Bioresource Technology 97 (2006) 1828–1833

Olive-mill wastewaters: a promising substratefor microbial lipase production

Alessandro D’Annibale, Giovanni Giovannozzi Sermanni,Federico Federici, Maurizio Petruccioli ¤

Dipartimento di Agrobiologia e Agrochimica, University of Tuscia, Via San Camillo De Lellis, 01100 Viterbo, Italy

Received 8 April 2004; received in revised form 16 June 2005; accepted 2 September 2005Available online 19 October 2005

Abstract

The present study investigated the valorization of olive-mill wastewater (OMW) by its use as a possible growth medium for the micro-bial production of extra-cellular lipase. To this end, strains of Geotrichum candidum (NRRL Y-552 and Y-553), Rhizopus arrhizus (NRRL2286 and ISRIM 383), Rhizopus oryzae (NRRL 6431), Aspergillus oryzae (NRRL 1988 and 495), Aspergillus niger (NRRL 334), Candidacylindracea (NRRL Y-17506) and Penicillium citrinum (NRRL 1841 and 3754, ISRIM 118) were screened. All strains were able to growon the undiluted OMW, producing extra-cellular lipase activity. C. cylindracea NRRL Y-17506 showed the highest lipase activity on allthe typologies of OMW used. Its lipase production on OMW was markedly aVected by the type of nitrogen source and was induced bythe addition of olive oil. The highest activity (9.23 IU ml¡1) of the yeast was obtained on OMW supplemented with NH4Cl (2.4 g l¡1) andolive oil (3.0 g l¡1).© 2005 Elsevier Ltd. All rights reserved.

Keywords: Olive-mill wastewater valorization; Fungal screening; Candida cylindracea; Lipase production; Medium optimization

1. Introduction

The three-phase olive-oil extraction process (TPOEP)generates a dark-colored eZuent usually termed olive-millwastewater (OMW). This eZuent is characterized by a highpolluting load and by the presence of compounds, phenols inparticular, with biostatic and phytotoxic activity (Capassoet al., 1995; Casa et al., 2003). With the only exception ofSpain, TPOEP is the most widespread olive oil manufactur-ing process (De Micheli and Bontoux, 1996). Therefore,OMW disposal is a large-scale environmental problem partic-ularly in the Mediterranean area where about 3£107 m3 ofthis eZuent are produced annually (Sayadi and Ellouz, 1995).

Several disposal methods have been proposed andmainly include physico-chemical treatments (decantationwith lime and/or chemical oxidation, concentration, drying

* Corresponding author. Tel.: +39 0761 357332; fax: +39 0761 357242.E-mail address: petrucci@unitus.it (M. Petruccioli).

0960-8524/$ - see front matter © 2005 Elsevier Ltd. All rights reserved.doi:10.1016/j.biortech.2005.09.001

and incineration; ultraWltration and reverse osmosis)(Rozzi and Malpei, 1996) and agronomic uses for ferti-irri-gation purposes (Cereti et al., 2004).

Besides being a serious environmental problem, how-ever, OMW can represent a possible resource for the pres-ence of simple and complex sugars that might be a basis forfermentation processes (Federici et al., 1988; Montedoroet al., 1993; Crognale et al., 2003; Fenice et al., 2003). Inaddition, OMW generally contains variable quantities ofresidual oil, the amount of which mainly depends on theextraction process eYciency. For these reasons, OMWcould be a putative candidate as a potentially suitable liq-uid growth medium for microorganisms able to producelipolytic enzymes (Scioli and Vollaro, 1997). It is notsurprising that lipolytic moulds are among the primarycolonizers of OMW during its storage either in tanks oraerated lagoons (Millan et al., 2000).

The aim of the present study was to assess OMW suit-ability as a growth medium for the fermentative productionof microbial enzymes and lipase (glycerol ester hydrolase

A. D'Annibale et al. / Bioresource Technology 97 (2006) 1828–1833 1829

E.C. 3.1.1.3) was taken as the reference model enzyme. Infact, this enzyme, which is characterized by a wide andincreasing number of applications (Jaeger and Eggert, 2002;Murty et al., 2002), has long been commercially producedthrough well-established fermentative processes. To thisend, 12 fungal strains belonging to well-known lipolyticspecies, such as Aspergillus oryzae, Aspergillus niger, Can-dida cylindracea, Geotrichum candidum, Penicillium citrinum,Rhizopus arrhizus and Rhizopus oryzae were preliminarilyscreened for their ability to grow on undiluted OMW and toproduce extra-cellular lipase activity. Among them, the mostpromising strain, i.e. C. cylindracea NRRL Y-17506, wasthen studied as the model organism and its lipase produc-tion further investigated in OMW-based medium.

2. Methods

2.1. Olive-mill wastewaters (OMW) and microorganisms

OMW samples, the composition of which is shown inTable 1, were withdrawn from four olive oil extractionplants that used either a conventional or modiWed three-phase extraction technology. Unless otherwise indicated,OMW was centrifuged (3800g, 20 min) and supplementedwith yeast extract, 0.5 g l¡1 and (NH4)2SO4, 1.0 g l¡1. Beforesterilization (121 °C for 20 min), the pH of OMW-basedmedia was adjusted to 6.1 with 1.0 M NaOH.

The fungal strains used in the present study wereobtained from the Culture Collections of the NorthRegional Research Laboratory (NRRL), Peoria, Illinois,USA (A. oryzae NRRL 1988 and NRRL 495, A. nigerNRRL 334, C. cylindracea NRRL Y-17506, G. candidumNRRL Y-552 and NRRL Y-553, P. citrinum NRRL 1841and NRRL 3754, R. arrhizus NRRL 2286 and R. oryzaeNRRL 6431) and from the Istituto Superiore di Ricerca eFormazione sui Materiali Speciali per le Tecnologie Avanz-ate (ISRIM), Terni, Italy (P. citrinum ISRIM 118 and Rhi-zopus sp. ISRIM 383). Stock cultures were maintained onPotato Dextrose Agar (PDA) (Difco, Detroit, Michigan,USA) at 4 °C and subcultured every month.

2.2. Culture conditions

Five-day-old PDA-slant cultures were suspended in 5 mlof sterile deionised water and used as the inoculum for pre-

cultures to obtain an initial cell (spores or yeast cells) con-centration of t1£ 106 ml¡1. Incubations were carried outat 28 °C for 72 h under orbital shaking (180 rpm) in Erlen-meyer Xasks (500 ml) containing 95 ml of 2-fold dilutedOMW-1. Pre-cultures (5 ml) were then used to inoculateErlenmeyer baZed-Xasks (500 ml) containing 95 ml ofOMW-based media which were incubated at 28 °C for168 h on a rotary shaker (180 rpm). Culture samples werewithdrawn on a daily basis starting from the 48th hour offermentation. All experiments were performed in triplicate.

To assess the suitability of OMW as a lipase productionmedium, four diVerent OMW samples were used (Table 1).The eVect of nitrogen was studied by adding to OMW-4 thesame amount of nitrogen (0.63 g l¡1) present as one of thefollowing compounds: NH4Cl, (NH4)2SO4, NaNO3 andurea. The inXuence of oil addition was evaluated by addingto OMW-4 one of the following oils (3.0 g l¡1): olive oil,corn oil and soybean oil.

2.3. Lipase assay

Lipase activity was determined spectrophotometricallyat 35 °C using �-naftilmyristate as a substrate, as describedelsewhere (Versaw et al., 1989). One international unit (IU)of lipase activity was deWned as the amount of enzyme pro-ducing 1�mol product min¡1 under the assay conditions.

2.4. Analytical methods

Microbial biomass concentration was measured by dryweight estimation: broth samples were Wltered through pre-weighed Whatman GF/C discs (diameter, 47 mm), the har-vested biomass was washed twice with distilled water andthe Wlter was dried at 105 °C for 24 h, cooled in a desiccatorand weighed. Chemical oxygen demand (COD), total phe-nols and total sugars were determined as described in thestandard methods for the examination of water and waste-water (APHA-AWWA-WPCE, 1995). Lipids were deter-mined gravimetrically after petroleum ether extraction(IRSA, 1985).

2.5. Statistical analysis

One way analysis of variance (ANOVA) and pairwisemultiple comparison procedures (Tukey test) were carried

Table 1Characteristics of the OMW used in the present study¤

¤ Data are means § standard deviations of three determinations. Column data followed by the same superscript letter were not signiWcantly diVerent(P 6 0.05; by Tukey test).¤¤ OMW-1, -3 and -4 came from a traditional three-phase extraction system; OMW-2 came from a modiWed three-phase system with low-water consump-

tion. With the exception of OMW-4, that was withdrawn from storage tanks, the other samples were taken from the decanter during the manufacturingprocess.

OMW¤¤ (no.) COD (g l¡1) Total sugars (g l¡1) COD/total sugars Total phenols (g l¡1) Lipids (g l¡1) pH

1 43.0 § 1.3a 17.4§ 0.5a 2.47 § 0.07a 2.52 § 0.11a 0.75§ 0.05a 5.042 51.5 § 2.1b 25.1§ 1.4b 2.05 § 0.09b 2.82 § 0.15b 0.81§ 0.04a 4.883 41.4 § 2.2a 13.2§ 0.4c 3.13 § 0.12c 2.77 § 0.08b 0.73§ 0.05a 4.924 25.6 § 1.1c 8.8 § 0.5d 2.90 § 0.13d 1.85 § 0.10c 0.40§ 0.03b 4.82

1830 A. D'Annibale et al. / Bioresource Technology 97 (2006) 1828–1833

out using the statistical software SigmaStat, version 2.0(Jandel Corp., San Rafael, CA, USA).

3. Results

3.1. Screening for lipase activity on OMW

Most of the microbial strains employed in this study wereselected on the basis of literature information (Baillargeonet al., 1989; Robert et al., 1989; Sztajer and Maliszewska,1989; Shaw et al., 1989; Pokorny et al., 1994; Macris et al.,1996; Kamini et al., 1998; Miranda et al., 1999; Elibol andOzer, 2000). P. citrinum ISRIM 118 was isolated from a soilthat underwent OMW land spreading while Rhizopus sp.ISRIM 383 was isolated from OMW storage tanks.

OMW-1 was used as the growth medium for the prelimi-nary screening; results are summarized in Table 2. All thestrains grew on OMW-1 and released a lipase activityhigher than 0.30 IU ml¡1. The highest lipase activities wereobtained with C. cylindracea NRRL Y-17506 and G. candi-dum NRRL Y-553 (0.46 and 0.52 IU ml¡1, respectively)after 168 h of fermentation. P. citrinum NRRL 3754 andISRIM 118 produced appreciable activity levels (0.34 and0.38 IU ml¡1, respectively), which were obtained in only72 h thus having volumetric productivities equal to 4.58 and5.42 IU l¡1 h¡1, respectively (Table 2).

3.2. EVect of diVerent OMW

The four selected strains indicated above were grown onthe four OMW samples which diVered signiWcantly in theirtotal sugar concentration and COD/total sugars ratio(Table 1). Fig. 1A shows the maximal levels of lipase activ-ity reached by the selected strains in the four OMW sam-ples. All the strains produced similar levels of lipase activityregardless of the OMW typology. The only exception wasC. cylindracea NRRL Y-17506 which released the highestlevels of lipase activity in OMW-3 and OMW-4 (0.82 and0.94 IU ml¡1, respectively) (Fig. 1A); these OMWs werecharacterized by the lowest initial content of total sugars

(13.2 and 8.8 g l¡1, respectively) and highest COD/totalsugar ratios (3.13 and 2.90, respectively).

In contrast, in the same OMW samples, C. cylindraceaNRRL Y-17506 exhibited the lowest growth (2.98 and2.70 g l¡1 in OMW-3 and OMW-4, respectively) (Fig. 1B).Under the experimental conditions tested and regardless of

Fig. 1. EVect of diVerent OMW typologies (see also Table 1) on lipase (A)and biomass (B) production by P. citrinum NRRL 3454 and ISRIM 118,G. candidum NRRL Y-553 and C. cylindracea NRRL Y-17506. Valuesrepresent means of three independent experiments and error bars indicatestandard deviations. For each strain, the same letter above bars indicatesthe absence of signiWcant diVerences (P < 0.01; Tukey test).

OMW-1 OMW-2 OMW-3 OMW-4

Bio

mas

s (g

l-1)

0

1

2

3

4

5

6

7

b

a

a

c c

a

a

b

d

b

c

bb

b

a

bLi

pase

act

ivity

(IU

ml-1

)

0.0

0.2

0.4

0.6

0.8

1.0NRRL 3754 ISRIM 118NRRL Y-17506NRRL Y-553

a aaa

aab

a

b

aba

abb

b

b

a

b A

B

Table 2Growth, lipase production and volumetric productivity (LP) by fungal strains cultured on OMW-1¤

¤ Values are given at the time (Tmax) of maximal enzyme production and are means of three independent experiments § standard deviation. Lipase activ-ity and LP followed by the same superscript letter were not signiWcantly diVerent (P 6 0.05; by Tukey test).

Strain Tmax (h) Biomass (g l¡1) Lipase (IU ml¡1) LP (IU l¡1 h¡1)

Penicillium citrinum NRRL 1841 120 5.88 § 0.02 0.37 § 0.02a 2.92 § 0.16a

Penicillium citrinum NRRL 3754 72 5.34 § 0.01 0.34 § 0.04ac 4.58 § 0.54b

Penicillium citrinum ISRIM 118 72 5.71 § 0.18 0.38 § 0.07a 5.42 § 1.00c

Aspergillus niger NRRL 334 168 7.07 § 0.63 0.33 § 0.01ac 2.08 § 0.06de

Aspergillus oryzae NRRL 485 168 5.39 § 0.42 0.38 § 0.02a 2.08 § 0.11de

Aspergillus oryzae NRRL 1988 168 5.73 § 0.18 0.34 § 0.03ac 2.08 § 0.18de

Candida cylindracea NRRL Y-17506 168 3.65 § 0.07 0.46 § 0.01b 2.92 § 0.06a

Geotrichum candidum NRRL Y-552 96 4.06 § 0.23 0.36 § 0.07ac 3.75 § 0.73f

Geotrichum candidum NRRL Y-553 168 3.16 § 0.05 0.52 § 0.01b 2.92 § 0.06a

Rhizopus arrhizus NRRL 2286 168 3.07 § 0.15 0.30 § 0.02c 1.67 § 0.11d

Rhizopus oryzae NRRL 6431 144 3.67 § 0.15 0.36 § 0.05ac 2.50 § 0.35ae

Rhizopus sp. ISRIM 383 120 4.22 § 0.16 0.35 § 0.03ac 2.92 § 0.25a

A. D'Annibale et al. / Bioresource Technology 97 (2006) 1828–1833 1831

the OMW typology, cell growth and lipase production bythis strain appeared to be negatively correlated (adjustedR2D0.91, PD0.03).

The best strain/OMW substrate combination appearedto be C. cylindracea NRRL Y-17506 on OMW-4. For thisreason, all subsequent experiments aimed at further study-ing the suitability of using OMW for lipase productionwere performed using that combination.

3.3. EVect of nitrogen source

Four diVerent nitrogen sources, including NH4Cl,(NH4)2SO4, NaNO3 and urea, were evaluated for their abil-ity to support lipase production by C. cylindracea NRRLY-17506 on OMW-4 at 0.63 g nitrogen l¡1.

Lipase production was signiWcantly favored by bothammonium salts, reaching 1.24 and 1.45 IU ml¡1 on(NH4)2SO4 and NH4Cl, respectively (Fig. 2). The activitieson media containing either urea or NaNO3 were signiW-cantly lower than those obtained with ammonium salts(0.32 and 0.65 IU ml¡1, respectively). In contrast, slightdiVerences in biomass production were observed by usingdiVerent nitrogen sources (Fig. 2).

3.4. EVect of oil addition

Three common oils, namely olive oil, corn oil and soy-bean oil, were added (3.0 g l¡1) to OMW-4 and their even-tual inducing eVect evaluated. Lipase production increasedupon oil addition (Fig. 3). The most eVective inducer wasolive oil which led to a 7.5-fold increase in activity peakwith respect to the OMW-based control medium (9.0 vs1.24 IU ml¡1). However, the addition of either corn or soy-bean oil led to an approximately 5-fold increase in the max-imal lipase production (6.42 and 6.34 IU ml¡1, respectively).

In all cases, oil addition resulted in an increase in cellbiomass (Fig. 3).

Fig. 2. EVect of nitrogen source on biomass and lipase production byC. cylindracea NRRL Y-17506 on OMW-4. Values represent means of threeindependent experiments and error bars indicate standard deviations. Thesame letter above bars with the same color indicates the absence of signiW-cant diVerences (P < 0.01; Tukey test).

Nitrogen source

Lipa

se a

ctiv

ity(I

U m

l-1)

0.0

0.2

0.4

0.6

0.8

1.0

1.2

1.4

1.6

1.8

Bio

mas

s(g

l-1)

0

1

2

3

4

5

(NH4)2SO4 NH4Cl NaNO3 Urea

a

ab

a

a

b

b

b

c

3.5. Time course of C. cylindracea lipase activity

The time course of growth and lipase production by C.cylindracea NRRL Y-17506 cultured on OMW-4 supple-mented with NH4Cl (2.4 g l¡1) and olive oil (3.0 g l¡1) is

Fig. 3. EVect of oil addition on biomass and lipase production byC. cylindracea NRRL Y-17506 on OMW-4. Values represent means of threeindependent experiments and error bars indicate standard deviations. Thesame letter above bars with the same color indicates the absence of signiW-cant diVerences (P < 0.01; Tukey test).

AdditionNone Olive oil Corn oil Soybean oil

Lipa

se a

ctiv

ity (

IU m

l-1)

0

2

4

6

8

10

12

Bio

mas

s (g

l-1)

0

1

2

3

4

5

6

a

a

bb

cc

b b

Fig. 4. Time course of growth (�) and lipase production (�) by C. cylindr-acea NRRL Y-17506 cultured on OMW-4 supplemented with NH4Cl(2.4 g l¡1) and olive oil (3.0 g l¡1). Residual lipids (�), total sugar (�) andphenol (�) concentrations and COD (�) are also reported. Values repre-sent means of three independent experiments and error bars indicate stan-dard deviations.

Time (h)0 24 48 72 96 120 144 168

Lipi

ds (

mg

l-1)

0

50

100

150

200

250

300

350

400

450C

OD

(g

l-1)

10

12

14

16

18

20

22

24

26

Tot

al p

heno

ls(g

l-1)

1.0

1.2

1.4

1.6

1.8

Lipa

se a

ctiv

ity (

IU m

l-1)

0

1

2

3

4

5

6

7

8

9

10

Tot

al s

ugar

(g

l-1)

0

1

2

3

4

5

6

7

8

9

Bio

mas

s (g

l-1)

0

1

2

3

4

5

6

7

1832 A. D'Annibale et al. / Bioresource Technology 97 (2006) 1828–1833

shown in Fig. 4. The yeast grew rapidly on the wastewaterreaching a biomass concentration of 5.3§0.15 g l¡1 after120 h of fermentation, following a time course similar tothat of the enzyme activity which increased up to9.23§ 0.46 IU ml¡1. Control cultures, grown on tap watercontaining 0.5 g l¡1 yeast extract and supplemented withNH4Cl and olive oil (both at the same concentration asabove), produced a very low lipase activity (0.067 IU ml¡1).

During the fermentation, the COD and total sugar con-tent of OMW decreased to 13.18§ 0.82 and 3.32§ 0.25 g l¡1,respectively (48.4% and 61.3% reduction), while the lipidswere exhausted within 96 h of fermentation. A reduction oftotal phenol content was also observed (36.2% with a Wnalconcentration of 1.18§0.03 g l¡1).

4. Discussion

This study shows that olive-mill wastewater can be avaluable liquid growth medium for the production ofmicrobial lipases. To the best of our knowledge this is theWrst report dealing with the use of OMW for this purpose.Although reliable fermentative processes for lipase produc-tion and eYcient protocols of heterologous expression ofthe enzyme are currently available (Ferrer et al., 2001),there is a growing interest in the search for cheap rawsubstrates. The economics of enzyme production usinginexpensive raw materials can make industrial enzymaticprocesses competitive with chemical ones (Miranda et al.,1999). In most instances, the carbon source has been shownto aVect the Wnal production cost of a bioprocess variably(Arbige and Pitcher, 1989; Ramalingam and Pennathur,2005), depending on down-stream costs (Saxena et al.,2003). In this respect, the use of several industrial by-prod-ucts, such as liquid reWnery wastes (Miranda et al., 1999),potato chips wastewater (Azab, 2000) and olive cake andbagasse (Cordova et al., 1998) was investigated and allowedthe production of signiWcant amounts of lipase.

Although all strains screened in this study were able togrow and to produce signiWcant levels of lipase activity inOMW, C. cylindracea NRRL Y-17506 appeared to be par-ticularly interesting. In fact, besides being the highest lipaseproducer among the strains investigated, it appeared to growand produce the enzyme on OMW samples widely diVeringin their COD, phenols, fats and sugar contents. With regardto the last parameter, the present study showed thatlipase production by C. cylindracea NRRL Y-17506 did notrequire sugar depletion from the medium since at the activitypeak, only 50% of the initial sugar content had beenconsumed. In addition, the enzyme was found to be prefer-entially produced in OMW-based media characterized bylow initial sugar contents and high COD/total sugars ratios.

Several studies report that type and sugar concentrationare important factors aVecting lipase production by Can-dida species; yet, it should be emphasised that resultsreported in the literature in this respect are not unequivo-cal. Lipase production by Candida rugosa was found to becompletely repressed by the presence of simple sugars

(Dalmau et al., 2000) and onset of C. paralipolytica lipaseactivity was shown to require the complete depletion ofsugar from the medium (Macris et al., 1996). Yet, an opti-mal glucose concentration as high as 36 g l¡1 in liquidcultures of C. cylindracea Y-17506 was found to yield17.3 IU ml¡1 (Muralidhar et al., 2001).

With regard to the nitrogen source, several investigatorsreported urea as the optimal one for lipase production inliquid cultures of C. rugosa (Obradors et al., 1993; Benjaminand Pandey, 1996). In addition, a high lipase production byC. cylindracea (i.e. 47.2 IU ml¡1) was obtained in a syntheticmedium containing peptone (3.7 g l¡1) and yeast extract(4.9 g l¡1) as nitrogen sources (Muralidhar et al., 2001). Thisstudy focused the attention on low-cost nitrogen com-pounds and showed that ammonium salts led to betterlipase yields than those obtained with either nitrate or urea.

Lipase production can be either constitutive or inducibleby the presence of their substrates or reaction products(Lotti et al., 1998). as a consequence, the presence of lipidsin growth media may represent a key factor in enzyme pro-duction. in this regard, several authors have reported theuse of vegetable oils either as growth substrates (Valeroet al., 1988; Benjamin and Pandey, 1996; Muralidhar et al.,2001; Wei et al., 2004) or inducers (Hegedus and Khacha-tourians, 1988; Zhang et al., 2003). High levels of lipase pro-duction by C. cylindracea liquid cultures were obtained in acomplex medium containing either 10% (Sokolovska et al.,1998) or 33% olive oil (Muralidhar et al., 2001).

The present study showed that lipase production by C.cylindracea NRRL Y-17506 was highly stimulated by thepresence of vegetable oils and that olive oil was the mosteVective. At concentrations lower than 0.3 g l¡1 the stimu-lating eVect was not evident possibly due to the intrinsicpresence of lipids in OMW (data not shown). Lipase pro-duction was considerably improved by the olive oil addi-tion at a concentration of 3 g l¡1 leading to activity levels of9.23 IU ml¡1. It has been shown that lipase is encoded bytwo diVerent gene groups in C. rugosa, one of which is con-stitutively expressed while the other is induced by fattyacids and repressed by the presence of glucose (Lotti et al.,1998). Fatty acids act at a transcriptional level in C. rugosaand this might explain why induction of C. cylindracealipase requires relatively high oil concentrations.

In conclusion, valorization of OMW by using it as afermentation medium for the production of lipase activitywith lipolytic fungi appears to be possible and promising.The lipolytic activity of C. cylindracea NRRL Y-17506 isfavored by the use of OMW with low sugar contents andwith residual oil concentration higher than 0.3 g l¡1. Theaddition of nitrogen might be required due to its generallylow content in OMW.

Acknowledgements

The research was supported by a grant from I.N.C.A.(Consorzio Interuniversitario per la Chimica dell’Ambi-ente), Project “Agro-Food National Plan”. The authors

A. D'Annibale et al. / Bioresource Technology 97 (2006) 1828–1833 1833

kindly thank Dr. F. Santori and Dr. A.R. Cicalini, IstitutoSuperiore di Ricerca e Formazione sui Materiali Specialiper le Tecnologie Avanzate, Terni, Italy, for the gift of twofungal strains.

References

APHA-AWWA-WPCE, 1995. Standard Methods for Examination ofWater and Wastewater, 19th ed. American Public Health Association,American Water Works Association, Water Environment ControlFederation, Washington, DC, USA.

Arbige, M.V., Pitcher, W.M., 1989. Industrial enzymology: a look towardsthe future. Tibtech 7, 331–356.

Azab, M.S., 2000. Re-use of wastewater of food industries for the produc-tion of fungal biomass and enzymes for the bio-treatment of wastewa-ter. Egyptian J. Biotechnol. 7, 163–179.

Baillargeon, M.W., Bistline, R.G., Sonnet, P.E., 1989. Evaluation of strainof Geotrichum candidum for lipase production and fatty acid speciWcity.Appl. Microbiol. Biotechnol. 30, 92–96.

Benjamin, S., Pandey, A., 1996. Optimization of liquid media for lipaseproduction by Candida rugosa. Bioresour. Technol. 55, 167–170.

Capasso, R., Evidente, A., Schivo, L., Orru, G., Marcialis, M.A., Cristinzio,G., 1995. Antibacterial polyphenols from olive oil mill waste waters. J.Appl. Bacteriol. 79, 393–398.

Casa, R., D’Annibale, A., Pieruccetti, F., Stazi, S.R., Giovannozzi Ser-manni, G., Locascio, B., 2003. Reduction of the phenolic componentsin olive-mill wastewater and its impact on durum wheat (Triticumdurum Desf.) germinability. Chemosphere 50 (8), 959–966.

Cereti, C.F., Rossini, F., Federici, F., Quaratino, D., Vassilev, N., Fenice,M., 2004. Reuse of microbially treated olive mill wastewater as fertiliserfor wheat (Triticum durum Desf.). Bioresour. Technol. 91, 135–140.

Cordova, J., Nemmaoui, M., Ismaili-Alaoui, M., Morin, A., Roussos, S.,Raimbault, M., Benjilali, B., 1998. Lipase production by solid-state fer-mentation of olive cake and sugarcane bagasse. J. Mol. Catal. B 5, 75–78.

Crognale, S., Federici, F., Petruccioli, M., 2003. �-glucan production byBotryospheria rhodina on undiluted olive-mill wastewaters. Biotechnol.Lett. 25, 2013–2015.

Dalmau, E., Montesinos, J.L., Lotti, M., Casas, C., 2000. EVect of diVerentcarbon source on lipase production by Candida rugosa. Enzyme Mic-rob. Technol. 26, 657–663.

De Micheli, M., Bontoux, L., 1996. Survey on current activity on the valo-rization of by-products from the olive oil industry. Final Report ofEuropean Commission DG XII.

Elibol, M., Ozer, D., 2000. Lipase production by immobilised Rhizopusarrhizus. Proc. Biochem. 36, 219–223.

Federici, F., Montedoro, G.F., Servili, M., Petruccioli, M., 1988. Pecticenzyme production by Cryptococcus albidus var. albidus on olive oilvegetation waters enriched with sunXower calathide meal. Biol. Wastes25, 291–301.

Fenice, M., Giovannozzi Sermanni, G., Federici, F., D’Annibale, A., 2003.Submerged and solid-state bioprocesses for laccase and manganese-peroxidase production by Panus tigrinus on olive-mill wastewater-based media. J. Biotechnol. 100, 77–85.

Ferrer, P., Montesinos, J.L., Valero, F., Sola, C., 2001. Production of nativeand recombinant lipases by Candida rugosa: a review. Appl. Biochem.Biotechnol. 95, 221–253.

Hegedus, D.D., Khachatourians, G.G., 1988. Production of an extracellu-lar lipase by Beauveria bassiana. Biotechnol. Lett. 10, 637–642.

IRSA, 1985. Metodi analitici per i fanghi. III. Parametri chimico-Wsici.Quaderni Istituto Ricerca sulle Acque, 64, CNR, Roma.

Jaeger, K.E., Eggert, T., 2002. Lipases for biotechnology. Curr. Opin. Bio-technol. 13, 390–397.

Kamini, N.R., Mala, J.G.S., Puvanakrishnan, R., 1998. Lipase productionfrom Aspergillus niger by solid-state fermentation using gingelly oilcake. Proc. Biochem. 33, 505–511.

Lotti, M., Monticelli, S., Montesinos, J., Brocca, S., Valero, F., Lafuente, J.,1998. Physiological control of the expression and secretion of Candidarugosa lipase. Chem. Phys. Lipids 93, 143–148.

Macris, J.B., Kourentzi, E., Hatzinikolaou, D.G., 1996. Studies on localiza-tion and regulation of lipase production by Aspergillus niger. Proc. Bio-chem. 31, 807–812.

Millan, B., Lucas, R., Robles, A., Garcia, T., de Cienfuegos, G.A., Galvez,A., 2000. A study on the microbiota from olive-mill wastewater(OMW) disposal lagoons with emphasis on Wlamentous fungi and bio-degradative potential. Microbiol. Res. 155, 143–147.

Miranda, O.A., Salgueiro, A.A., Pimentel, M.C.B., Lima Filho, J.L., Melo,E.H.M., Duràn, N., 1999. Lipase production by a Brazilian strain ofPenicillium citrinum using an industrial residue. Bioresour. Technol. 69,145–147.

Montedoro, G., Begliomini, A.L., Servili, M., Petruccioli, M., Federici, F.,1993. Pectinase production from olive vegetation waters and its use inthe mechanical olive oil extraction process to increase oil yield andimprove quality. Ital. J. Food Sci. 4, 355–362.

Muralidhar, R.V., Chirumamila, R.R., Marchant, R., Nigam, P., 2001. Aresponse surface approach for the comparison of lipase production byCandida cylindracea using two diVerent carbon sources. Biochem. Eng.J. 9, 17–23.

Murty, V.R., Bhat, J., Muniswaran, P.K.A., 2002. Hydrolysis of oils byusing immobilized lipase enzyme: a review. Biotechnol. Bioproc. Eng. 7,57–66.

Obradors, N., Montesinos, J.L., Valero, F., Lafuente, F.J., Solà, C., 1993.EVect of diVerent fatty acids in lipase production by Candida rugosa.Biotechnol. Lett. 15, 357–360.

Pokorny, D., Friedrich, J., Cimerman, A., 1994. EVect of nutritional factorson lipase biosynthesis by Aspergillus niger. Biotechnol. Lett. 16, 363–366.

Ramalingam, K., Pennathur, G., 2005. Enhanced lipase production byCandida rugosa using novel cost-eVective substrates. In: Proc. 229thACS National Meeting, San Diego, CA, March 13–17, 2005. AmericanChemical Society, Washington, p. 234.

Robert, P.W., Douglas, B.K., Morris, J.G., James, R., Adams, J.M., 1989.Immobilisation of Candida cylindracea lipase on a new range ofceramic supports. Biotechnol. Tech. 3, 345–348.

Rozzi, A., Malpei, F., 1996. Treatment and disposal of olive mill eZuents.Int. Biodeterior. Biodegr. 38, 135–144.

Saxena, R.K., Sheoran, A., Bhoopander, G., Davidson, W.S., 2003. PuriW-cation strategies for microbial lipases. J. Microbiol. Methods 52, 1–18.

Sayadi, S., Ellouz, R., 1995. Role of lignin peroxidase and manganese per-oxidase from Phanerochaete chrysosporium in the decolorization ofolive mill wastewaters. Appl. Environ. Microbiol. 61, 1098–1103.

Scioli, C., Vollaro, L., 1997. The use of Yarrowia lipolytica to reduce pollu-tion in olive mill wastewaters. Water Res. 31, 2520–2524.

Shaw, J.F., Chang, C.H., Wang, Y.J., 1989. Characterization of three dis-tinct forms of lipolytic enzyme in a commercial Candida lipase prepa-ration. Biotechnol. Lett. 11, 779–784.

Sokolovska, I., Albasi, C., Riba, J.P., Bales, V., 1998. Production of extra-cellular lipase by Candida cylindracea CBS 6330. Bioproc. Eng. 19, 179–186.

Sztajer, H., Maliszewska, I., 1989. The eVect of culture conditions onlipolytic productivity of Penicillium citrinum. Biotechnol. Lett. 11,895–898.

Valero, F., Ayats, F., Lòpez-Santìn, J., Poch, M., 1988. Lipase productionby Candida rugosa. Biotechnol. Lett. 19, 741–744.

Versaw, W.K., Cuppett, S.L., Winters, D.D., Williams, L.E., 1989. Animproved colorimetric assay for bacterial lipase in nonfat dry milk. J.Food Sci. 54, 1557–1558.

Wei, D., Zhang, L.-Y., Song, Q., 2004. Studies on a novel carbon sourceand cosolvent for lipase production by Candida rugosa. J. Ind. Micro-biol. Biotechnol. 31, 133–136.

Zhang, L.Y., Wei, D.Z., Tong, W.Y., 2003. EVective inducers for lipase pro-duction by Candida rugosa. Ann. Microbiol. 53, 499–504.