Purification of an organic solvent-tolerant lipase from Aspergillus niger MYA 135 and its application in ester synthesis

Biocatalysis and Agricultural Biotechnology 1 (2012) 25–31

Contents lists available at SciVerse ScienceDirect

Biocatalysis and Agricultural Biotechnology

1878-81

doi:10.1

n Corr

E-m1 Te2 Te

journal homepage: www.elsevier.com/locate/bab

Purification of an organic solvent-tolerant lipase from Aspergillus niger MYA135 and its application in ester synthesis

Cintia M. Romero a, Licia M. Pera a, Flavia Loto a, Cecilia Vallejos b,1,Guillermo Castro c,2, Mario D. Baigori a,n

a Planta Piloto de Procesos Industriales Microbiologicos (PROIMI-CONICET), Av. Belgrano y Pasaje Caseros, T4001MVB Tucuman, Argentinab Departamento de Bioquımica de la Nutricion, Instituto Superior de Investigaciones Biologicas (INSIBIO), Chacabuco 461, 4000 S. M. de Tucuman, Argentinac CINDEFI, Centro de Investigacion y Desarrollo en Fermentaciones Industriales Facultad de Ciencias Exactas (UNLP) 50 y 115 (B1900AJL) La Plata, Buenos Aires, Argentina

a r t i c l e i n f o

Article history:

Received 7 February 2011

Received in revised form

24 May 2011

Accepted 17 August 2011Available online 30 August 2011

Keywords:

Lipase

Aspergillus niger

Purification

Organic solvent-tolerant

81/$ - see front matter & 2011 Elsevier Ltd. A

016/j.bcab.2011.08.013

esponding author. Tel.: þ54 381 43 44 888; f

ail address: [email protected] (M.D. Baigori

l.: þ54 381 4248921.

l./fax: þ54 221 4833794.

a b s t r a c t

An organic solvent-tolerant lipase from olive oil-induced Aspergillus niger MYA 135 supernatant was

purified using two methods: electroelution and ion-exchange chromatography. With electroelution

purification was 8.4-fold and recovery 47% and with ion-exchange 16.6-fold and 53.4%, respectively.

The purified enzyme showed a prominent single band with SDS-PAGE and was a monomeric protein of

68 kDa. The isoelectric point (pI) of the lipase was 5.1 and optimum pH and temperature for activity

were 7.0 and 37 1C, respectively. The lipase showed affinity for esters with long acyl chains, with a Km of

0.99 mM for C18. Substrate specificity of the immobilized lipase was highest for C18 among the various

a- and b-naphthyl esters assayed. Substrate specificity agreed with kinetics parameters of long-chain

fatty acids (C18). Transesterification activity of the A. niger MYA 135 lipase indicates that it could be a

potential biocatalyst for biodiesel production.

& 2011 Elsevier Ltd. All rights reserved.

1. Introduction

Lipases [EC 3.1.1.3] catalyze hydrolysis of triglycerides at oil–water interfaces. This is a reversible reaction and the enzyme alsocatalyzes ester synthesis and transesterification under microaqu-eous conditions. Lipases are presently considered as the tool ofchoice of chemists owing to their ability to catalyze a variety ofsynthetic reactions in non-aqueous environments (Saxena et al.,2003).

Although reactions catalyzed by lipases in the presence oforganic solvents have many advantages (Gotor-Ferndez et al.,2006), most enzymes, including lipases, are not stable in organicsolvents, especially hydrophilic organic solvents. This is due to thetendency of these solvents to strip water molecules of the enzymesurface, which leads to inactivation of the enzyme (Yang et al.,2004). In order to overcome this limitation, several strategies likechemical modification, immobilization and protein engineeringhave been employed to stabilize enzymes for use in organicsolvents (Polizzi et al., 2007). Alternatively, it has been proposed thatrather than modifying enzymes to increase their solvent stability, itwould be more desirable to screen directly for solvent-tolerant

ll rights reserved.

ax: þ54 381 43 44 887.

).

enzymes for their application in non-aqueous enzymatic reactions(Ji et al., 2010).

There exist several reports on the purification and character-ization of solvent-tolerant lipases, classified by the degree ofsolvent tolerance, which have been used in the synthesis of manyuseful products such as biodiesel (Ji et al., 2010; Yang et al., 2007;Wang et al., 2009).

Filamentous fungi are widely recognized and used for theirextracellular lipases, a feature that facilitates enzyme recoveryfrom the broth. A. niger is one of the most important microorgan-isms used in biotechnology and this microorganism producesmany extracellular enzymes that are recognized as GRAS (gen-erally recognized as safe) by the FDA (Mhetras et al., 2009).

Lipases have been purified from several sources using a varietyof methods involving ammonium sulfate precipitation and ionexchange chromatography followed by gel filtration. In recentyears, new techniques have been studied to help reduce thenumber of steps necessary for lipase purification and increase theyield of purified enzymes (Saxena et al., 2003).

In a previous study, we reported the presence of three enzymaticbands on nPAGE with lipolytic activity in olive oil-induced super-natant from A. niger MYA 135 (Pera et al., 2006). In the currentstudy, one of them, lipase I, was selected for purification andcharacterization, because it had shown to be stable in the presenceof various organic solvents. This lipase is significantly different fromthose reported from other Aspergillus strains, and it has shown to be

C.M. Romero et al. / Biocatalysis and Agricultural Biotechnology 1 (2012) 25–3126

potentially useful for biodiesel synthesis due to its excellent solventtolerance.

2. Materials and methods

2.1. Enzyme production and enzyme assaying

A. niger ATCC MYA 135 was obtained from the PROIMI culturecollection. Enzyme production was performed under submergedconditions using a saline medium supplemented with 2% olive oil(Pera et al., 2006). Lipase activity was measured spectrophotome-trically at 405 nm with p-nitrophenyl palmitate (p-NPP) as sub-strate at 37 1C in 100 mM phosphate buffer (pH¼7.0), 0.1% (bymass per volume) gum arabic and 0.4% (by mass per volume)Triton X-100 according to the method by Winkler and Stuckman(1979). One unit of hydrolytic activity is expressed as the amountof enzyme that released 1 mmol of p-nitrophenol per min underthe given assay conditions. The protein concentration was deter-mined according to Bradford (1976) with bovine serum albuminas the standard.

2.2. Lipase stability in organic solvents

Supernatants were run on native polyacrylamide gels. After-wards, each gel was incubated in the corresponding solvent underconstant agitation for 1 h at 37 1C. Residual enzyme activity wasmeasured using a-naphthyl acetate as substrate and comparedwith lipolytic activity of the enzyme in the absence of organicsolvents. The following solvents were assayed: methanol, ethanol,1-propanol, 2-propanol, 1-butanol, 2,3-butanediol, acetone, n-hexane,hexanol and heptane (100%).

The intensity of the dyed bands was analyzed by densitometryusing a Gel Doc 2000 equipment (Bio-Rad) with Quantity Onesoftware.

2.3. First purification technique

The fermented broth was filtered and centrifuged to get a clearsupernatant, which was then concentrated with PEG 20,000 over-night at 4 1C. Samples were recovered in 10 mM phosphate buffer,pH 7. Preparative electrophoresis of an appropriate dilution of thecrude enzyme preparation was run for about 5 h at 50 V. After thegel was run, a slice of about 2 cm was cut vertically along the gel,which was subsequently stained for lipolytic activity and proteinswith a-naphthyl acetate and Coomassie Brilliant Blue according toBradford (1976), respectively. The stained slice was aligned with theremainder of the gel and then each band corresponding to thestained proteins was excised horizontally. Each slice was thenintroduced into a dialysis membrane, previously filled with 10 mM(NH4)2CO3. The membrane (with the sample) was placed in ahorizontal gel electrophoresis equipment also filled with 10 mM(NH4)2CO3. The proteins were eluted from the gel slices at 75 Vduring 3 h and a reverse current for 5 min. The contents of thedialysis membrane were recovered and the gels discarded. The(NH4)2CO3 solution containing the enzyme was vacuum concen-trated. Lipase activity of the purified enzyme was measured with theassay described above in a native polyacrylamide gel. Purity wasconfirmed by migration of a single band.

2.4. Isoelectric focusing

Isoelectric focusing polyacrylamide gel electrophoresis(IEF–PAGE) was performed in a vertical gel equipment using40% ampholytes (Sigma) with a pH range of 3–10, 20 mM aceticacid as anodic solution and 20 mM NaOH as cathodic solution.

The purified protein was loaded onto the gel and focused at 200 Vfor a period of 1.5 h. For detection of protein bands, gels wereincubated previously in 10% trichloroacetic acid (TCA) during30 min for silver staining and in 10 mM phosphate buffer forlipase activity staining.

2.5. Second purification technique

In this technique, the fermented broth was previously dialyzedagainst water with dialysis buffer to reduce salt contents and thenconcentrated as described above until a concentration between1 and 10 mg of protein/ml. The concentrated protein was col-lected in a minimum volume of 50 mM Tris–HCl buffer, pH 7.0.The concentrated sample was loaded onto a DEAE-sepharoseCL-6B column (1.6�20 cm2) (Pharmacia), previously equilibratedwith 50 mM Tris–HCl buffer (pH 7). After washing with two bedvolumes of the initial buffer, elution was performed with agradient of 0–500 mM NaCl at a flow rate of 0.5 ml/min. Fractionsshowing lipase activity were pooled, vacuum concentrated andthen stored at �20 1C.

2.6. Molecular mass determination

The molecular mass of the lipase was determined by SDS-PAGEaccording to Laemmli (1970). Proteins were stained with silver(Deutscher, 1990). The molecular mass was also determined bymatrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) mass spectrophotometry with an Ultraflex II (Bruker), at theCEQUIBIEM mass spectrometry center, Buenos Aires, Argentina.

2.7. Effect of pH and temperature on enzyme activity

The effect of pH on the enzyme activity was assayed at 37 1C ina pH range of 3.0–9.0, using the following buffers (100 mM):citrate–phosphate (pH 3.0 and 5.0), phosphate (from pH 6.0 to8.0) and borate–HCl (pH 9.0). The optimum temperature of theenzyme was determined by measuring the enzyme activity atvarious temperatures (25–45 1C).

The activity was assayed under standard assay conditions(Pera et al., 2006).

2.8. Lipase kinetics

Kinetics constants of the lipase were determined by directregression of the Michaelis–Menten hyperbola obtained experi-mentally. The assays were carried out according to the standardreaction (Pera et al., 2006) using a pure lipase sample and p-NPderivatives with fatty acids of different chain length (C2, C3, C16and C18) in concentrations that varied from 0.1 to 2.0 mM.The apparent kinetics parameters Km and Vmax were calculatedfor each experiment by non-linear regression analysis, whicheffectively fits the data directly to the best hyperbola. GraphPadPrism version 4.00 for Windows, GraphPad Software, San DiegoCalifornia USA, was used.

2.9. Immobilization of the lipase I

Immobilization was carried out by entrapping the lipase in apolyacrylamide gel. Concentrated supernatant with lipase I wasloaded onto a native polyacrylamide gel and electrophoresis wascarried out. After the gel was run, a slice of about 2 cm was cutvertically along the gel and lipase I was detected using a-naphthylacetate as substrate. The stained slice was aligned with theremainder of the gel and then the band corresponding to the stainedprotein was excised horizontally. Slices were subsequently intro-duced into 100 mM phosphate buffer, pH 7.0.

C.M. Romero et al. / Biocatalysis and Agricultural Biotechnology 1 (2012) 25–31 27

The residual hydrolytic activity of the immobilized enzymewas tested using p-NPP as substrate. Thus, the immobilized lipaseI retained 36% (0.37 U/L) of the concentrated supernatant lipaseactivity (1.04 U/L).

2.10. Effect of organic solvents on immobilized lipase I

The effect of various organic solvents with different log P value(Table 1) on the immobilized lipase I stability was also investi-gated. To 1 ml of each organic solvent, a gel slice containing thelipase I was added. The system was incubated in a shake tube(150 rpm) for 1 h at 37 1C, and the residual hydrolytic activity wasthen quantified using p-NPP as substrate. For control distilledwater was added instead of solvent.

2.11. Chain length and positional specificity of immobilized lipase I

Substrate preference of immobilized lipase was determined bymeasuring naphthol formation from a- and b-naphthyl esters(1.3 mM): acetate (C2), myristate (C14), palmitate (C16) andstearate (C18). Lipase activity was assayed spectrophotometri-cally at 560 nm. Released naphthol was bound to 1 mM Fast Bluein 100 mM phosphate buffer (pH 7.0), to give a colored product.The results are expressed as the percentage of the relative lipaseactivity (%) measured with a- and b-naphthyl esters consideringthe preferred substrate (highest enzyme activity) as 100% (Baigorıet al., 1996).

2.12. Production of biodiesel catalyzed by lipase I

Enzymatic solvent-free transesterification was carried out byaddition of 100 ml of 20 mM b-naphthyl stearate (acyl donor) to900 ml of ethanol, serving as reaction medium and acyl acceptor.Immobilized lipase I entrapped in a polyacrylamide gel was usedas biocatalyst. The reaction mixture was shaken (150 rpm) for 2 hat 37 1C and the naphthol concentration in the supernatant wasmeasured as above. A reaction mixture with gel slices withoutenzyme was used as hydrolysis control. In the absence of theenzyme, no reaction was observed. One unit of transesterificationactivity is defined as the amount of biocatalyst that released 1 mmolof naphthol per min. The operational stability of lipase I wasevaluated following repeated cycles of transesterification. Eachreaction was carried out for 2 h at 37 1C as previously described.Between two consecutive catalytic cycles, the immobilized enzyme

Table 1Densitometry analysis expressed as the percentage of intensity (%) of the dyed

bands corresponding to residual activity using a-naphthyl acetate as substrate.

Three lipolytic enzymes from olive oil-induced supernatant from A. niger MYA 135

were assayed for activity after treatment in different organic solvents. Log P values

of the solvents used are shown.

Solvent Intensity of the dyed band (%)a

Log P Band I Band II Band III

Control 40.5 32.8 26.7Methanol �0.77 38.5 34.1 27.4Ethanol �0.31 36.5 33.2 30.2Acetone �0.20 38.4 32.7 28.91-Propanol 0.28 46.5 39.5 13.92-Propanol 0.38 38.0 32.7 29.22.3-Butanediol 0.35 39.2 31.0 29.8Butanol 0.80 – – –Hexanol 2.03 100 – –n-Hexane 3.60 60.2 – 39.7Heptane 4.32 58.1 – 41.9

a The intensity of the dyed band (%) was calculated for each of the bands I, II

and III (the sum of the three bands was 100%).

was washed exhaustingly with water and the residual enzymeactivity was determined based on the transesterification assay, asdescribed before.

3. Results and discussion

3.1. Lipase stability in organic solvents

In a former study, we observed that lipase activity in super-natant from A. niger MYA 135 was almost unaffected in thepresence of 50% water-miscible organic solvents. Water-immis-cible aliphatic solvents reduced the lipase activity between 20%and 80% (Pera et al., 2006).

To observe the behavior of the three lipolytic supernatantenzymes, their stability in organic solvents was studied withnative polyacrylamide gel electrophoresis. Previously, three activ-ity bands were observed with an apparent MW of 104.3 kDa,77.4 kDa and 73.3 kDa for band I, II and III, respectively (Peraet al., 2006). Densitometry analysis revealed that all three bandspresent in the supernatant were stable in water-miscible solvents(Table 1). These results are similar to those reported previouslyfor a lipase from Serratia marcense (Zhao et al., 2008). Enzymestability in hydrophilic solvents is especially interesting, becausethis property could be useful in the production of lipase-catalyzedbiodiesel; methanol or ethanol tolerance is a key factor in thisprocess (Nie et al., 2006). Although hexanol and n-butanol arepolar solvents, they present low water solubility. This couldexplain the fact that only band I was active in hexanol and noneof them in n-butanol (Table 1). Band I showed interesting activityin heptane and n-hexane, two non-polar solvents of low watersolubility (Table 1). Enzymes I and III were able to preserveactivity due to the hydrophobicity of these solvents. In general, ahydrophobic solvent does not capture enzyme-bound watermolecules. Water is necessary for the active conformation of theenzyme, because it allows the enzyme to acquire a suitable three-dimensional conformation for catalysis (Zaks and Klibanov, 1988).A lipase from Bacillus sp. 42 showed an increase in activity of 100%with different non-polar solvents compared with the control(Eltaweel et al., 2005) and a lipase from Pseudomonas sp. showedactivity in the presence of n-hexane (Rahman et al., 2006). Band Ipresented residual activity in all the solvents assayed, except forn-butanol. Densitometry analysis revealed that band I showedhighest intensity after treatment in organic solvents with low andhigh log P values (Table 1). In this connection, the effect of organicsolvents on immobilized lipase I was also evaluated using p-NPPas substrate. As shown in Fig. 1, high enzyme stability wasobserved in the presence of water miscible organic solvents aswell, since it retained almost 100% of its activity after exposurefor 1 h at 37 1C in 100% methanol or ethanol. Interestingly,enzyme preincubation in the presence of n-hexane causes anincrease of the initial hydrolytic activity of 38%.

3.2. Purification of lipase I

Because lipase I showed to be tolerant to organic solvents, it wasfurther analyzed. This extracellular lipase was first purified byelectrophoretic elution. The supernatant was concentrated withPEG 20,000 and loaded onto a native polyacrylamide gel. Afterelectrophoresis the activity band was cut out and electroeluted. Theeluted enzyme was visualized with lipase activity staining of the geland showed as a single band (Fig. 2a). Subsequently, the band wasinoculated on a second polyacrylamide gel with pH gradient. Asingle band was visualized with silver staining and the pI was foundto be 5.1 (Fig. 2b). The result of SDS-PAGE of the purified A. niger

MYA 135 lipase is shown in Fig. 2c. The lipase was homogeneous

0

20

40

60

80

100

120

140

160

Contro

l

Meth

anol

Ethano

l

Aceton

e

1-Pro

pano

l

2-Pro

pano

l

2.3-B

utano

diol

Butano

l

Hexan

ol

n-Hex

ane

n-Hep

tane

Res

idua

l Act

ivity

(%

)

Fig. 1. Residual hydrolytic activity of immobilized lipase I from A. niger MYA 135 after preincubation in 100% organic solvents using p-nitrophenyl palmitate as substrate.

Remaining activity was compared with the control incubated in the presence of distilled water. Error bars represent the standard deviation calculated from at least three

independent experiments.

116.2 kDa

97.4 kDa

66.2 kDa

45 kDa

31 kDa

21.5 kDa

Fig. 2. Purified lipase I from A. niger MYA 135 by electroelution. (a) Single band

recovered after electroelution. A homogeneous band was observed for activity

with a-naphthyl acetate. (b) A single band was observed after isoelectric focusing

and silver staining. (c) SDS-PAGE of the purified lipase. (d) SDS-PAGE molecular

weight standards (Bio-Rad).

C.M. Romero et al. / Biocatalysis and Agricultural Biotechnology 1 (2012) 25–3128

and the molecular mass was estimated at 65 kDa. MALDI-TOF/TOFanalysis of the enzyme activity band electroeluted from the gelrevealed a molecular mass of 68 kDa, which was close to the valueestimated with SDS-PAGE. The molecular mass of most fungal lipasesis in the range of 25–65 kDa. Several purified Aspergillus lipases aresummarized in Table 2. The lipase isolated in the current studyseems to be related to the category with the highest molecular mass.

The purified protein showed to be a monomer with an acid pIof 5.1, which agrees with other findings as most lipases fromAspergillus strains exhibit acidic pI (Table 2).

With the electroelution technique the lipase was purified8.4 times with a recovery of 47%, which is a high recovery for asingle step of purification (Table 3).

To improve the recovery after purification, DEAE-Sepharoseanion-exchange chromatography was assayed (Table 3). Becausethe protein concentration in the supernatant was very low, it madeus assume that lipase I was responsible for most of the proteinconcentration. With this information and that of the acid pI of thelipase, the supernatant was adjusted to pH 7, so the lipase becamenegatively charged to improve bonding to the column. A single bandwas obtained after SDS-PAGE with 16.6-fold purification and 53.4%recovery. Shu et al. (2007) reported a recovery of 33.99% for a lipasefrom A. niger F044 after purification with four successive steps.Mhetras et al. (2009) obtained a recovery of 54% of a lipase from

A. niger NCIM 1207 using several purification steps: precipitationwith ammonium sulfate followed by phenyl sepharose and Sepha-cryl-100 gel chromatography. Our result showed that the similarrecovery was achieved by single step purification.

3.3. Effect of temperature and pH on the lipase activity

The temperature effect on the lipase activity was comparedwith that measured at 37 1C (100%; optimum temperature). Thepurified enzyme was more active at temperatures between 35and 37 1C (Fig. 3a), which is in agreement with data for otherAspergillus lipases like that from Aspergillus carneus, which alsoshowed optimum activity at 37 1C (Table 2). The pH effect on thelipase activity is relative to that measured at pH 7.0 (100%;optimum pH) (Fig. 3b). The optimum is similar to that observedin lipases from A. oryzae and A. niger F044 (Table 2).

3.4. Enzyme kinetics

The effect of the substrate concentration on the reaction rate ofthe purified lipase was assayed using fatty acid esters (p-NPderivatives) with different chain length (Table 4). The enzymeshowed the highest Vmax with short-chain fatty acid esters (p-NPpropionate; C3), corresponding to 0.51 mmol/min, and a lowest Km

of 0.99 mM for p-NP stearate (C18). This low Km value indicates agreat affinity of the enzyme for this ester. Several lipases have beenpurified from A. niger, but substrate affinity varied greatly. Lip1 andLip2 lipases from A. niger CICC 4009 showed affinity for p-NP caprate(C10) and p-NP caprylate (C8), respectively, and the optimalsubstrate for Lip2 showed a Km of 6 mM (Yang et al., 2010). A lipasefrom A. niger F044 showed Km values of 7.37 mM with p-NPP (C16)(Shu et al., 2007). Lipase from A. niger MYA 135 showed the highestaffinity toward long-chain fatty acid esters (C18).

3.5. Substrate specificity of the immobilized enzyme

The purified lipase was immobilized by entrapment in apolyacrylamide gel and substrate specificity was examined.Lipase I was able to hydrolyze substrates with acyl chain lengthsof C2, C14, C16 and C18 bound in the a- and b-position. Highestrelative activity was observed for C18 chains with preference forthe acyl chain bound in the b-position (p¼0.0002) (Fig. 4).Consequently, enzyme specificity was higher toward long-chainsaturated fatty acid esters than to short- or middle-chain fatty

Table 2Optimun temperature, optimum pH, pI and molecular mass of lipases isolated from Aspergillus strains.

Microorganism fromAspergillus genus

Optimun temperature (1C) Optimun pH pI Mass molecular (kDa) Reference

A. niger (Amano) 30 4.1 Not reported 35 (Sugihara et al., 1988)

A. niger Not reported acid Not reported Not reported (Torossian and Bella 1991)

A. oryzae 30 7 Not reported 39 (Toida et al., 1995)

A. niger 25 6.5–7.0 Not reported Not reported (Hatzinikolaou et al., 1996)

A. niger strain MZKI A116 Not reported Not reported 4.1 43 (Pokorny et al., 1997)

A. niger strain MZKI A116 Not reported Not reported 4.2 65 (Pokorny et al., 1997)

A. terreus Not reported Not reported Not reported 41 (Yadav et al., 1998)

A. oryzae RIB128 40 5.5 Not reported 29 (Toida et al., 1998)

A. niger 35–55 5.0–6.0 4.4 35.5 (Namboodiri and Chattopadhyaya 2000)

A. nidulans 40 6.5 4.85 29 (Mayordomo et al., 2000)

A. carneus 37 9.0 4.3 27 (Saxena et al., 2003)

A. niger from Fluka Ltd Not reported Not reported Not reported 31 (Fernandez-Lorente et al., 2005)

A. niger from Fluka Ltd Not reported Not reported Not reported 43 (Fernandez-Lorente et al., 2005)

A. niger from Fluka Ltd Not reported Not reported Not reported 65 (Fernandez-Lorente et al., 2005)

A. niger NRRL3 60 7.2 Not reported Not reported (Adham and Ahmed 2007)

A. niger F044 45 7.0 Not reported 35–40 (Shu et al., 2007)

A. niger NCIM 1207 50 2.5 8.5 32.2 (Mhetras et al., 2009)

A. niger MYA 135 37 7 5.1 68 This article

Table 3Purification of lipase I from A. niger MYA 135.

Purificationtechnique

Total activity(U)

Total protein(mg)

Specific activity(U/mg)

Purification(fold)

Recovery(%)

Supernatant 59.29 14.21 4.17 1.0 100.0Electro-elution 27.87 0.79 35.12 8.4 47.0DEAE-sepharose 31.64 0.46 69.28 16.6 53.4

0

20

40

60

80

100

120

25

Temperature (ºC)

Rel

ativ

e ac

tivity

(%

)

0

20

40

60

80

100

120

3

pH

Rel

ativ

e ac

tivity

(%

)

30 35 37 40 45

5 6 6.5 7 8 9

Fig. 3. (a) Effect of temperature on lipase I activity from A. niger MYA 135. The

activities are shown as relative values of that measured at 37 1C (100%). (b) Effect of

pH on lipase I activity. The activities are shown as relative values of that measured at

pH 7.0 (100%). p-Nitrophenyl palmitate was used as substrate. Error bars represent

the standard deviation calculated from at least three independent experiments.

Table 4Kinetics of A. niger MYA 135 lipase I activity with different p-nitrophenyl esters

as substrate. p-NPA (p-nitrophenyl acetate), p-NPPr (p-nitrophenyl propionate),

p-NPP (p-nitrophenyl palmitate) and p-NPS (p-nitrophenyl stearate). Standard

deviations were calculated from at least three independent experiments.

Substrate Lipase I

Km (mM) Vmax

(lmol/ml/min)

p-NPA (C2) 2.0970.23 0.3770.06

p-NPPr (C3) 1.7470.14 0.5170.07

p-NPP (C16) 2.8170.27 0.02570.004

p-NPS (C18) 0.9970.0.10 0.01070.001

C.M. Romero et al. / Biocatalysis and Agricultural Biotechnology 1 (2012) 25–31 29

acid esters. A different behavior was observed for other purifiedAspergillus lipases like that from A. oryzae RIB 128. This enzymeshowed higher specificity toward substrates of middle-chainsaturated fatty acids than to short- or long-chain fatty acids

(Toida et al., 1998). A lipase from A. nidulans catalyzed moreefficiently hydrolysis of esters of short- and middle-chain fattyacids, with relative hydrolysis rates of 100%, 76%, 29%, 11% and 3%for C5:0, C8:0, C12:0, C16:0 and C18:0, respectively (Mayordomoet al., 2000). A lipase from A. niger MZKI Al 16 demonstratedhigher activity toward short chains (C2–C4) (Hatzinikolaou et al.,1996). Activity of a lipase from A. carneus with p-nitrophenylderivatives as substrates showed enzyme preference for lauricacid (12:0) (Saxena et al., 2003) and finally, a A. niger (AmanoPharmaceuticals Co.) lipase showed specificity toward middle-chain fatty acids (C8–C10) (Sugihara et al., 1988).

The substrate specificity observed in lipase from A. niger MYA135 agrees with kinetics parameters of long-chain fatty acids(C18) and to our knowledge this is the most selective lipasetoward long-chain fatty acids that has been reported so far.

3.6. Application of the lipase in transesterification: biodiesel

production

Considering that the immobilized lipase was stable in ethanol andthat the active site seemed to accommodate better for long-chain

0

20

40

60

80

100

120

Rel

ativ

e ac

tivity

(%

)

α-nap

hthyl

aceta

te

α-nap

hthyl

myrist

ate

α-nap

hthyl

palm

itate

α-nap

hthyl

steara

te

β-nap

hthyl

aceta

te

β-nap

hthyl

myrist

ate

β-nap

hthyl

palm

itate

β-nap

hthyl

steara

te

Fig. 4. Substrate specificity of lipase I from A. niger MYA 135 toward different a- and b-naphthyl esters. The activities are shown as relative values of the maximum activity

measured with b-naphthyl stearate (100%). Error bars represent the standard deviation calculated from at least three independent experiments.

Table 5Retention of transesterification activity by immobilized

lipase I from A. niger MYA 135 in continuous cycles.

Cycles Transesterification

activity (U/L)

Residual

activity (%)

1 1.0170.2 100

2 0.6570.02 64.2

3 0.3170.05 31.0

4 Without activity 0

C.M. Romero et al. / Biocatalysis and Agricultural Biotechnology 1 (2012) 25–3130

substrates, it was thought that the lipase would be suitable forbiodiesel production. Solvent-free transesterification with ethanolwas performed to reduce the influence of the solvent on theimmobilized lipase. In this case ethanol was the acyl acceptor andreaction solvent at the same time. Ethanol is also preferred tomethanol in the transesterification process because it is derivedfrom agricultural products and it is renewable and biologicallyless objectionable regarding the environment (Demirbas, 2006;Chaibakhsh et al., 2009).

Lipase I showed affinity and specificity for long-chain fattyacids (C18), so b-naphthyl steareate was used as acyl donor. Thelipase was able to produce biodiesel with a transesterification rateof 0.7870.01 U/L. In addition, the TLC profile of the syntheticreaction shows a new spot that could correspond to the ethylester production (data not shown) (Romero et al., 2007). On theother hand, the reusability of the immobilized enzyme wasshown in Table 5. After 1 cycle of consecutive operation, theresidual activity remained higher than 60% of the original.

4. Conclusion

In this study, an organic solvent-stable lipase from A. niger MYA135 was 16.6 times purified with an overall yield of 53.4%. Thelipase is a single polypeptide with a molecular weight of 68 kDa anda pI of 5.1. The immobilized lipase was able to catalyze biodieselsynthesis in a solvent-free transesterification reaction. These resultsmake the solvent-tolerant lipase even more potentially valuable forbiotechnological applications in non-aqueous catalysis.

Acknowledgments

This work was supported by grants from CIUNT 261/D409(UNT), PICT 14-32491 (ANCyT) and PIP 297 (CONICET).

References

Adham, N.Z., Ahmed, E.M., 2007. Extracellular lipase of Aspergillus niger NRRL3;production, partial purification and properties. Indian Journal of Microbiology49, 77–83.

Baigorı, M.D., et al., 1996. Purification and characterization of an intracellularesterase from Bacillus subtilis MIR-16. Biotechnology and Applied Biochemistry24, 7–11.

Bradford, M.M., 1976. A rapid and sensitive method for the quantitation ofmicrogram quantities of protein utilizing the principle of protein–dye binding.Analytical Biochemistry 72, 248–254.

Chaibakhsh, N., et al., 2009. Effect of alcohol chain length on the optimumconditions for lipase catalyzed synthesis of adipate esters. Biocatalysis andBiotransformation 27, 303–308.

Demirbas, A., 2006. Biodiesel production via non-catalytic SCF method andbiodiesel fuel characteristics. Energy Conversion and Management 47,2271–2282.

Deutscher, M.P., 1990. Guide to protein purification. Methods in Enzymology 182,430.

Eltaweel, M.A., et al., 2005. An organic solvent-stable lipase from Bacillus sp Strain42. Annals of Microbiology 55, 187–192.

Fernandez-Lorente, G., et al., 2005. Purification of different lipases from Aspergillusniger by using a highly selective adsorption on hydrophobic supports.Biotechnology and Bioengineering 92, 773–779.

Gotor-Ferndez, V., et al., 2006. Lipases: useful biocatalysts for the prepara-tion of pharmaceuticals. Journal of Molecular Catalysis B—Enzymatic 40,111–120.

Hatzinikolaou, D.G., et al., 1996. Production and partial characterization ofextracellular lipase from Aspergillus niger. Biotechnology Letters 18, 547–552.

Ji, Q., et al., 2010. Purification and characterization of an organic solvent-tolerantlipase from Pseudomonas aeruginosa LX1 and its application for biodieselproduction. Journal of Molecular Catalysis B—Enzymatic 66, 264–269.

Laemmli, U.K., 1970. Cleavage of structural proteins during the assembly of thehead of bacteriophage T4. Nature 227, 680–685.

Mayordomo, I., et al., 2000. Isolation, purification and characterization of a cold-active lipase from Aspergillus nidulans. Journal of Agricultural and FoodChemistry 48, 105–109.

Mhetras, N.C., et al., 2009. Purification and characterization of acidic lipase fromAspergillus niger NCIM 1207. Bioresource Technology 100, 1486–1490.

Namboodiri, V.M., Chattopadhyaya, R., 2000. Purification and biochemical char-acterization of a novel thermostable lipase from Aspergillus niger. Lipids 35,495–502.

Nie, K., et al., 2006. Lipase catalyzed methanolysis to produce biodiesel. Optimiza-tion of the biodiesel production. Journal of Molecular Catalysis B—Enzymatic43, 142–147.

Pera, L.M., et al., 2006. Catalytic properties of lipase extracts from Aspergillus niger.Food Technology and Biotechnology 44, 247–252.

Pokorny, D., et al., 1997. Aspergillus niger lipases: induction, isolation andcharacterization of two lipases from a MZKI A116 strain. Journal of MolecularCatalysis B—Enzymatic 2, 215–222.

Polizzi, K., et al., 2007. Stability of enzymes. Current Opinion in Chemical Biology11, 220–225.

Rahman, R.N.Z.R.A., et al., 2006. S5 lipase: an organic solvent tolerant enzyme.Journal of Microbiology 44, 583–590.

Romero, C.M., et al., 2007. Catalytic properties of mycelium-bound lipases fromAspergillus niger MYA 135. Applied Microbiology and Biotechnology 76,861–866.

Saxena, R.K., et al., 2003. Purification strategies for microbial lipases. Journal ofMicrobiological Methods 52, 1–18.

C.M. Romero et al. / Biocatalysis and Agricultural Biotechnology 1 (2012) 25–31 31

Saxena, R.K., et al., 2003. Purification and characterization of an alkaline thermo-stable lipase from Aspergillus carneus. Process Biochemistry 39, 239–247.

Shu, Z.Y., et al., 2007. Purification and characterization of a lipase from Aspergillus

niger F044. Chinese Journal of Biotechnology 23, 96–101.Sugihara, A., et al., 1988. Purification and characterization of Aspergillus niger

Lipase. Agricultural Biology and Chemistry 52, 1591–1592.

Toida, J., et al., 1995. Purification and characterization of a lipase from Aspergillus

oryzae. Bioscience, Biotechnology, and Biochemistry 59, 1199–1203.Toida, J., et al., 1998. Purification and characterization of triacylglycerol lipase

from Aspergillus oryzae. Bioscience, Biotechnology, and Biochemistry 62,759–763.

Torossian, K.K., Bella, W., 1991. Purification and characterization of an acid-resistant triacylglycerol lipase from Aspergillus niger. Biotechnology andApplied Biochemistry 13, 205–211.

Wang, S.L., et al., 2009. Purification and characterization of extracellular lipasesfrom Pseudomonas monteilii TKU009 by the use of soybeans as the substrate.Journal of Industrial Microbiology and Biotechnology 36, 65–73.

Winkler, U.K., Stuckman, M., 1979. Glycogen, hyalunorate, and some otherpolysaccharides greatly enhance the formation of exo-lipase by serratiamarescens. Journal of Bacteriology 138, 663–670.

Yadav, R.P., et al., 1998. Characterization of a regiospecific lipase from Aspergillusterreus. Biotechnology and Applied Biochemistry 28, 243–249.

Yang, J., et al., 2010. lip2, a novel lipase gene cloned from Aspergillus niger exhibitsenzymatic characteristics distinct from its previously identified family mem-ber. Biotechnology Letters 32, 951–956.

Yang, J.K., et al., 2007. Cloning, expression and characterization of a novel thermalstable and short-chain alcohol tolerant lipase from Burkholderia cepacia strainG63. Journal of Molecular Catalysis B—Enzymatic 45, 91–96.

Yang, L., et al., 2004. Hydration of enzyme in non-aqueous media is consistentwith solvent dependence of its activity. Biophysical Journal 87, 812–821.

Zaks, A., Klibanov, A.M., 1988. The effect of water on enzyme action in organicmedia. Journal of Biological Chemistry 263, 8017–8021.

Zhao, L., et al., 2008. Biochemical properties and potential applications of anorganic solvent-tolerant lipase isolated from Serratia marcescens ECU1010.Process Biochemistry 43, 626–633.