvette(42°C), and the decrease in absorbance at 340 nm (UV) wasfollowed for 6 minutes. The absorbance decrease per unit oftime was calculated from the slope of the curve. The first 2min of analysis were not used in slope determination. Thismethod was correlated with the titrimetric method, as pre-viously described [21]. One unit of extracellular lipase ac-tivity was defined as the amount of lipase necessary tohydrolyse 1mmol of ester bond per minute under assayconditions. If the activity was higher than 0.5 units/ml, thesample had to be diluted to be included in the linear rangeof the assay.

2.3.2. Extracellular esterase activity assayThe esterase activity was measured from the enzymatic

hydrolysis ofp-nitrophenyl propionate (PNPP) at 25°C. Inthis assay 750ml of 0.4 mM PNPP in 50 mM phosphatebuffer (pH 7.0; 80%) and acetonitrile (20%) solution wereadded to 250ml of sample that was previously filteredthrough a 0.45mm filter. The formation rate ofp-nitrophe-nol (« 5 6.517 mM z cm) was measured at 348 nm for 6min. At this wavelength, the absorbance is pH-independent.One unit of extracellular esterase activity was defined as thequantity of enzyme necessary to release 1mmol of p-nitrophenol per minute under assay conditions. If the activ-ity was higher than 0.050 units/ml, the sample had to bediluted to be included in the linear range of the analysis.

2.3.3. Cell-bound lipase analysisCells were harvested by centrifugation at 50003 g for

10 min at 4°C, washed in 10 mM Tris-HCl buffer (pH 8.0),and resuspended to a 5-ml final volume with the samebuffer. The cell suspension was disrupted with glass beadsfor 8 periods of 30 s. The disrupted cells were centrifugedat 40003 g for 10 min at 4°C. The supernatant was used asthe cell extract for the determination of cell-bound lipase byusing the turbidimetric lipase analysis described above.Cell-bound concentration units are given per g or mg of drybiomass.

2.3.4. Western blotting analysisCells extracts and concentrated culture media were sub-

jected to SDS-PAGE electrophoresis in 15% acrylamidegel. After separation, proteins were blotted onto a nitrocel-lulose membrane (Amersham International, Amersham,UK) for 1 h at 4°C and 500 mA.After overnight membranesaturation with 5% nonfat milk at 4°C, lipase detection wasperformed with polyclonal antibodies raised against the

658 E. Dalmau et al. / Enzyme and Microbial Technology 26 (2000) 657–663

homologous lipase fromGeotrichum candidum[22] diluted1:8000 in a buffer containing Tris-HCl pH 7.4, 0.9% NaCl,and 5% nonfat milk. Detection was performed by using aECL chemiluminescent kit (Amersham International, Am-ersham, UK).

2.4. Analytical procedure

2.4.1. Protein analysisExtracellular protein concentration was determined with

the bicinchoninic acid protein assay kit (Pierce BCA ProteinAssay, Prod. No. 23225, Rockford, IL, USA), according tothe manufacturer’s instructions and using the enhanced pro-tocol. Bovine serum albumin was used as the protein stan-dard for the calibration curve.

2.4.2. Oleic acid analysis

Oleic acid concentration was determined with modifiedLowry-Tinsley method [23] composed of two stages. First,samples were pretreated extracting oleic acid with two liq-uid-liquid chloroform (5 ml) extractions in acid medium (1M HCl, pH 2). Then, chloroform was eliminated to drynessin a rotary evaporator, and finally, n-hexane was added toresuspend the dry sample.

2.4.3. Glucose analysisGlucose concentration was determined with a Biochem-

istry Analyser (YSI model 2700 SELECT, Yellow Springs,Ohio, USA) based on the principle of immobilised glucoseoxidase.

3. Results

3.1. Lipase production on simple substrate use

A range of different carbon sources, mainly carbohy-drates, alcohols, acids, and lipids were screened for their

capacity to support growth ofC. rugosacultures and lipaseproduction (Table 1). Arabinose, xylose, oxalic acid, andcetyl and stearyl alcohols were not used by the microorgan-ism as carbon sources. On the basis of enzyme activityassays, it was concluded that good growth, but no lipaseactivity, was obtained on media supplemented with glucose,sorbitol, glycerol, and acetic, lactic, citric, and succinicacids as sole carbon sources.

As indicated in Table 1, high biomass, but low enzymeactivity, were obtained with galactose and mannitol,whereas lipid-related substrates supported cell growth andmaximum activity levels. Among them, caprylic acid wasnot used by the microorganism as a carbon source becauseof its possible toxicity [20], and oleyl alcohol gave thelowest activity level, about 0.2 units/ml (data not shown).On the other hand, palmitic acid gave the highest enzymeactivity observed, approximately 10-fold higher, and alsothe maximum activity yield (4.3 units/mg biomass).

However, the total protein of the culture supernatants didnot correlate with extracellular lipase activities, with galac-tose exhibiting maximum protein production with minimumlipase activity, and palmitic acid giving maximum specificactivity of the supernatant (0.7 units/mgprotein).

As a further and complementary assay, the presence oflipase protein by Western blotting hybridisation with lipasecross-reacting antibodies (Fig. 1) was investigated. In gen-eral, the immunological detection confirms the results basedon enzyme activity with the exception of the samples ob-

Table 1Effect of different C-sources (2 g/l) in the medium on the growth and production of lipase byC. rugosa

Carbon source Biomass(g/l)

Extracel. lipaseactivity (units/ml)

Cell-bound lipase(units/gcells)

Extracel. esteraseactivity (units/ml)

Extracel. protein(mg/ml)

Glucose 0.97 0 0 ND* NDGalactose 0.85 0.1 0 0.001 52.7Mannitol 1.10 0.1 0 0.003 14.7Glycerol 1.25 0 0 0.002 5.9Sodium lactate 0.44 0 0 0 0.6Palmitic acid 1.23 5.3 52 0.036 7.5Oleic acid 1.55 0.4 25 0.013 39.5Triolein 1.61 0.3 72 0.004 3.7Tween 80 0.75 0.4 1 0.004 2.4

* ND: not determined.a The microorganism was grown in 1 liter flasks with 100 ml of medium, at 30°C. All the cultures were incubated for 48 h, except glucose, galactose,

mannitol and glycerol, that were maintained for 24 h.

Fig. 1. Western blotting analysis of proteins secreted byC. rugosa incultures grown as in Table 1. Each lane contains 200ml of culture medium.Lanes: C, control (500 ng of purified lipase); lanes 1 and 2, cells grown onmannitol-containing medium for 24 and 48 h; lanes 3 and 4, galactose 24and 48 h; lanes 5 and 6, glycerol 24 and 48 h; lane 7, lactate 48 h; lane 8,palmitate 48 h; lane 9, Tween 80 48 h; lane 10, oleic acid 48 h; lane 11,triolein 48 h.

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tained from culture media of cells growing on glycerol-containing medium: the level of lipase protein (lane 5) wascomparable to that obtained in the experiments with man-nitol (lane 1), whereas no significant level of lipase activitywas observed in this sample (Table 1).

In good agreement with the results presented in Table 1,the highest level of extracellular lipase protein was detectedin medium supplemented with palmitic acid as a sole carbonsource. Interestingly, two proteins differing slightly in sizewere observed. This may be explained by a microheteroge-neity in the protein glycosylation. The amount of lipaseprotein detected when using palmitic acid as the carbonsource appeared to be at least 2-fold higher than that ob-served with oleic acid, the inducer more commonly used inthis kind of experiment.

Measurement of cell-bound lipase activity showed thatdetectable levels were attained only with substrates thatgave maximum extracellular activity. Growth on Tween 80as the source of carbon exhibited a high extracellular lipaseof 0.4 units/ml, but at the same time, showed a low cell-bound lipase production, probably indicating an additionaleffect of Tween 80 on permeabilisation of the yeast cells. InWestern blotting, very faint reacting bands were detected inthe cells grown in palmitic, oleic acid, and triolein (notshown).

The highest esterase activity (0.036 units/ml) was alsoreached with palmitic acid, whereas lower results wereobtained with oleic acid (0.013 units/ml) and other lipidcompounds (0.004 units/ml). Soluble carbon sourcesshowed lower esterase activity (,0.004 units/ml) than didlipidic substances.

3.2. Lipase production on mixed substrate use

To elucidate some aspects of the induction effect oflipid-related substrates on lipase production, mixed carbonsources media were prepared by using a soluble compound,selected from previous experiments for its growth promot-

ing capacity, and a fatty acid selected because of its influ-ence on the induction of lipolytic activity. That is to say, thetwo substrates, soluble compound and fatty acid, wereadded together to the medium from the beginning of thecultures, giving the microorganism the possibility to useboth substrates in a sequential or simultaneous way, depend-ing on its metabolism. This issue would have importantconsequences from the production point of view. If theinducer was not consumed during lipase production, itwould be relatively easy to control the production system,whatever the operational mode used, batch, fed-batch, orcontinuous.

The use of mixed carbon sources did not improve pre-vious results obtained by using sole lipid related substratesas a carbon source. No increase in activity was observed inthese cultures (Table 2), although biomass production washigher in most cases, suggesting a possible competing effectof some soluble carbon sources or a close relation betweenextracellular lipase activity production and fatty acids con-sumption. In glucose medium with oleic acid added, lipaseyields related to oleic acid were similar to that obtained witholeic acid alone, provided that enough time to achieve thetotal exhaustion of both carbon sources was supplied. In-creased activity relative to biomass was observed by usingsodium lactate and oleic, palmitic, or stearic acids comparedto other soluble substrates, indicating a possible no-repres-sor effect of this carbon source. Again, palmitic acid in themedium, comparised with other acids, increased the level ofsupernatant activity detected. In spite of promoting highgrowth and lipase yield, the use of palmitic acid as the solecarbon source has operational problems because of its in-solubility.

The effect of glucose on lipase production was studiedby carrying out three types of experiments. First,C. rugosawas grown in a bioreactor with a mixture of glucose (2 g/l)and oleic acid (1 g/l). When both substrates where addedtogether, glucose was used preferentially to oleic acid,which was metabolized once the glucose was rather ex-

Table 2Effect of mixed carbon-sources in the medium on the growth and production of lipase byC. rugosa

No-lipid substrate(2 g/l)

Fatty acid(0.5 g/l)

Biomass(g/l)

Extracel. lipase activity(units/ml)

YLip/X

(units/mgcells)

Glucose Oleic acid 1.48 0.1 0.07Glucose Palmitic acid 1.11 0.4 0.36Glucose Stearic acid 0.99 0.4 0.40Galactose Oleic acid 1.36 0.4 0.29Galactose Palmitic acid 1.09 0.4 0.37Galactose Stearic acid 0.92 0.3 0.33Mannitol Oleic acid 1.57 0.3 0.19Mannitol Palmitic acid 1.28 0.4 0.31Mannitol Stearic acid 1.21 0.4 0.33Sodium lactate Oleic acid 0.81 0.5 0.62Sodium lactate Palmitic acid 0.51 0.7 1.37Sodium lactate Stearic acid 0.23 0.5 2.17

a The microorganism was grown in flasks with 100 ml of medium at 30°C for 48 h.

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hausted and without any significant lag period, as can beseen in Fig. 2. In each phase, characteristic specific growthand substrate consumption rates were observed, beinghigher with glucose than for oleic acid. Final biomass typ-ically ranges from 2.0 to 2.5 g of dry weight per liter. Lipaseactivity was not detected until the glucose had been com-pletely exhausted and oleic acid started to be consumed. Thevalues of lipase production in the bioreactor were signifi-cantly higher than those observed in shake-flask studies.

The second experiment was planned to study the effectof glucose addition to a culture growing on oleic acid. Twodifferent 100-ml samples were withdrawn at 18 and 19 hpost inoculation and cultivated with added glucose (seeTable 3) in shake flasks. Oleic acid was present in the firstsample, whereas it was completely exhausted in the secondone. After 2 h of incubation, growth went on, and extracel-lular lipase activity increased in both cultures. However,data presented in Table 3 indicate that total lipase amountdid not changed significantly, but cell-bound lipase de-creased approximately to the same extent that extracellular

lipase increased. In the second culture a higher level of totallipase activity was obtained. This was probably due to thelonger time that cells had been in the presence of theinducer. Thus, the cell-bound lipase-specific band observedin Western blotting vanished in the final sample (Fig. 3).These results suggested that glucose addition to oleic acid-growing cells stopped lipase synthesis but facilitated itssecretion to the extracellular medium.

To verify the effect of glucose addition in a culture witha high content of both biomass and lipase activity, a thirdtype of experiment was carried out. A fed-batch strategywith oleic acid as a sole carbon source, according to previ-ous experiments by Gordillo et al. [19], was used. In thiscase, the oleic acid feeding was stopped and 6 h later, 10 g/lof glucose was added. This fact could stimulate the cells tosecret cell-bound lipase once the oleic acid was exhausted.

Just before adding the glucose, the biomass concentra-tion was 7.8 g/l, and 3 h later, when the glucose wasexhausted, the biomass reached a concentration of 10.9 g/l.In Fig. 4 the time course of cell-bound, extracellular, andtotal lipase activity are presented. The general behaviour ofthe cell-bound and extracellular lipase activities was similarbut with opposite variations, while total activity kept con-stant. These results are in concordance with those obtainedin the shake-flask experiments presented above. Taking intoaccount that any variation in cell-bound and extracellularlipase was not detected for the late, nonfeeding period, itwas concluded that differences obtained after glucose addi-

Fig. 2. Time course of growth (F) and extracellular lipase activity (Œ) ofCandida rugosaculture growing in a mixture of glucose (■) and oleic acid(}) as carbon sources.

Table 3Effect of additional glucose consumption in flasks on the growth, production and secretion of lipase byC. rugosagrowing previously with oleic acid (3g/l) in a Bioflo III bioreactor of 1 l of working volume

Flask 1 Flask 2

Initialt 5 18 h

Finalt 5 20 h

Initialt 5 19 h

Finalt 5 21 h

Glucose (g/l) 4.32 1.97 5.26 2.60Oleic acid (g/l) 0.21 0 0 0Biomass (g/l) 2.45 3.62 2.75 3.82Extracel. lipase activity (units/ml) 4.0 5.0 6.0 6.4Cell-bound lipase (units/mgcells) 0.76 0.24 0.67 0.27Total lipase (units/ml) 5.9 5.9 7.8 7.4

Fig. 3. Western blotting analysis ofC. rugosacultures grown on oleicacid-containing medium for 18 h and then supplemented with glucose.Samples were withdrawn 2 h after glucose addition. Lanes 1 and 3, 0.5 mgof cell extracts (cell-bound protein); lanes 2 and 4, 200ml of culturemedium (extracellular proteins); lane C, control with 500 ng of purifiedlipase.

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tion were a direct consequence of glucose effect on lipasesecretion.

4. Discussion

Previous works on the physiology of lipase productionshowed that the mechanisms regulating biosynthesis varywidely in different microorganisms. Results obtained withCalvatia [24], Rhizopus [11], Aspergillus [9], andRhodotorula[25], showed that lipase production seems tobe constitutive and independent of the addition of lipidicsubstrates to the culture medium, although their presenceenhanced the level of lipase activity produced. On the otherhand, it is well known that, in other microorganisms such asG. candidum[26], lipidic substrates are necessary for lipaseproduction, and also, carbohydrates can act as repressors ofits biosynthesis inFusarium[27].

This paper describes how, inC. rugosa, lipidic substratesand their metabolites (long-chain fatty acids) participate inthe expression of lipase. Some substrates, typically carbo-hydrates and acids nonrelated to fats, support good growth,but very low lipase production was obtained. Lipidic sub-strates induced lipase production, the fatty acids being themost effective inducers. These results are comparable withthat reported by Nahas [10] onRhizopus oligosporusand byShimada [26] et al. onG. candidum.

Data obtained with medium supplemented with Tween80 showed similar extracellular enzyme activities and lowercell-bound lipase compared to media containing other lip-idic compounds, indicating that enzyme biosynthesis wasfollowed by almost its total secretion. It is well known thatvarious compounds, such as surfactants, can increase yeast

cells permeability, facilitating the export of several com-pounds across the cell through its membrane [28]. There-fore, it is likely that Tween 80, in addition to inducing lipasebiosynthesis, increases cell permeability, and consequentlycell-bound lipase decreased so significantly.

Comparison of lipase and esterase activities in superna-tants of different cultures showed that both types of activitywere produced (except for glycerol-supplemented medium),but a close quantitative relationship between them was notobtained, probably indicating that different enzymes areresponsible for both activities.

Combinations of two types of substrates do not improvelipase production. When using glucose and oleic acid asmixed substrates as carbon-sources, their consumption isproduced in a sequential pattern. From experiments onmixed glucose and oleic acid use, it is apparent that lipaseproduction is repressed by the glucose present in the definedmedium. These results are in accordance with those ofNahas [10] forR. oligosporus, Baillargeon et al. [29] forG.candidum, and Rapp [27] forFusarium, in spite of theirbeing related to other microorganisms, but differ from thoseobtained forC. rugosaby Chang et al. [16] who do notreport a repressive effect of glucose.

Extracellular lipase normally appears in the culture me-dium at the end of the exponential growth phase. Simulta-neously, a significant level of cell-bound lipase was accu-mulated inside the cells. Addition of glucose to oleic acid-growing cells strongly represses lipase synthesis, suggestinga mechanism of catabolite repression for the control of thisenzyme production. Nevertheless, glucose stimulates thesecretion of lipase accumulated inside the cells, but theexact mechanism for this stimulation has not been studied.

The present study clearly demonstrates the request forlipidic substrates (or their metabolites, such as fatty acids)and the inhibition effect of glucose in the production ofsignificant amounts of lipase byC. rugosa. These data willbe used for future improvements in lipase production in thelaboratory or in a pilot plant bioreactor by using differentoperational strategies.

References

[1] Jaeger KE, Ransac S, Dijkstra BW, Colson C, van Heuvel M, MissetO. Bacterial lipases. FEMS Microbiol Rev 1994;15:29–63.

[2] Rapp P, Backhaus S. Formation of extracellular lipases by filamen-tous fungi, yeasts and bacteria. Enzyme Microb Technol 1992;14:938–43.

[3] Bjorkling F, Godtfredsen SE, Kirk O. The future impact of industriallipases. Trends Biotechnol 1991;9:360–63.

[4] Malcata FX. Engineering of/with lipases: scope and strategies. In:Malcata FX, editor, Engineering of/with lipases. Dordrecht, Nether-lands: Kluwer Academic Publishers, 1996. p. 1–16.

[5] Gilbert EJ, Drozd JW, Jones CW. Physiological regulation and opti-mization of lipase activity inPseudomonas aeruginosaEF2. J GenMicrobiol 1991;137:2215–21.

Fig. 4. Time course of cell-bound (ƒ), extracellular (Œ), and total (■)lipase activity after glucose addition (10 g/l) to an oleic acid fed-batchculture starved for 6 h.

662 E. Dalmau et al. / Enzyme and Microbial Technology 26 (2000) 657–663

[6] Marcin C, Katz L, Greasham R, Chartrain M. Optimization of lipaseproduction byPseudomonas aeruginosaMB 5001 in batch cultiva-tion. J Ind Microbiol 1993;12:29–34.

[7] Chartrain M, Marcin C, Katz L, et al. Enhancement of lipase produc-tion during fed-batch cultivation ofPseudomonas aeruginosaMB5001. J Ferment Bioeng 1993;76:487–92.

[8] Long K, Ghazali HM, Ariff A, Ampon K, Bucke C. Mycelium-boundlipase from a locally isolated strain ofAspergillus flavuslink: patternand factors involved in its production. J Chem Technol Biotechnol1996;67:157–63.

[9] Pokorny D, Friedrich J, Cimerman A. Effect of nutritional factors onlipase biosynthesis byAspergillus niger. Biotechnol Lett 1994;16:363–66.

[10] Nahas E. Control of lipase production byRhizopus oligosporusundervarious growth conditions. J Gen Microbiol 1988;134:227–33.

[11] Salleh AB, Musani R, Basri M, et al. Extra- and intra-cellular lipasesfrom a thermophilicRhizopus oryzaeand factors affecting their pro-duction. Can J Microbiol 1993;39:978–81.

[12] Rua ML, Dıaz-Maurino T, Ferna´ndez VM, Otero C, Ballesteros A.Purification of characterization of two distinct lipases fromCandidacylindracea. Biochim Biophys Acta 1993;1156:181–89.

[13] Grochulski P, Li Y, Schrag JD, Cygler M. Two conformational statesof Candida rugosalipase. Prot Sci 1994;3:82–91.

[14] Lotti M, Alberghina L. Candida rugosalipase isoenzymes. In: Mal-cata FX, editor. Engineering of/with lipases. Dordrecht, Netherlands:Kluwer Academic Publishers, 1996. p. 115–24.

[15] Lotti M, Grandori R, Fusetti F, et al. Cloning and analysis ofCandidacylindracealipase sequences. Gene 1993;124:45–55.

[16] Chang RC, Chou SJ, Shaw JF. Multiple forms and functions ofCandida rugosalipase. Biotechnol Appl Biochem 1994;19:93–97.

[17] Valero F, Del Rio JL, Poch M, Sola` C. Fermentation behavior oflipase production byCandida rugosagrowing on different mix-tures of glucose and olive oil. J Ferment Bioeng 1991;72:399 –401.

[18] Lotti M, Monticelli S, Montesinos JL, et al. Physiological control on

the expression and secretion ofCandida rugosalipase. Chem PhysLipids 1998;93:143–48.

[19] Gordillo MA, Sanz A, Sa´nchez A, et al. Enhancement ofCandidarugosa lipase production by using a different control fed-batch op-erational strategies. Biotechnol Bioeng 1998;60:156–68.

[20] Obradors N, Montesinos JL, Valero F, Lafuente FJ, Sola´ C. Effects ofdifferent fatty acids in lipase production byCandida rugosa. Bio-technol Lett 1993;15:357–60.

[21] Valero F, Ayats F, Lopez-Santı´n J, Poch M. Lipase production byCandida rugosa: fermentation behavior. Biotechnol Lett 1998;10:741–44.

[22] Charton E, Davies C, Macrae R. Use of specific polyclonal antibodiesto detect heterogeneous lipases fromGeotrichum candidum.BiochimBiophys Acta 1992;112:191–208.

[23] Lowry RR, Tinsley IJ. Rapid colorimetric determination of free fattyacids. J Am Oil Chem Soc 1976;5:470–72.

[24] Christakopoulos P, Tzia C, Kekos D, Macris BJ. Production andcharacterization of extracellular lipase fromCalvatia gigantea. ApplMicrobiol Biotechnol 1992;38:194–97.

[25] Papaparaskevas D, Christakopoulos P, Kekos D, Macris BJ. Optimiz-ing production of extracellular lipase fromRhodotorula glutinis.Biotechnol Lett 1992;14:397–402.

[26] Shimada Y, Sugihara A, Nagao T, Tominaga Y. Induction ofGeotri-chum candidumlipase by long-chain fatty acids. J Fermet Bioeng1992;74:77–80.

[27] Rapp P. Production, regulation, and some properties of lipase activityfrom Fusarium oxysporumf.sp. vasinfectum. Enzyme Microb Tech-nol 1995;17:832–38.

[28] Christova N, Tuleva B, Galabova D. Dependence of yeast permeabi-lization with Triton X-100 on the cell age. Biotechnol Tech 1996;10:77–8.

[29] Baillargeon MW, Bistline RG, Sonnet PE. Evaluation of strains ofGeotrichum candidumfor lipase production and fatty acid specificity.Appl Microbiol Biotechnol 1989;30:92–6.

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