Enzymatic hydrolysis of olive wastewater for hydroxytyrosol enrichment

Bioresource Technology 102 (2011) 9050–9058

Contents lists available at ScienceDirect

Bioresource Technology journal homepage: www.elsevier.com/locate/biortech

Enzymatic hydrolysis of olive wastewater for hydroxytyrosol enrichment Sonia Khoufi ⇑, Manel Hamza, Sami Sayadi Laboratoire des Bioprocédés Environnementaux, Pôle d’Excellence Régional (PER, AUF), Centre de Biotechnologie de Sfax, Université de Sfax, B.P. 1117, 3018 Sfax, Tunisia

a r t i c l e

i n f o

Article history: Received 27 April 2011 Received in revised form 11 July 2011 Accepted 15 July 2011 Available online 24 July 2011 Keywords: Olive wastewater Enzyme preparation Hydrolysis Hydroxytyrosol recovery

a b s t r a c t Aspergillus niger broth culture on wheat bran was assessed for olive wastewater (OW) hydrolysis in order to release hydroxytyrosol (HT). The enzyme profiles of this culture broth gave essentially (IU/L): 3000 bglucosidase and 100 esterase. Hydrolysis activity of A. niger enzyme preparation was evaluated by using three substrates: raw OW, phenolic fraction extracted from OW by ethyl acetate and its corresponding exhausted fraction. Large amounts of free simple phenolics were released from exhausted fraction and raw OW after enzymatic treatment. HPLC analyses show that HT was the main phenolic compound. One step of ethyl acetate extraction of hydrolysed OW allowed the recovery of 0.8 g of HT per litre of OW. The antioxidant activity of extracts from OW and exhausted fraction, measured by DPPH method, was drastically enhanced after hydrolysis treatment. This study demonstrates that hydrolysed OW is a potential source of bioactive phenolic compounds with promising applications in food and pharmaceutical industries. Ó 2011 Elsevier Ltd. All rights reserved.

1. Introduction Olive wastewater (OW) represents a serious problem with crucial negative impacts on soil and water quality, and thus on agriculture and environment. The high-polluting power of this wastewater is generally associated with the high chemical (COD) and biochemical (BOD) oxygen demand, total solids and the slightly acidic character of OW (Khoufi et al., 2008). Concerning the organic load of OW, phenolics and related compounds were reported to be present at very high concentrations. Indeed, most of the problems associated with OW pollution can be attributed to these phenolic compounds that are rather recalcitrant to biological degradation. In fact, phenolic compounds are responsible for several biological effects, including microtoxicity and phytotoxicity (Khoufi et al., 2008; Mekki et al., 2007). On the other hand, OW phenolic compounds have ideal structural chemistry for free radical-scavenging and metal-chelating properties (Fki et al., 2005), and have been shown to be more effective antioxidants in vitro than vitamins E and C on a molar basis (De Leonardis et al., 2007). The interest in natural antioxidants is increasing thanks to the evidence for the involvement of oxygenderived free radicals in several pathological processes. Epidemiological studies have suggested a relation between the consumption

Abbreviations: BG, b-glucosidase; BHT, butylated hydroxytoluene; DPPH, 1,1diphenyl-2-picrylhydrazyl; EA, ethyl acetate; HT, hydroxytyrosol; OW, olive wastewater; p-NPG, p-nitrophenyl-b-D-glucopyranoside; RS, reducing sugar. ⇑ Corresponding author. Tel./fax: +216 74 874 452. E-mail address: sonia.khoufi@cbs.rnrt.tn (S. Khoufi). 0960-8524/$ - see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.biortech.2011.07.048

of polyphenol-rich food and the prevention of diseases associated with oxidative stress, such as cancers, cardiovascular diseases, inflammations and others (Agalias et al., 2007). According to these notes, OW could represent an interesting and alternative source of biologically active polyphenols. Several researches have evaluated the feasibility and the economic processes for recovering olive phenols from OW (Reis et al., 2006) or solid wastes (Fernandez-Bolanos et al., 2002; Rodríguez et al., 2008). The main systems proposed to recover the phenols from OW are: resin chromatography; selective concentration by ultra-filtration and reverse osmosis; solid–liquid or liquid–liquid extraction with solvents and supercritical fluid extraction (Gonzalez-Munos et al., 2003). In previous investigations, it was reported that among all procedures that were taken for natural antioxidants recovery, liquid–liquid solvent extraction represented a simple and convenient alternative, and it was widely used in pilot-scale production and in ultimate commercial recovery (Reis et al., 2006). Fresh OW is rich in polyphenols especially oleuropein which is the major bioactive compound of this by-product in the early harvest season. The molecule consists of three structural subunits: a polyphenol, namely 4-(2-hydroxyethyl) benzene-1,2-diol which is also known as hydroxytyrosol (HT), a secoiridoid called elenolic acid and a glucose molecule (Fig. 1). HT can be found in olive products either as the simple phenol or esterified with elenolic acid to form oleuropein and its aglycon and as a part of the verbascoside molecule. In addition, HT is the major bioactive metabolite of oleuropein and it is considered as one of the most powerful naturally derived antioxidants (Ziogas et al., 2010) and it has been demonstrated that the other structural subunit, elenolic

S. Khoufi et al. / Bioresource Technology 102 (2011) 9050–9058

9051

Fig. 1. Oleuropein and its derivatives after enzymatic hydrolysis by b-glycosidase and esterase.

acid, exhibits strong antiviral properties (Micol et al., 2005; Yamada et al., 2009). However, regarding the beneficial effects of oleuropein and its degradation products especially the HT, several methods have been developed to produce this compound by means of chemical synthesis (Tuck et al., 2000), enzymatic conversion (Briante et al., 2000), photochemical synthesis (Azabou et al., 2007) and bench scale purification from olive leaves (Lee and Lee, 2010) and OW (Savournin et al., 2001). Other biological methods have been also developed to produce HT (Brooks et al., 2006; Bouallagui and Sayadi, 2006). Aspergillus species have been studied thoroughly with respect to their ability to produce extracellular enzymes such as cellulases or pectinases which can be applied easily in food processing, as they have achieved FDA recognition as GRAS (Generally Regarded as Safe) substances (Mazzeia et al., 2009). This fungus is indeed efficient producers of cellulolytic enzymes which comprise four types of enzymes acting synergistically: endo-1,4-b-glucanase (EC 3.2.1.4), cellobiohydrolase (EC 3.2.1.91) exo-1,4-b-glucosidase (EC 3.2.1.74) and b-glucosidase (EC 3.2.1.21). b-Glucosidases have been also studied due to its potential to release flavour compounds such as terpenes from odourless non-volatile glycosidic precursors in fruit juices and wines (Maicas and Mateo, 2005) and release phenolic compounds with antioxidant, nutraceutical and flavourant properties from their glycosylated forms in fruit and vegetable residues (Zheng and Shetty, 2000). In the present work, Aspergillus niger culture broth on wheat bran, particularly rich on b-glucosidase and esterase was tested for its ability to release HT from raw OW. Its hydrolysis activity was also evaluated on a phenolic fraction extracted from OW by ethyl acetate and its corresponding exhausted fraction (OW after EA extraction). Optimisation of the enzymatic hydrolysis was studied in order to improve HT yield in OW. 2. Methods 2.1. Material Olive fruits used in this study were collected from olive trees grown under linear irrigation treatment in the region of Sfax (south of Tunisia) in the 2009–2010 harvest season (September). The olives were washed, deleafed, and then crushed with a hammer crusher. Five litres of water were added to 1 kg of homogeneous olive paste, well mixed and then centrifuged to separate oil and suspended solids from the aqueous phase. This olive wastewater (OW) was stored at 20 °C. Wheat bran was obtained locally and stored in a dark room at 4 °C. HT, used as a standard, was purified from olive mill wastewater in the laboratory (Allouche and Fki, 2004).

p-Nitrophenol (p-NP); p-nitrophenyl-b-D-glucopyranoside (pNPG); p-nitrophenyl acetate (p-NPA); butylated hydroxytoluene (BHT); 2,6-dimethoxyphenol (DMP); Folin–Ciocalteu phenol reagent and gallic acid were purchased from Sigma–Aldrich (Switzerland). Finally, 1,1-diphenyl-2-picrylhydrazyl (DPPH) was purchased from Fluka (Switzerland). 2.2. A. niger enzymes production A. niger strain (CTM 10099) was maintained on Potato Dextrose Agar (PDA) at room temperature with periodic transfer. Spores from A. niger were produced by growth on PDA for 7 days at 30 °C and harvested by suspending with a sterilised solution of 0.1% Tween 80. Culture broths of A. niger were obtained after growth on wheat bran as carbon source in 500 mL Erlenmeyer flasks containing 100 mL of basal medium for the fungal growth. A C/N ratio equal to 10 was maintained for this fermentation by adding vegetal peptone and urea as sources of nitrogen. The medium was adjusted to pH 4.0 and sterilized by autoclaving (20 min at 121 °C). Erlenmeyer flasks were inoculated to give 107 spores/ mL as final concentration and incubated at 30 °C for 10 days in an orbital incubator shaker operating at 160 rpm. Every day two flasks from the batch cultivations were carried for enzymes activity analysis and for bioconversion reactions. The culture broth was filtered through muslin cloth to give a cultivation solid fraction (mycelia or biomass) and a cultivation filtrate representing the extracellular medium (filtrate). b-Glucosidase and esterase activities, extracellular protein and reducing sugars (RS) concentrations were measured in the filtrate. 2.3. Enzymatic hydrolysis experiments Enzymatic hydrolysis experiments were conducted in 500 mL Erlenmeyer flasks. Hydrolysis activity of A. niger biomass (mycelium separated from the culture broth and washed with citrate buffer, pH 4.8), filtrate (enzyme preparation) and the whole culture broth (biomass and extracellular medium) were tested. The total amount of bio-catalyst present in the batch cultivation (filtrate, biomass or whole culture broth) was added to 50 mL of substrate. The bioconversion reactions were carried out at 50 °C for 2 h. The OW control was incubated at the same conditions by adding water instead of enzyme. Samples were withdrawn periodically from the biotransformation medium for further analysis. Three different substrates were tested for the proposed enzymatic hydrolysis: raw OW, exhausted fraction and phenolic fraction. The phenolic fraction was obtained by extracting OW with ethyl acetate in a proportion of 1:1 (v/v). The ethyl acetate extract

9052

S. Khoufi et al. / Bioresource Technology 102 (2011) 9050–9058

was evaporated to dryness by rotary evaporation and the residue was resuspended in distilled water with the same initial proportion. After OW extraction, the remaining exhausted OW fraction was also used as substrate for bioconversion.

procedure, a mixture of pyridine (40 lL) and N,O-bis(trimethylsilyl)trifluoroacetamide (200 lL) were added to the sample (200 lL) and vortexed in screw cap glass tubes and consecutively placed in a water bath at 80 °C during 45 min. From the silylated mixture 1 lL was directly analysed by GC–MS.

2.4. b-Glucosidase activity 2.9. High performance liquid chromatography (HPLC) analysis b-Glucosidase (BG) activity was determined according to Norkrans (1950) by measuring the hydrolysis of p-NPG. For the enzymatic assay, the incubation mixture contained 0.9 mL of 5 mM p-NPG in 50 mM citrate buffer (pH 4.8), and appropriately diluted enzyme solution in a total volume of 1 mL. The reaction was performed at 50 °C within 10 min and terminated by the addition of 2 mL of Na2CO3 solution (1.0 M) followed by the addition of 10 mL of water. The amount of p-NP released was determined spectrophotometrically by measuring the absorbance of the solution at 405 nm. One unit (IU) of enzyme activity was defined as the amount of enzyme that produced 1 lmol of p-NP under the conditions of the assay. 2.5. Esterase activity Esterase activity was carried out according to the method of Mackness et al. (1983). The reaction mixture contained 50 lL of crude enzyme, 50 lL of 150 mM p-NPA in ethanol, and 2.9 mL of Tris–HCl buffer (9.2 mM, pH 7.5) to give a final concentration of 2.5 mM p-NPA in a total volume of 3 mL. The reaction was initiated by the addition of enzyme extract and incubated at 25 °C during 4 min. Absorbance was determined at 405 nm. A standard curve was prepared in the range of 0–220 nmol of p-NP. 2.6. Protein analysis Extracellular proteins were determined by the Bradford method (Bradford, 1976), using Bio-Rad protein assay and bovine serum albumin as standard. 2.7. Reducing sugars analysis DNS reagent was prepared by mixing 10.6 g of 3,5-dinitrosalicylic acid and 19.8 NaOH into 1416 mL of distilled water in a stirred beaker. Rochelle salts (potassium sodium tartrate) (306 g), phenol (8.1 g) and sodium sulphite (8.3 g) were then added (AlZuhair, 2008). A sample volume of 100 lL was mixed with 900 lL of citrate buffer (50 mM, pH 4.8). Then, 1.5 mL of DNS solution was added to the mixture and heated in boiling water for 10 min. The samples were then cooled to room temperature with 10 mL of distilled water and their absorbances at 550 nm were determined using a spectrophotometer. A blank solution, consisted of 100 lL of distilled water was used to zero the spectrophotometer before analysis. A standard curve was prepared in the range of 0–2 g/L of glucose. 2.8. Gas chromatography–mass spectrometry (GC–MS) analysis GC–MS analysis was performed with a HP model 5975B inert MSD, equipped with a capillary DB-5MS column with 30 m length, 0.25 mm internal diameter and 0.25 mm film thickness (Agilent Technology, USA). The carrier gas was He used at 1 mL/min flow rate. The oven temperature program was as follows: 1 min at 100 °C, ramped from 100 to 260 °C at 4 °C/min and 10 min at 260 °C. The chromatograph was equipped with a split/splitless injector used in the split mode. The split ratio was 100:1. OW samples (20 mL) were extracted with ethyl acetate (60 mL). The organic phase was collected and reduced to 10 mL by rotary evaporation (37 °C) and was then silylated. For the silylation

The identification and quantification of phenolic monomers were carried out by HPLC. The assays were performed on a Shimadzu apparatus composed of an LC-10ATvp pump and an SPD10Avp UV/Visible detector. The column was a C-18 (4.6  250 mm; Shimpack VP-ODS), and its temperature was maintained at 40 °C. The flow rate was 0.5 mL/min. The mobile phase used was 0.1% phosphoric acid in water (A) versus 70% acetonitrile in water (B) for a total running time of 50 min, and the following proportions of solvent B were used for the elution: 0–30 min, 20–50%; 30–35 min, 50%; and 35–50 min, 50–20%. The flow rate was 0.6 mL/min, and the injection volume was 20 lL. Identification and quantification of HT was based on its spectrum and on its retention time in comparison with standard analysed under the same conditions. 2.10. Total phenol content determination Concentration of total phenols was measured as gallic acid equivalents (Randhir and Shetty, 2007). One millilitre of diluted sample was transferred to a test tube and 1 mL of 95% ethanol; 5 mL of distilled water and 0.5 mL of Folin–Ciocalteu phenol reagent were added. After an incubation period of 5 min, 1 mL of 5% Na2CO3 was added, well mixed and kept in the dark for an hour. Then the samples were vortexed and the absorbance was measured at 725 nm using an UV spectrophotometer. 2.11. DPPH (2,2-diphenyl-1-picrylhydrazyl) radical scavenging assay The DPPH radical-scavenging effect was evaluated according to Bouaziz et al. (2005). A volume of 4 mL from each diluted sample (phenolic extract) was added to a 10 mL of DPPH methanol solution (1.5  10 4 M). After mixing the two solutions softly and after being left for 30 min at room temperature, the optical density was measured at 520 nm, using a spectrophotometer. The antioxidant activity of each sample was expressed in terms of IC50 (microgrammes per mL required to inhibit DPPH radical formation by 50%) and calculated from the log-dose inhibition curve. 3. Results and discussion 3.1. Characteristics of A. niger culture broth on wheat bran Wheat bran, an available by-product supplemented with basal medium was used as a cultivation medium for A. niger aiming to obtain an enzyme mixture optimal for enzymatic hydrolysis of OW. BG production by A. niger grown on wheat bran was enhanced by optimisation of initial pH of the culture medium, temperature cultivation, inoculum size, initial C/N ratio and other parameters (data not shown). The fungal enzyme production was monitored by the measurement of BG and esterase activities. Protein and reducing sugars (RS) contents in the filtrate of the fungus broth culture were also determined. Fig. 2 shows the time course of a typical batch fermentation of A. niger on wheat bran under the optimal conditions. During this fermentation, the fungus showed a rapid growth from the third day of inoculation and the mycelium efficiently colonised the entire substrate by day 8. Fig. 2a, presenting the evolution of the RS and protein concentrations during the

9053

S. Khoufi et al. / Bioresource Technology 102 (2011) 9050–9058

a

RS 10

140 120

8

100 80

6

60

4

40 2

20 0

RS concentration (g/L)

Protein concentration (µg/mL)

Protein

0 0

2

4

6

8

10

Incubation time (days)

b

Esterase

7000

300

6000

250

5000

200

4000 150 3000 100

2000

50

1000

Esterase activity (IU/mL)

B-glucosidase activity (IU/mL)

B-glucosidase

0

0 0

2

4

6

8

10

Incubation time (days) Fig. 2. Protein and sugar concentrations (a) and activities of b-glycosidase (BG) and esterase (b) in the culture broth of A. niger during the incubation time.

incubation time, shows that the RS concentration increased during the first time of incubation reaching 8 g/L at day 2 of fermentation. However, since day 3, this concentration decreased and then stabilized at a value of 1 g/L during all the period of incubation. This result could be explained by the release of RS from wheat bran polysaccharides by the A. niger cellulase and their consumption by the mycelium in the same time of the release. Determination of BG and esterase activities showed interesting values during the fermentation of wheat bran (Fig. 2b). In fact, BG activity increased in a steep manner from 3 to 7 days incubation time when the protein concentration was stable at a concentration of 80 lg/ mL. After this time, the BG production took a less steep behaviour. The highest BG activity (6000 IU/mL) was found in the culture broth of 10-day incubation. However, the peak of esterase activity (230 IU/mL) is essentially sustained by a protein concentration present after 2 days of incubation time (Fig. 2b). These results indicate an efficient colonisation of the fungus on the wheat bran substrate with an efficient production of two enzymatic activities of interest that their appearance is in two well separated moments of the incubation time. Analyses of other enzyme activities have shown that A. niger broth culture on wheat bran is a multienzyme mixture containing pectinolytic, cellulolytic, amylolytic and xylanolytic enzymes (data not shown). These enzymes are known to play an important role in the mobilisation of the substrate phenolic compounds during fungal bioprocessing (Randhir and Shetty, 2007). The use of A. niger enzyme preparation rich on BG and esterase may prove useful in solving the problem of HT low concentrations in fresh OW and technical industrial problems relating to chemical hydrolysis of OW. 3.2. Enzymatic hydrolysis of OW by A. niger culture broth HT is the main natural phenolic compound occurring in OW (Marco et al., 2007), and it is also present in virgin olive oil, albeit

in very small amounts and mainly in the form of natural derivatives. HT is incorporated in the aglycon of oleuropein and is thought to be released by hydrolysis from this glycoside during olive storage and pressing, due to the action of cellular enzymatic or acidic catalysis (Brenes et al., 2001; Jemai et al., 2009). Due to the rising interest in sustainable development and environmentally friendly practices, microbial enzyme transformation processes are generally preferred over the conventional chemical conversion process. For this reason, the study of OW hydrolysis by A. niger broth culture for HT enrichment was suggested. Optimisation of operation conditions was also investigated. 3.2.1. Substrate selection for HT bioformation The first possibility for increasing the recovery of HT from OW is to proceed with an ethyl acetate (EA) extraction and then to expose the extract to the hydrolytic activity of the A. niger enzymes. For this reason, the organic fraction recovered by EA extraction was dried by rotovapor and dissolved in a fixed volume of distilled water. The second and third alternatives were to expose raw OW and exhausted fraction (OW obtained after EA extraction) to the enzymatic hydrolysis. A. niger culture broth (5-days incubation) was used for enzymatic reactions of the three samples during 2 h. These preliminary enzymatic reactions were catalysed by the whole culture broth. The filtrate of this enzymatic bio-catalyst has a BG and esterase activities of 3000 and 100 IU/mL, respectively (Fig. 2b). Concentrations of total phenols in the phenolic fraction, exhausted fraction and raw OW were 1.2, 4.6 and 5.8 g/ L, respectively (Table 1). As shown in Table 1, depending on the substrate, the total phenolic content after hydrolysis ranged between 1.5 and 7.1 g/L. All of these were higher than that of the non-hydrolysed substrate. These data indicate that exhausted fraction releases the highest quantity of free phenols. The change of the total phenolic content in raw OW and exhausted fraction was due to the mobilisation of simple phenolic compounds from the polyphenol complexes whereas in the case of the phenolic fraction was due to the phenolic compounds present in the enzyme preparation which were released during the hydrolysis of wheat bran (Williamson et al., 1998). This enhanced phenolic content can improve the antioxidant value of OW particularly when the released phenol is HT. Qualitative and quantitative analyses of simple phenolics recovered from samples by EA extraction procedure were monitored by HPLC. The HPLC chromatograms of raw OW, phenolic fraction and exhausted fraction before and after hydrolysis are shown in Fig. 3. The identification of the different phenolics was confirmed by GC– MS analysis and by comparison with reference compounds (Fig. 4). The main phenolic compounds identified were: tyrosol, HT, p-coumaric acid and caffeic acid (Fig. 4). Some fatty acids were also detected (Fig. 4). HPLC profile of raw OW shows that oleuropein and HT were the major compounds and they represented 66.5% and 2%, respectively. Tyrosol is also detected but its concentration is very low in comparison with the other phenolics. Fig. 3 shows that HPLC chromatograms exhibited an increase of HT peak with a concomitant decrease of the oleuropein peak after hydrolysis treatment of

Table 1 Total phenols content and hydroxytyrosol (HT) concentration of phenolic fraction, exhausted fraction, raw OW and acidified OW before and after enzymatic hydrolysis treatment. Phenols (g/L)

HT (g/L)

Hydrolysis treatment

Before

After

Before

After

Phenolic fraction Exhausted fraction Raw OW Acidified OW (pH 2)

1.20 4.60 5.80 6.40

1.50 6.20 7.10 –

0.05 0.02 0.05 0.18

0.15 0.75 0.56 –

9054

S. Khoufi et al. / Bioresource Technology 102 (2011) 9050–9058

Fig. 3. HPLC chromatograms (UV 280 nm) of the phenolic fraction (EA extract) (a), hydrolysed phenolic fraction (b), exhausted fraction (c), hydrolysed exhausted fraction (d), hydrolysed raw OW (e) and acidified (pH 2) OW (f). Tyrosol: Ty; hydroxytyrosol: HT; para-coumaric acid: PC; caffeic acid: CA and oleuropein: OE.

samples. This can be explicated by the action of BG and esterase for the cleavage of the glucoside and the ester links present in oleuropein that leads to the production of HT, elenolic acid and glucose (Fig. 1). As can be seen in the HPLC chromatograms, hydrolysed OW exhibited a high HT content (0.56 g/L) in comparison to control and acidified OW. Also, these profiles show an increase of tyrosol (Ty), p-coumaric acid (PC) and caffeic acid (CA) concentrations

(Fig. 3). Furthermore, results presented in Table 1 note that enzymatic bioconversion increased the HT concentration about 37.5 and 11 times respectively in the presence of exhausted fraction and raw OW. Using the exhausted fraction as substrate, 0.75 g/L of HT were produced suggesting that enzymatic hydrolysis can be a very promising technology that may be used for increasing the recovery of HT from exhausted OW obtained after a first set of solvent extraction of OW. The use of raw OW for enzymatic

S. Khoufi et al. / Bioresource Technology 102 (2011) 9050–9058

9055

Fig. 4. Total ion gas chromatogram of ethyl acetate-extractable products obtained from raw (a) and hydrolysed OW (b). The MS-identified compounds with respect to their retention time are the following: 15.69: tyrosol (1); 21.05: hydroxytyrosol (2); 25.16: para-coumaric acid (3); 27.59: palmitic acid (4); 29.9: caffeic acid (5); 31.39: oleic acid (6).

biotransformation to release maximum concentration of HT could be also very realistic. Our results confirmed that BG and esterase are the key factor of HT release from oleuropein while an other study has demonstrated the enzymatic bioconversion of this compound by using homogeneous recombinant BG from hyperthermophilic archaeon Sulfolobus solfactaricus expressed in Escherichia coli (Briante et al., 2000). Here, it can be noted that under the biotransformation conditions used in this study, HT was obtained from oleuropein and other complex molecules present in OW, which are less extractable by the EA solvent. In fact, OW generally contains in addition to oleuropein other glucosylated phenolic molecules, such as ligstroside and demethyl-oleuropein that, when they are hydrolysed give rise to HT and several forms of elenolic acid (Mazzeia et al., 2009). The enhancement of substrate bioconversion to HT is probably due to the action of BG, esterase and other enzymes. This study has also demonstrated that acidic hydrolysis cannot release significant amount of HT (only 0.18 g/L) in comparison with the enzymatic pre-treatment adopted (0.56 g/L). The acidification–

extraction process was usually used for the recovery of OW phenolics (Allouche and Fki, 2004; Lesage-Meessen et al., 2001). OW acidification before extraction was necessary to remain more phenolic compounds as HT. This OW extraction process resulted in an EA extract containing the target compounds and an acidified exhausted fraction representing the aqueous phase of OW. This fraction is characterised by a very high acidity, salinity and COD values, in addition to the presence of other recalcitrant organic molecules (Khoufi et al., 2008). These characteristics make the acidified wastewater very difficult to treat; so the development of a suitable sequence to carry out in a depuration process appears of great interest whereas in the case of enzymatic hydrolysis, the acidification problem is avoided. The recovery of HT from OW or exhausted fraction pre-treated by A. niger enzyme preparation is of great importance, not only because of its aforementioned significant properties, but also because it could exploit a large amount of OW. In the basis of this part of study, large amount of HT together with other monomeric phenols could be found in OW after hydrolysis treatment by a convenient enzymatic process. This procedure

9056

S. Khoufi et al. / Bioresource Technology 102 (2011) 9050–9058

is environmentally friendly and is adaptable to industrial processes. 3.2.2. Effect of bio-catalyst nature, reaction time and incubation time of A. niger culture on HT bioformation In order to improve HT production, several operating factors could be studied, mainly the nature of the bio-catalyst, the bioconversion reaction time and the incubation time of A. niger culture. A. niger enzymes can be present free in the aqueous culture or fixed on the biomass. In this part of study, characterisation of active enzymes location (free or fixed on the biomass) was investigated. These enzymes were considered as bio-catalysts for the target biotransformation. For this reason, bioconversion reactions of OW by biomass (mycelia), culture filtrate and the whole culture broth (mixture) were investigated. Fig. 5 shows the time course of HT formation during these reactions. Blank control without enzyme was also run, and no significant HT production was observed (Fig. 5). The HT measurement of the blank showed a value of 0.05 g/L corresponding only to 6.25% of its final value in hydrolysed OW. It should be noted that in the absence of broth culture, substrate was not converted to HT even with high temperature condition (50 °C). Bioconversion of OW by filtrate was characterised by a rapid HT production with concentrations higher than those obtained in similar experiments in the presence of the whole culture broth and the washed biomass. The amount of HT produced in the presence of filtrate (0.80 g/L) increased by 42.8% compared to mixed culture (0.56 g/L). Here, we can explain this difference by the degradation of HT by the biomass. The use of biomass, previously washed by citrate buffer, resulted in a less HT release (0.08 g/L) during the early times of reaction. However, the HT concentration had increased to 0.27 g/L after 6 h of bioconversion reaction (Fig. 4). This can indicates that the strain still produced its enzyme during the incubation with OW. In this case, it was noted an increase of BG activity in the enzymatic react (220 IU/ mL). It can be concluded that the highest concentration of HT (0.8 g/L) was obtained when using the enzyme filtrate produced by A. niger having a BG activity of 3000 IU/mL. Therefore, the subsequent enzymatic hydrolyses of OW were performed using enzymes from cultivation filtrate. According to these results, the filtrate is the best bio-catalyst for the HT release. To achieve maximum HT yield, the growth state of A. niger culture used for OW conversion was also optimised. Batch fermentations of wheat bran with A. niger were investigated under the same controlled conditions. Culture filtrates obtained at different incubation times were tested for OW bioconversion. Results showed that HT concentration increases with the use of the filtrate in the first day of incubation. This filtrate presents only esterase activity (130 UI/mL). After this time, the release of HT was concomitant with the BG activity present in the culture broth (Figs. 2b and 6).

Hydroxytyrosol concentration (g/L)

Mixture

Biomass

Filtrate

Control

1 0.8 0.6 0.4 0.2 0 0

2

4

6

8

10

Bioconversion time (h) Fig. 5. Evolution of hydroxytyrosol concentration in OW during hydrolysis reaction by the mixture, filtrate and biomass of A. niger culture broth at 50 °C.

An association between elevated BG activity (day 2–5) and HT mobilisation was observed. Esterase activity was constant during this time of incubation. Nevertheless, after 5 days of incubation, no further HT increasing was observed. At higher enzyme loading (4800 IU/mL) using filtrate of broth culture at day 8, the HT concentration was 0.5 g/L, being less than the one on day 5. Indeed, HT concentration was found to decrease slowly. This could be due to the fact that HT is degraded by the spores which have appeared from day 6. These findings indicate that BG is the key enzyme in this process; also the BG concentration is essential for OW polymers bioconversion to release HT. This was demonstrated by Capasso et al. (1997) who have studied the enzymatic hydrolysis of oleuropein, extracted from olive plant, by free BG from almond. It should also be taken into account that the amount of HT in OW comes not only from the hydrolysis of oleuropein and verbascoside but also from the hydrolysis of hydroxytyrosol 4-b-D-glucoside (Romero et al., 2002) where the BG is the key enzyme for HT release. The presence of relatively high levels of BG activity together with esterase seems to be essential for efficient enzymatic hydrolysis of OW. The use of A. niger enzyme preparation obtained after 5 days of incubation is an effective bio-catalyst for the OW bioconversion. Under these biotransformation conditions, high amounts of HT (0.8 g/L) are collected by a very simple procedure (enzymatic hydrolysis – EA extraction) which may prove useful for a further application for recycling OW. Homogeneous recombinant BG from hyperthermophilic archaeon S. solfactaricus, expressed in E. coli, was immobilized on chitosan support and used to produce hydroxytyrosol from commercially available oleuropein (Briante et al., 2000). In addition, the antioxidant properties of the main products obtained were purified and characterised (Briante et al., 2001) until a production of hydroxytyrosol of 91–94% in weight was achieved (Briante et al., 2004). 3.3. Antioxidant activity Phenolic compounds are known to exhibit free-radical scavenging (antioxidant activity), which is determined by their reactivity as hydrogen or electron donors, the stability of the resulting antioxidant-derived radical, their reactivity with other antioxidants and their metal chelation properties (De Leonardis et al., 2007). In the present study, antioxidant activity of extracts from raw OW, hydrolysed OW, exhausted fraction, hydrolysed exhausted fraction, purified HT and BHT were measured by DPPH free radical scavenging method and the IC50 are presented in Table 2. The DPPH method estimates the ability of the extract to quench the DPPH free radical. All extracts showed antioxidant potential in the DPPH radical scavenging assay (Table 2). BHT showed strong antioxidant activity with an IC50 value of 29 lg/mL. The antioxidant activity of raw OW was naturally very low (IC50 = 650 lg/ mL) in comparison with the activity of BHT. However, this activity increased after enzymatic hydrolysis with an IC50 value of 75.2 lg/ mL (Table 2). These antioxidant activities of phenolic extracts depend on the qualitative characteristics of phenolic profile and not just on the total phenolic content (Randhir and Shetty, 2007; Ziogas et al., 2010; Lee and Lee, 2010). The IC50 value of hydrolysed OW extract proves that this extract contains compounds of high antioxidant activity as HT (IC50 = 23 lg/mL). Table 2 shows also the antioxidant activity of extracts from exhausted fraction and hydrolysed exhausted fraction. From these results we have noted the very low antioxidant activity of extract from exhausted fraction (IC50 = 805 lg/mL) this means that the most phenolic compounds having the propriety of high antioxidant activity were extracted during the first stage of EA extraction. Hydrolysing this exhausted fraction using the enzyme preparation of A. niger, a high antioxidant activity (IC50 = 20 lg/mL) was registered witch is higher than

9057

0.8 0.6 0.4 0.2

10

9

ay D

D

ay

8

7

ay D

ay

6 D

5

ay D

ay D

ay

4

3 D

ay D

ay D

ay D

C

on tro

2

1

0

l

Hydroxytyrosol concentration (g/L)

S. Khoufi et al. / Bioresource Technology 102 (2011) 9050–9058

Fig. 6. Hydroxytyrosol content after bioconversion of OW using filtrate of A. niger culture broth at different times of incubation.

Table 2 Radical scavenging effect of raw OW, hydrolysed OW, exhausted fraction, hydrolysed exhausted fraction, BHT and purified hydroxytyrosol. Samples

IC50 (lg/mL)

Raw OW Hydrolysed OW Exhausted fraction Hydrolysed exhausted fraction BHT Hydroxytyrosol

650 75.2 805 20 29 23

the activity of hydrolysed OW, BHT and HT. This could be explicated by the synergy reaction between HT and other compounds also released during the enzymatic hydrolysis as caffeic acid and p-coumaric acid (Fig. 2). It is known from literature that, among OW phenolic compounds, HT and caffeic acid exhibited good antioxidant properties, but in all cases, tyrosol and ferulic acid are considered to be nutraceutically positive. Previous results showed that olive phenols recovered by EA extraction from OW are good antioxidants for lard (De Leonardis et al., 2007). Others studies showed that OW polyphenols have an antioxidant effect on intestinal human epithelial cells (Manna et al., 1997) and a cytostatic action on some tumour cells (Owen et al., 2000). Fki et al. (2005) showed that OW extract can be used as alternative natural antioxidants to stabilize edible oils, while at the same time appeasing a major concern of consumers over the use of synthetic antioxidants in food products. On the basis of these results, the proposed enzymatic hydrolysis of OW will improve the biological value of its phenolics that are useful for industrial and pharmaceutical applications. 4. Conclusion This study suggested enzymatic treatment for OW with the aim of increasing HT concentration. Hydrolysis treatments were investigated by using culture broth of A. niger on wheat bran. The use of filtrate from A. niger culture broth (5 days) as a bio-catalyst releases 0.8 g of HT per litre of OW after 2 h of hydrolysis reaction. This process produced a natural and a bioactive product from a vegetal source as opposed to the molecule obtainable through chemical synthesis. The proposed enzymatic pre-treatment may prove useful not only for laboratory applications, but also for potential industrial application for recycling OW. Acknowledgement This research was supported by Contrats Programmes Ministère de l’Enseignement Supérieur et de la Recherche Scientifique,

Tunisie. The authors would like to thank Mr. Adel Gargoubi for his technical assistance in HPLC analysis.

References Agalias, A., Magiatis, P., Skaltsounis, A.L., Mikros, E., Tsarbopoulos, A., Gikas, E., Spanos, I., Manios, T.A., 2007. New process for the management of olive oil mill waste water and recovery of natural antioxidants. J. Agric. Food Chem. 55, 2671–2676. Allouche, N., Fki, I., Sayadi, S., 2004. Toward a high yield recovery of antioxidants and purified hydroxytyrosol from olive mill wastewaters. J. Agric. Food Chem. 52, 267–273. Al-Zuhair, S., 2008. The effect of crystallinity of cellulose on the rate of reducing sugars production by heterogeneous enzymatic hydrolysis. Bioresour. Technol. 99, 4078–4085. Azabou, S., Najjar, W., Ghorbel, A., Sayadi, S., 2007. Mild photochemical synthesis of the antioxidant hydroxytyrosol via conversion of tyrosol. J. Agric. Food Chem. 55, 4877–4882. Bouallagui, Z., Sayadi, S., 2006. Production of high hydroxytyrosol yields via tyrosol conversion by Pseudomonas aeruginosa immobilized resting cells. J. Agric. Food Chem. 54, 9906–9911. Bouaziz, M., Grayer, R.J., Simmonds, M.S.J., Damak, M., Sayadi, S., 2005. Identification and antioxidant potential of flavonoids and low molecular weight phenols in olive cultivar chemlali growing in Tunisia. J. Agric. Food Chem. 53 (2), 236–241. Bradford, M.M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254. Brenes, M., Garcia, A., Garcia, P., Garrido, A., 2001. Acid hydrolysis of secoiridoid aglycons during storage of virgin olive oil. J. Agric. Food Chem. 49, 5609–5614. Briante, R., La Cara, F., Febbraio, F., Barone, R., Piccialli, G., Carolla, R., Mainolfi, P., De Napoli, L., Patumi, M., Fontanazza, G., Nucci, R., 2000. Hydrolysis of oleuropein by recombinant b-glycosidase from hyperthermophilic archae on Sulfolobus solfataricus immobilised on chitosan matrix. J. Biotechnol. 77, 275–286. Briante, R., La Cara, F., Tonziello, M.P., Febbraio, F., Nucci, R., 2001. Antioxidant activity of the main bioactive derivates from oleuropein hydrolysis by hyperthermophilic b-glycosidase. J. Agric. Food Chem., 3198–3203. Briante, R., Patumi, M., Febbraio, F., Nucci, R., 2004. Production of highly purified hydroxytyrosol from Olea europaea leaf extract biotransformed by hyperthermophilic b-glycosidase. J. Biotechnol. 111, 67–77. Brooks, S.J., Doyle, E.M., O’Connor, K.E., 2006. Tyrosol to hydroxytyrosol biotransformation by immobilised cell extracts of Pseudomonas putida F6. Enzyme Microbiol. Technol. 39, 191–196. Capasso, R., Evidente, V.C., Gianfreda, L., Maremonti, M., Greco, J.R.G., 1997. Production of glucose and bioactive aglycone by chemical and enzymatic hydrolysis of purified oleuropein from Olea europea. Appl. Biochem. Biotechnol. 61, 365–377. De Leonardis, A., Macciola, V., Lembo, G., Aretini, A., Nag, A., 2007. Studies on oxidative stabilisation of lard by natural antioxidants recovered from olive-oil mill wastewater. Food Chem. 100, 998–1004. Fernandez-Bolanos, J., Rodrıguez, G., Rodrıguez, R., Heredia, A., Guillen, R., Jimenez, A., 2002. Production in large quantities of highly purified hydroxytyrosol from liquid–solid waste of two-phase olive oil processing or ‘‘Alperujio’’. J. Agric. Food Chem. 50, 6804–6811. Fki, I., Allouche, N., Sayadi, S., 2005. The use of polyphenolic extract, purified hydroxytyrosol and 3,4-dihydroxyphenyl acetic acid from olive mill wastewater for the stabilization of refined oils: a potential alternative to synthetic antioxidants. Food Chem. 93, 197–204. Gonzalez-Munoz, M.J., Luque, S., Alvarez, J.R., Coca, J., 2003. Recovery of phenol from aqueous solutions using hollow-fibre contactors. J. Membr. Sci. 213, 181–193.

9058

S. Khoufi et al. / Bioresource Technology 102 (2011) 9050–9058

Jemai, H., Bouaziz, M., Sayadi, S., 2009. Phenolic composition, sugar contents and antioxidant activity of Tunisian sweet olive cultivar with regard to fruit ripening. J. Agric. Food Chem. 57, 2961–2968. Khoufi, S., Aloui, F., Sayadi, S., 2008. Extraction of antioxidants from olive mill wastewater and electro-coagulation of exhausted fraction to reduce its toxicity on anaerobic digestion. J. Hazard. Mater. 151, 531–539. Lee, O.-H., Lee, B.-Y., 2010. Antioxidant and antimicrobial activities of individual and combined phenolics in Olea europea leaf extract. Bioresour. Technol. 101 (10), 3751–3754. Lesage-Meessen, L., Navarro, D., Maunier, S., Sigoillot, J.C., Lorquin, J., Delattre, M., Simon, J.L., Asther, M., Labat, M., 2001. Simple phenolic content in olive oil residues as a function of extraction systems. Food Chem. 75, 501–509. Mackness, M.I., Walker, C.H., Rowlands, D.C., Price, N.R., 1983. Esterase activity in homogenates of three strains of the rust red flour beetle Tribolium castaneum (Herbst). Comp. Biochem. Physiol. 74C, 65–68. Maicas, S., Mateo, J.J., 2005. Hydrolysis of terpenyl glycosides in grape juice and other fruit juices: a review. Appl. Microbiol. Biotechnol. 67, 322–335. Manna, C., Galletti, P., Cucciolla, V., Moltedo, O., Leone, A., Zappia, V., 1997. The protective effect of the olive oil polyphenols (3–4-dyhydroxyphenyl) – ethanol counteracts reactive oxygen metabolite induced cytotoxicity in caco-2-cells. J. Nutr. 12, 286–292. Marco, E.D., Savarese, M., Paduano, A., Sacchi, R., 2007. Characterization and fractionation of phenolic compounds extracted from olive oil mill wastewaters. Food Chem. 104, 858–867. Mazzeia, R., Giornoa, L., Piacentinia, E., Mazzucab, S., Drioli, E., 2009. Kinetic study of a biocatalytic membrane reactor containing immobilized b-glucosidase for the hydrolysis of oleuropein. J. Membr. Sci. 339, 215–223. Mekki, A., Dhouib, A., Sayadi, S., 2007. Polyphenols dynamics and phytotoxicity in a soil amended by olive mill wastewaters. J. Environ. Manag. 84, 134–140. Micol, V., Caturla, N., Pérez-Fons, L., Mas, V., Pérez, L., Estepa, A., 2005. The olive leaf extract exhibits antiviral activity against viral haemorrhagic septicaemia rhabdovirus (VHSV). Antiviral Res. 66 (2–3), 129–136. Norkrans, B., 1950. Influence of cellulolytic enzymes from Hymenocetes on cellulose preparations of different crystallinity. Physiol. Plant 3, 75–81.

Owen, R.W., Giacosa, A., Hull, W.E., Haubner, R., Spiegelhalder, B., Bartsch, H., 2000. The antioxidant/anticancer potential of phenolic compounds isolated from olive oil. Eur. J. Cancer 36, 1235–1247. Randhir, R., Shetty, K., 2007. Mung beans processed by solid-state bioconversion improves phenolic content and functionality relevant for diabetes and ulcer management. Innov. Food Sci. Emerg. Technol. 8, 197–204. Reis, M.A.T., De Freitas, O.M.F., Ferreira, L.M., Carvalho, J.M.R., 2006. Extraction of 2(4-hydroxyphenyl) ethanol from aqueous solution by emulsion liquid membrane. J. Membr. Sci. 269, 161–170. Rodríguez, G., Lama, A., Rodríguez, R., Jiménez, A., Guillén, R., Fernández-Bolaños, J., 2008. Olive stone an attractive source of bioactive and valuable compounds. Bioresour. Technol. 99 (13), 5261–5269. Romero, C., Brenes, M., Garciäa, P., Garrido, A., 2002. Hydroxytyrosol 4-â-Dglucoside, an important phenolic compound in olive fruits and derived products. J. Agric. Food Chem. 50, 3835–3839. Savournin, C., Baghdikian, B., Elias, R., Dargouth-Kesraoui, F., Boukef, K., Balansard, G., 2001. Rapid high-performance liquid chromatography analysis for the quantitative determination of oleuropein in Olea europaea leaves. J. Agric. Food Chem. 49 (2), 618–621. Tuck, K.L., Tan, H.W., Hayball, P.J., 2000. Synthesis of tritium-labeled hydroxytyrosol, a phenolic compound found in olive oil. J. Agric. Food Chem. 48 (9), 4087–4090. Williamson, G., Faulds, C.B., Kroon, P.A., 1998. Specificity of ferulic acid (feruloyl) esterases. Biochem. Soc. Trans. 26, 205–209. Yamada, K., Ogawa, H., Hara, A., Yoshida, Y., Yonezawa, Y., Karibe, K., Nghia, V.B., Yoshimura, H., Yamamoto, Y., Yamada, M., Nakamura, K., Imai, K., 2009. Mechanism of the antiviral effect of hydroxytyrosol on influenza virus appears to involve morphological change of the virus. Antiviral Res. 83 (1), 35–44. Zheng, Z., Shetty, K., 2000. Solid-state bioconversion of phenolics from cranberry pomace and role of Lentinus edodes b-glucosidase. J. Agric. Food Chem. 48, 895– 900. Ziogas, V., Tanou, G., Molassiotis, A., Diamantidis, G., Vasilakakis, M., 2010. Antioxidant and free radical-scavenging activities of phenolic extracts of olive fruits. Food Chem. 120, 1097–1103.