Changes in Phenolic Compounds and Phytotoxicity of the Spanish

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Biotechnology and Biological Transformations

Changes in phenolic compounds and phytotoxicity of the Spanish -style green olive processing wastewaters by Aspergillus niger B60 Eugenia Papadaki, Maria Z. Tsimidou, and Fani Th. Mantzouridou J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.8b00918 • Publication Date (Web): 26 Apr 2018 Downloaded from http://pubs.acs.org on April 26, 2018

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Journal of Agricultural and Food Chemistry

Changes in phenolic compounds and phytotoxicity of the Spanish−style green olive processing wastewaters by Aspergillus niger B60

Eugenia Papadaki, Maria Z. Tsimidou, and Fani Th. Mantzouridou* Laboratory of Food Chemistry and Technology, School of Chemistry, Aristotle University of Thessaloniki, 541 24 Thessaloniki, Greece

Corresponding author *Phone: +302310997774; Fax: +302310997847; E−mail: [email protected]

ACS Paragon Plus Environment


1

ABSTRACT

2

This study systematically investigated the degradation kinetics and changes in the

3

composition of phenolic compounds in Spanish−style Chalkidiki green olive

4

processing wastewaters (TOPWs) during treatment using Aspergillus niger B60. The

5

fungal growth and phenol degradation kinetics were described sufficiently by the

6

Logistic and Edward models, respectively. The maximum specific growth rate (2.626

7

1/d) and the maximum degradation rate (0.690 1/h) were observed at 1500 mg/L of

8

total polar phenols, indicating the applicability of the process in TOPWs with high

9

concentration of phenolic compounds. Hydroxytyrosol and the other simple phenols

10

were depleted after 3−8 days. The newly−formed secoiridoid derivatives identified by

11

HPLC−DAD−FLD and LC−MS are likely produced by oleoside and oleuropein

12

aglycon via the action of fungal β−glucosidase and esterase. The treated streams were

13

found to be less phytotoxic with reduced chemical oxygen demand by up to 76%.

14

Findings will provide useful information for the subsequent treatment of residual

15

contaminants.

16 17

KEYWORDS: table olive processing wastewaters, Aspergillus niger, phenol

18

degradation kinetics, hydroxytyrosol degradation, phytotoxicity

19

2 ACS Paragon Plus Environment

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INTRODUCTION

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Table olive processing wastewaters (TOPWs) generated from the olive production

22

plants create severe environmental problems in the major table olive producing areas,

23

especially of the Mediterranean countries.1 The volumes of TOPWs produced and

24

their composition vary widely depending on the processing methods applied.

25

According to our previous study,2 the most polluting ones are those that involve lye

26

treatment and exhaustive washings of the fruit (i.e. 6 and 4 m3/ton of Californian–

27

style black–ripe and Spanish–style green olives, respectively). Also, considering that

28

Spanish−style method is the most commonly applied one worldwide,2 it generates, in

29

absolute terms (liters of effluent generated), the largest volume of TOPWs with the

30

highest chemical oxygen demand (COD) values. Among the resulting wastewaters,

31

lye (L) and washing water (WW) effluents constitute the 75% of the total volume of

32

wastewater production.2 These data, along with the fact that disposal of large volumes

33

of wastewaters is demanded by small–sized enterprises within a short period of time

34

(~ 1 month), reflect the need to give priority to their treatment. Unlike olive mill

35

wastewaters (OMWs), until now, there are no specific regulations regarding the

36

principles for TOPW management. The practices currently applied include land

37

disposal, discharge into rivers or the sea, and storage in evaporation ponds.2

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Among the organic molecules found in TOPWs, the phenolic compounds merit

39

special consideration as they are not easily biodegradable.3 Hydroxytyrosol (HTyr) is

40

one of the main phenolic compounds found in TOPWs, originating from the alkaline

41

hydrolysis of the bitter−tasting oleuropein (Ole).2,4 Its presence in OMWs has been

42

related to the phytotoxic and antibacterial properties of the effluent.5,6

43

Research in the detoxification of olive processing wastewaters is mainly focused

44

on OMW. Our previous work2 highlighted the current advances and challenges of the



TOPW

remediation.

Various

physical

(e.g.

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effective

membrane filtration,

46

evaporation) and chemical methods (e.g. advanced oxidation processes) have been

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applied with varying degree of success.2,4 These treatments offer distinct

48

disadvantages such as generation of polluted and difficult to handle side streams,

49

and/or relatively high energy/cost requirements. Αlternatively, biological treatment of

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TOPWs was recognized as an economically and ecologically viable option. In the

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limited number of publications, activated sludge from municipal wastewater treatment

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plants,3,7 white−rot fungi,8 Aspergillus niger,9–11 Geotrichum candidum12 and the

53

microalga Nannochloropsis gaditana13 were used. Activated sludge, white−rot fungi

54

and microalgae can effectively detoxify the streams, but suffer from drawbacks such

55

as long adaptation periods,3,7,8,13 while G. candidum is effective only after sterilization

56

and supplementation of the target streams with growth factors.12

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In the present study, our interest was focused on A. niger. This microorganism has

58

been used to aid in remediation of TOPWs removing up to 86% of the COD and 48%

59

of phenolic content in 4–days batches.10,11 However, the existing knowledge in the

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TOPW treatment with this fungus is limited, especially with regard to the degradation

61

of phenolic compounds and their metabolites. From the qualitative and quantitative

62

point of view, remarkable differences are noted among the results obtained by

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different authors when studying the phenolic compound composition of the treated

64

effluents. Thus, whereas some authors confirm the ability of A. niger to degrade the

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phytotoxic HTyr in the streams,11 other authors do not.10 Moreover, in the above

66

mentioned studies the TOPWs contained levels of the target compound (< 50 mg/L)

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well below the highest values reported in literature for similar wastewaters (900,

68

1800, 3400 mg/L).4,14,15 What is even more important is that no previous studies have

69

extensively investigated the new compounds produced from the transformation of


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phenolic compounds in TOPWs during fermentation. Last but not least, despite the

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research interest that phytotoxicity of OMW has attracted, relevant studies on

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untreated and treated TOPWs practically do not exist.8

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All the above observations let us to investigate systematically the degradation

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kinetics and changes in the composition of phenolic compounds that occur in L and

75

WW effluents from the processing of Spanish−style Chalkidiki green olives during

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treatment using A. niger B60. This type of product accounts for more than 50% of the

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Greek table olive production and 43% of the table olive exports.16 The potential

78

phytotoxic effect of the untreated and treated streams was evaluated for the first time,

79

to assess the impact of the biological treatment on wastewater quality.

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MATERIALS AND METHODS

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Samples. Fresh L and WW effluents from Spanish−style processing of green

82

olives (cv. Chalkidiki) were obtained from an industrial plant located in Chalkidiki

83

(Northern Greece). Representative samples (20 L) were obtained from the effluents (5

84

m3 of L and 10 m3 of WW) of each tank (8 tons). Sampling was from three different

85

tanks processed in parallel. They were collected just after olive treatment with 2%

86

NaOH aqueous solutions for 11 h (L) and two water changes at 8 and 16 h (WW), and

87

stored immediately at −20 °C. For the preparation of phenol−enriched WW, the dry

88

phenolic extract from WW −obtained as described below− was redissolved in WW at

89

a final concentration of 1500 mg/L. The sampling was repeated for three production

90

seasons (2014−2015, 2015−2016 and 2016−2017).

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Chemicals. Hydroxytyrosol (HTyr), tyrosol (Tyr), p−coumaric acid (CuA), caffeic

92

acid (CA), luteolin−7−O−glucoside (LuG) and oleuropein (Ole) were supplied by

93

Extrasynthѐse S.A. (Genay, France). LC−MS grade formic acid (FA) and acetonitrile

94

were

obtained

from

Merck

(Darmstadt,

Germany).


Phosphoric

acid,


95

2,2′−azino−bis(3−ethylbenzothiazoline−6−sulfonic acid) diammonium salt (ABTS),

96

4−nitrophenyl−β−D−glucopyranoside (PNPG), p−nitrophenyl acetate (PNPA) and

97

p−nitrophenol were from Sigma−Aldrich (Steinheim, Germany). Folin−Ciocalteu

98

reagent, sodium carbonate, HPLC grade acetonitrile and methanol were purchased

99

from Chem−Lab NV (Zedelgem, Belgium). Potato dextrose agar (PDA) was supplied

100

by Lab M Limited (Heywood, UK). All of the other reagents and solvents of

101

appropriate grade were purchased from various producers.

102

Microorganism. A. niger strain B60 (ATCC 201573), generously provided by

103

Prof. T. Roukas (Department of Food Science & Technology, School of Agriculture,

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Aristotle University of Thessaloniki), was regularly subcultured every 2 to 3 months

105

on PDA plates and maintained at 4 °C.

106

Submerged Fermentation. Submerged fermentation experiments were carried out

107

at optimum conditions for TOPW treatment by A. niger reported previously.11

108

Specifically, experiments were conducted under aerobic conditions in Erlenmeyer

109

flasks (250 mL) containing 50 mL of unsterile L or WW without and after enrichment

110

with its own phenols. The initial pH value of the effluents was adjusted to 5 with

111

concentrated HCl (12 N). The effluents were inoculated with a suspension containing

112

2 × 107 spores/mL. A Neubauer hemocytometer (BlauBrand, Wertheim, Germany)

113

was used to count fungal spores. The inoculated streams were incubated at 30 °C for 8

114

(L), 6 (WW) or 12 d (enriched WW) on a rotary shaker (KS 4000i control, IKA,

115

Wilmington, NC) operating at 160 rpm. The non−inoculated effluents were incubated

116

under the same conditions and used as controls. The flasks were withdrawn at the

117

defined time points and the fermentation broth was filtered under reduced pressure

118

(Pump V−700, Büchi, Flawil, Switzerland) through a Whatman 1 filter paper. The


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filtrates were used for further analysis. Mycelium cells were washed with distilled

120

water and dried at 103 °C for the determination of biomass dry weight (g/L).

121

Determination of Total Solids (TS), Total Dissolved Solids (TDS) and Total

122

Suspended Solids (TSS). TS, TDS and TSS (g/L) of the untreated effluents were

123

determined following the procedures described in Standard Methods.17

124

Determination of COD, pH and Electrical Conductivity. COD (g O2/L) was

125

determined by the potassium dichromate method using test tubes and an AL200 COD

126

VARIO Set−Up (Aqualytic, Dortmund, Germany). pH value was measured with a

127

MP 220 pH meter (Mettler−Toledo, Greifensee, Switzerland). Electrical conductivity

128

(mS/cm) was measured using the portable conductivity meter CM 35 (Crison,

129

Barcelona, Spain).

130

Determination of Enzyme Activity. The β−glucosidase, esterase and laccase

131

activities (U/L) were determined according to standard procedures, using PNPG,

132

PNPA and ABTS as a substrate, respectively.11,18 One unit (U) of β−glucosidase or

133

esterase activity was defined as the amount of enzyme required to release 1 µmol of

134

p−nitrophenol per minute under assay conditions. One unit of laccase was defined as

135

the amount of enzyme required to oxidize 1 µmol of ABTS substrate per min.

136

Determination of Sugar and Nitrogen Content. Glucose, fructose and total

137

sugars were quantified by HPLC analysis as described elsewhere.19 Quantitation was

138

performed using external calibration curves for glucose and fructose (CV% = 1.3 and

139

1.7 for glucose and fructose content of the effluent, respectively, n = 5). Total

140

nitrogen was determined by the persulfate digestion method using LCK 338 cuvette

141

tests and a DR3900 spectrophotometer (Hach Lange, Düsseldorf, Germany).20

142

Color Measurement. Color was measured using a portable spectrophotometer

143

(MiniScan, HunterLab, Murnau, Germany) and expressed in terms of the CIELAB



144

parameters (L*, a* and b*). Chroma value ( = ∗ + ∗ ), hue angle (ℎ° =

145

( ∗ ⁄∗ )) and the color difference of the streams due to biological treatment

146

( = ∗ + ∗ + ∗ ) were calculated.21

147

Extraction of Phenolic Compounds. The phenolic extracts of the effluents were

148

obtained according to the liquid–liquid extraction protocol described in El−Abbassi et

149

al.,22 with some minor modifications. Samples (10 mL) were acidified to pH 2 with

150

HCl (6 N) and extracted with petroleum ether (10 mL, once) in order to remove traces

151

of lipids. Then, the polar phenols of the aqueous phase were extracted by ethyl acetate

152

(10 mL, three times). Each mixture was vortexed and subsequently centrifuged at

153

3500 g for 10 min (SL 16R Thermo Fisher Scientific, Darmstadt, Germany). The

154

ethyl acetate extracts were collected and evaporated under vacuum at ~35 °C

155

(Rotavapor, Büchi). The dry residue was dissolved in methanol and finally filtered

156

through a 0.45 µm PTFE filter (Waters). Repeatability of the extraction procedure

157

relating to the total polar phenol (TPP) content was satisfactory (CV% = 4.4, n = 5).

158

Determination of TPP Content. TPP content of the extracts was estimated by the

159

Folin−Ciocalteu (F−C) colorimetric assay.23 A calibration curve was constructed

160

using CA solutions in the range of 50−500 mg/L. Repeatability of measurements was

161

satisfactory (CV% = 2.0 for L and 2.4 for WW, n = 5).

162

RP–HPLC Analysis of Phenolic Compounds. Analysis was performed on an

163

HPLC system equipped with a P4000 pump, a SCM1000 vacuum membrane

164

degasser, a Midas autosampler (Spark, Emmen, The Netherlands), and a UV 6000 LP

165

diode array detector (DAD; Thermo Separation Products, San Jose, CA), connected in

166

series with an SSI 502 fluorescence detector (FLD; Scientific Systems Inc., State

167

College, PA). Separation was achieved on a Discovery HS column C18 (250 x 4.6 mm

168

i.d., 5 µm) (Supelco, Sigma–Aldrich) and the elution system consisted of 0.2% 8 ACS Paragon Plus Environment

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aqueous phosphoric acid (solvent A) and acetonitrile (solvent B). The gradient was as

170

follows: 0 min 10% B, 1 min 10% B, 10 min 20% B, 43 min 50% B, 48 min 95% B,

171

52 min 95% B, 60 min 10% B, at a flow rate of 0.5 mL/min. The injection volume

172

was 10 µL. Peak identification was based on standards available, relative retention

173

times, spectra matching and literature. Quantitation was performed using external

174

calibration curves for Ole (at 240 nm), HTyr and Tyr (exc 280 nm/em 320 nm) in the

175

range of 50–1000, 50–1200 and 20–800 mg/L, respectively (CV% = 1.5 for HTyr at

176

exc 280 nm/em 320 nm, n = 5). Ole was used for oleoside−11−methyl ester (OME),

177

decarboxymethyl oleuropein aglycon (DOA), elenolic acid (EA) and the reduced form

178

of elenolic acid monoaldehyde (rEAM) quantitation at 240 nm, multiplied by their

179

molecular weight ratio (404:540, 320:540, 242:540 and 244:540, respectively).24

180

LC−MS Analysis of Phenolic Compounds. The identity of major peaks was

181

examined using an HPLC system (Shimadzu, Kyoto, Japan) composed of a LC−20

182

AB pump and a SIL 20A autosampler. Detection was realized with a SPD 20A UV–

183

Vis detector coupled in series with LC–MS 2010EV mass selective detector

184

(Shimadzu), equipped with an atmospheric pressure electrospray ionization source

185

(ESI). MS measurements were carried out in negative ionization (PI) mode.

186

Separation was carried out using the same column as above. The eluents were 0.5%

187

aqueous formic acid (solvent A) and 0.5% formic acid in acetonitrile (solvent B) and

188

the gradient program had the same segments as the mentioned above. The flow rate

189

was 0.4 mL/min. Identification was based on comparison of MS data with available

190

ones from literature.

191

Phytotoxicity Test. The phytotoxicity of the untreated alkaline and treated

192

effluents was assessed on commercially sold seeds of Lepidium sativum and Lactuca

193

sativa species, according to standard procedures.25 Prior to phytotoxicity tests, the



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biologically treated effluents were sterilized by filtration though a 0.22 µm pore

195

filtrate membrane. Aliquots (5 mL) of undiluted (100%) or diluted with deionized

196

water (2−75%), were then transferred into 90–mm Petri dishes containing one piece

197

of Whatman 1 filter paper. Ten seeds of each species were placed in each disk. Dishes

198

containing 5 mL of deionized water (control) or sodium hydroxide solutions with an

199

electrical conductivity value of 13 and 29 mS/cm were also run. Germination was

200

conducted over 72 h in darkness at 25 °C. Results were expressed as percentage of

201

relative index of germination (GI) according to equation (Eq.) 1:6

% =

× × 100

(1)

202

where GS and GC are the number of germinated seeds in the sample and control,

203

respectively, while LS and LC are the average root elongation (mm) for the sample and

204

control seeds, respectively. Moreover, EC50 and EC30 values expressed as wastewater

205

concentrations causing 50% and 30% germination inhibition, respectively, were

206

calculated by plotting the % inhibition values against the loge of the concentrations.

207

Each sample test for the calculation of GI% was carried out in triplicate (CV% = 4 on

208

L. sativa seeds at 20% dilution, n = 5).

209

Kinetics of Growth and Phenol Degradation. Among the different models

210

(Logistic, Gompertz and Richards) tested to describe the growth kinetics of A. niger

211

in the streams, the modified Logistic model (Eq. 2) was the most appropriate one:26,27

=

4 # 1 + exp ! $ (% − ' + 2)

(2)

212

where y is the amount of biomass (g/L) at time t, A is the upper asymptote value (g/L),

213

µm is the maximum specific biomass formation rate (1/d), λ is the lag phase duration

214

(d) and t is the time (d).


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The degradation of phenolic compounds was determined by the time segment

216

method (TSM).28 According to that, the degradation rate q is calculated from the

217

changes of TPP content for each time segment during the biological treatment and the

218

kinetic parameters are estimated by the Edward model (Eq. 3):

* = *$ +exp ,

−−0 − exp , 01 ./ .

(3)

219

where q is the specific degradation rate (1/h) at time t, qm is the maximum specific

220

degradation rate (1/h), S is the TPP content (mg/L) at time t, KI is the TPP inhibition

221

constant (mg/L), KS is the TPP affinity constant (mg/L) and t is the time (h).

222

Statistical Analysis. All measurements and treatments were performed in

223

triplicate. Statistical comparisons of the mean values were carried out either by one–

224

way ANOVA, followed by the Duncan’s test, or by Student’s t–test using the SPSS

225

20.0 software (SPSS Inc., Chicago, IL). Results were considered statistically

226

significant at p < 0.05. The kinetic models were fitted to the experimental data with

227

Microsoft Excel spreadsheet using Solver function (Microsoft Corp., Redmond, WA).

228

RESULTS AND DISCUSSION

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Physicochemical Characteristics of Untreated TOPWs. Table 1 shows the

230

physicochemical characteristics of the Spanish−style Chalkidiki green olive

231

processing wastewaters for three consecutive production seasons (2014−2015,

232

2015−2016, 2016−2017). According to data, elevated values of critical wastewater

233

quality indicators such as pH, electrical conductivity, COD and solids have been

234

found in all the TOPW streams from the three tested processing periods. The extreme

235

alkaline pH (10−13) of both L and WW streams can be attributed to the excess of

236

hydroxyl ions. The electrical conductivity of the effluents was elevated (18−29 and

237

8−13 mS/cm for the L and WW, respectively) due to the presence of high



238

concentration of NaOH as a source of sodium ions,2 mainly in the L effluent.

239

Moreover, the L and WW effluents contained significant amounts of TSS (4−7 and

240

1−4 g/L, respectively). Organic matter in terms of COD was between 15−18 g O2/L in

241

L and 11−16 g O2/L in WW. According to the minimum requirements for the

242

discharge of agro–food industrial wastewaters set by the European Union, the TSS

243

and COD values of the streams are far above the maximum prescribed standard limits

244

(TSS < 35 mg/L, COD < 125 mg O2/L).29 The TPP content in L (0.3−0.4 g/L) and

245

WW (0.2−0.7 g/L) varied between the processing periods, probably due to differences

246

in climatic conditions, but was within the wide range of the reported values for the

247

same type of wastewaters (0.2−1.4 g/L).4,8,9,11,12,30 Among the easily assimilable

248

nutrients, L and WW streams contained a relative low amount of sugars (mainly

249

glucose and fructose), in accordance with previous findings.2,15,30 Next to sugars, the

250

levels of nitrogenous compounds (0.1−0.2 g/L of nitrogen) were considered

251

satisfactory for microbial growth.31 The above findings support our previous point

252

regarding the magnitude of TOPW disposal problem and demand for the development

253

of effective bioremediation processes before disposal into the environment.

254

Kinetics of Microbial Growth and Biodegradation of Phenolic Compounds.

255

The kinetics of fungal growth and phenolic compound degradation by A. niger are

256

expected to be influenced by the observed physicochemical characteristics of the L

257

and WW streams. To the best of our knowledge, the related scientific information

258

about the degradation of phenolic compounds is limited to synthetic wastewaters

259

containing phenol as a sole antimicrobial factor.32–34 The L and WW effluents from

260

2014−2015 were used for the preliminary experiments. The findings showed a

261

satisfactory fungal growth (3 g/L biomass) that coincided with TPP content decrease

262

of about 57% and 85% in L and WW, respectively, after completion of the biological


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treatment. Based on these promising results, the L and WW streams from 2015−2016

264

were used for the exhaustive kinetic study. Key findings of this study were verified

265

using the respective streams of the following processing period (2016−2017).

266

Growth Kinetics. The experimental data on biomass formation (Figure 1A, B) were

267

interpreted using the modified Logistic sigmoid function. This non−linear

268

three−parameter model, with R2 equal to 0.993 and 0.995 for L and WW, respectively,

269

showed a good consistency with experimental cell growth data. The derived

270

biological parameters are given in Table 2. Fitted biomass formation responses

271

demonstrated that WW provided faster fungal growth rate (µm) than L (2.040 vs.1.470

272

l/d), achieving also a significantly higher biomass yield (A) (3.243 vs. 2.841 g/L).

273

This might be attributed to the high levels of NaOH and suspended solids in the L

274

stream that prevent microbial growth by increasing turbidity and lowering the

275

dissolved oxygen levels.35 On the other hand, the lag phase period required for

276

adaptation to the L stream was found to be shorter (0.601 d) than that in WW (0.933

277

d) due to the lower initial TPP content in the former (270.4 vs. 561.3 mg/L).32,34 Same

278

trend was also recorded for effluents from the 2016−2017 processing period with a

279

maximum biomass yield (A) of 3.0 and 3.5 g/L in L and WW, respectively. The above

280

findings indicate that the suspended cells could overcome the inhibitory effect of

281

phenolic compounds in WW even though the growth of cells was delayed. The

282

growth performance of A. niger in L and WW, in terms of the estimated parameters,

283

was similar or even better than that reported in synthetic medium with lactose as a

284

sole carbon source31 or in undersized semolina27 under submerged conditions.

285

Phenol Degradation Kinetics. The growth of A. niger in WW resulted in a

286

sequential consumption of sugars and polar phenolic compounds; sugars were almost

287

depleted within the first day and then phenol degradation occurred (Figure 1B, D). In



288

L, TPP content showed a strong fluctuation throughout the treatment (Figure 1C).

289

Specifically, although a decrease was observed for the first day (up to 14%), it

290

increased within the next two days and then an upward trend was evidenced up to a

291

final value of 102.9 mg/L. Considering the significant acidification of both streams

292

during the first two days of treatment (Figure 1A, B), it can be deduced that carbon is

293

directed to primary metabolism yielding biomass and organic acids.9

294

The estimation of phenol degradation kinetic parameters (qm, KI, KS) is essential to

295

evaluate the overall fungal performance in TOPWs. However, no data exist in the

296

literature on these parameters. In this study, the above mentioned parameters were

297

determined for the first time by Edward model fit to data obtained by TSM (Table 2).

298

By applying this segmentation method, realistic values of parameters derive due to the

299

large number of data points (15) considering also the variation of TPP content during

300

the treatment.28 The goodness of fit (R2) (0.734−0.800) was acceptable and

301

comparable to that for other datasets estimated by TSM using bacterial species

302

(0.76−0.77).28 The value of qm in WW was 2 fold higher than that in L (0.297 vs.

303

0.129 1/h), indicating that the degradation rate was positively related to the initial TPP

304

content. However, the lower KI values in TOPWs (113−120 mg/L) compared with the

305

respective values reported for this fungus in synthetic medium containing a mixture of

306

phenol and p−cresol as the only inhibitor (450−600 mg/L)36 indicated a lower

307

tolerance level in the target waste streams. This is probable due to the presence of

308

high levels of other inhibitory factors (i.e. NaOH) that stimulate further the inhibitory

309

effects in TOPWs. On the other hand, the KS values (11−12 mg/L) designate the

310

capability of A. niger B60 to be active at low phenol concentrations. Similar or even

311

higher KS values concerning degradation of phenolics by fungal (50−300 mg/L)36 and

312

bacterial species (1−130 mg/L)28,37 have been also reported.


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313

A TPP content reduction of 62% and 78% in L and WW, respectively, was

314

recorded at the end of incubation (6−8 d) (Figure 1C, D). A similar reduction of

315

phenolics in L (65%) and WW (80%) effluents of the following processing period

316

(2016−2017), ensures a reproducible process. The estimated percent reduction values

317

are around 1.5 fold higher than the mean value reported previously in TOPW with

318

similar composition, but treated with a different A. niger strain (41−48%).9,11 This

319

finding indicates that the fungal potential to degrade the phenolic compounds of this

320

type of waste streams is strain−dependent and highlights the appropriateness of A.

321

niger B60 for this purpose. The latter is strengthened by the fact that A. niger could

322

withstand the toxic effects of the high phenolic concentrations in WW enriched with

323

its own phenolics up to 1500 mg/L. The fungus exhibited similar growth pattern and

324

increased metabolic activity compared to those in original WW (Table 2), resulting in

325

TPP content reduction of 84% after 12 d. This is attributed to the enhanced expression

326

of phenol degrading enzymes to overcome the severe environmental stress.38

327

Changes in Phenolic Compounds during the Biological Treatment of TOPWs.

328

In the limited available reports regarding TOPW treatment with A. niger there is

329

insufficient discussion about the transformation of phenolic compounds during the

330

biological treatment9–11 and chemical structure of the remaining non−biodegradable

331

compounds in the treated streams.9 Also, the ability of the fungus to degrade the

332

phytotoxic hydroxytyrosol (HTyr) is being questioned among the different studies.10,11

333

In the current study, HPLC analysis with fluorescence (FL) and UV multi–wavelength

334

detection at key time intervals gave a new insight into the interpretation of the results.

335

The data of the findings are detailed below.

336

Degradation of Simple Phenols. RP–HPLC profiles of the L and WW from all

337

production seasons at 280 nm and using FL detection (280 exc/320 em nm) were



338

qualitatively similar and indicated that the individual phenolics in untreated streams

339

consisted mainly of simple phenols (Figures 2 and 3). Using FL detection, HTyr (peak

340

2) (Table 3) was found to be the most abundant phenolic compound in both untreated

341

effluents of all processing periods as a result of the debittering process (95–380

342

mg/L). Also present in both effluents was tyrosol (Tyr) (peak 5) (λmax at 239 and 275

343

nm) (60–180 mg/L). The total content of HTyr and Tyr in both untreated effluents

344

was estimated to be comparable to their TPP content (Table 1). Α non−fluorescent

345

compound (peak 3) present in the L stream, eluted at a retention time very close to

346

that of HTyr (Figure 2A, B). The compound exhibits UV bands at 239 and 267 nm,

347

with a slight shoulder at 389 nm, and has a molecular ion with an m/z value of 167

348

(Table 3), that could be assigned as a methoxylated derivative of HTyr. Peak 12 was

349

assigned as acetylhydroxypinoresinol (AHPin) based on absorption spectra (244 and

350

275 nm) and [M–H]– value at m/z 431 (Table 3).39 Peak 3 and AHPin could be

351

artifacts that were formed during the table olive production process and/or the

352

extraction of phenolics from the streams. RP−HPLC chromatogram recorded at 280

353

and 320 nm of both effluents exhibited also phenolic acids and flavonoids that

354

included caffeic acid (CA), luteolin−7−O−glucoside (LuG) and p−coumaric acid

355

(CuA) (peaks 7−9) (Figure 2B, C and 3B, C) in rather low amounts (< 25 mg/L).

356

The parallel monitoring of changes in the phenolic compounds of non−inoculated

357

streams (2015–2016) showed that at the end of incubation of controls, TPP content

358

was slightly reduced (< 20%), whereas HTyr concentration remained the same (< 3%

359

reduction). As far as it concerns the inoculated samples, dramatic changes were

360

observed in the content and profile of phenolic and related compounds. The treatment

361

with A. niger resulted in the total disappearance of the above mentioned compounds

362

in both effluents (Figure 2B−D and 3B−D) and in enriched stream (contained HTyr at


Page 16 of 42

Page 17 of 42


363

about 1000 mg/L) (Supporting Information Figure S1). The time pattern observed for

364

the reduction of the HTyr and Tyr levels in the streams during the treatment was

365

similar to that for TPP content (Figure 1C, D). The degradation of simple phenols by

366

A. niger involves as key steps the aromatic ring hydroxylation and fission catalysed

367

by hydroxylase and dioxygenase.33 This mechanism leads to the transformation of the

368

substrates into intermediates of central metabolic pathways (e.g. the tricarboxylic acid

369

cycle).32 The lack of laccase activity observed in this study (Supporting Information

370

Table S1) was in line with Ayed et al. findings.11

371

The content of TPP (211 to 1384 mg/L) or individual phenols (10 to 1820 mg/L for

372

HTyr) in TOPWs varies considerably among different studies.4,8,10–12,14,30 HTyr values

373

as high as 3.4 g/L have also been reported for wastewaters from Spanish−style

374

processing of green olives of the Manzanilla de Sevilla cultivar.15 To the best of our

375

knowledge, this is the first time to report the ability of A. niger B60 to degrade HTyr

376

in TOPWs with relatively high initial levels of the target compound. Up to now, in

377

TOPW with HTyr content above 45 mg/L, no considerable degradation of the target

378

compound was recorded by a strain of A. niger isolated from TOPWs.10,11

379

Quantitative data from similar studies indicate a decrease in the degradation

380

efficiency of A. niger and other Aspergillus species in synthetic32,33 or industrial

381

media such as OMWs rich in phenolics compounds.20 Whether these differences are

382

correlated with the type of strain and the nutritional composition of the substrate

383

needs further investigation. Moreover, the partial removal of phenolic compounds

384

through adsorption by the fungal biomass attributable to the hydrogen bonding

385

capacity of cell wall material should not be precluded.11

386

Newly−Formed Compounds. The presence of A. niger was responsible not only for

387

the degradation of phenolic compounds, but also for the synthesis of new phenolic



388

and non–phenolic ones (Table 3 and Figures 1C, D; 2A, B; 3A, B). On the HPLC

389

profile of the treated WW at 240 nm (Figure 3A), new peaks were detected (peaks 10,

390

11, 15 and 16) that were assigned as metabolites of oleoside−11−methyl ester (OME)

391

(peak 6)40 (Table 3) with the concomitant disappearance of the latter (Figure 1D). The

392

fragmentation behavior of peaks 10 and 11 with a molecular ion [M–H]– at m/z 243,

393

an adduct ion [M–H+FA]– at m/z 289 and an absorption band at 243 nm, could be

394

explained by assuming the structures of two isomeric reduced forms of elenolic acid

395

monoaldehyde (rEAM) (Table 3).41 Moreover, the structures of compounds 15 and 16

396

with a [M–H]– value at m/z 241 and λmax at 241 nm were elenolic acid (EA) isomers

397

(Table 3).42 Compounds 10 and 11 were also detected in treated L stream (Figure 2A),

398

despite the absence of OME from the untreated one, probably due to its fast

399

degradation under the strong alkaline conditions.43 This finding indicates involvement

400

of a range of different chemical precursors of peaks 10 and 11. This suggestion is

401

strengthened by the detection of two newly−formed compounds in the stream, peaks

402

13 and 14 (Figure 2A). These compounds were assigned as two isomeric dialdehydic

403

forms of decarboxymethyl oleuropein aglycon (DOA) by a λmax at 241 and 286 nm,

404

with a molecular ion [M–H]– at m/z 319, a dimer [2M–H]– at m/z 639, and

405

characteristic fragment ions of [M–H+FA]– at m/z 365 and of [M–H–C2H3O]– at m/z

406

275 (Table 3).44,45 Their disappearance after 6 days of L treatment was followed by

407

the simultaneous appearance of peaks 10 and 11, which suggests that they may play a

408

role in the formation of the latter. The degradation of phenolics in both streams was

409

also accompanied by the generation of a conjugated diene (peak 4) (Figures 2A, 3A)

410

with a λmax at 241 nm, and an unknown compound (peak 1) (Figures 2B, 3B) with λmax

411

values at 239 and 277 nm, a shoulder at 330 nm, and [M–H]– value at m/z 155 (Table

412

3). The latter has a very low retention time and thus a strong polarity, and was


Page 18 of 42

Page 19 of 42


413

abundant during the treatment of L effluent. This compound was also observed in the

414

current study after acid hydrolysis of the TOPW phenolic extracts (data not shown).

415

The formation of EA isomers 1 and 2 in WW can be mainly assigned to the

416

activity of A. niger β–glucosidase (maximum activity of 32 U/L at day 3) (Supporting

417

Information Table S1), according to the scheme proposed in Figure 4. Nevertheless,

418

the hydrolysis of the glycosidic bond of the OME (R−1) can be also promoted by the

419

lower pH value environment of the TOPWs due to the production of acids by fungal

420

metabolism.46 In L, the enzymatic cleavage of the ester bond in the DOA (R−2) gave

421

rise to the formation of HTyr and EA isomer 2, with esterase exhibiting its highest

422

activity (36 U/L) on day 6 (Supporting Information Table S1). Thus, the constant or

423

even slightly increased content of HTyr during the first 3 days of L treatment should

424

result from its simultaneous biosynthesis and degradation, reflecting the trend of TPP

425

content (Figure 1C). Using EA isomer 2 as substrate, elenolic acid monoaldehyde

426

(EAM) was formed via a highly selective conjugate addition reaction (R−3). The

427

mechanism involves carbonyl oxygen lone pair attack at the electrophilic β carbon of

428

the α, β – unsaturated carbonyl group.45 Finally, aldose reductase catalysed the

429

reduction of the EAM carbonyl group and the formation of rEAM in both effluents

430

(R−4).41 In contrast to WW, EA in L was quickly transformed into rEAM, thus only

431

the latter compound was detected in this stream (Figure 1C, 2A). Quantitative data for

432

the metabolites produced after the biological treatment highlighted that the

433

newly−formed secoiridoid derivatives in L (81.8 mg/L rEAM isomers) and WW

434

(112.5 mg/L EA and 51.4 mg/L rEAM isomers) contribute to the TPP content of the

435

treated streams (102.9 and 124.7 mg/L, respectively) (Figure 1C, D). All the above changes were verified by using streams of 2016−2017 production

436 437

season (data not shown), showcasing reproducibility of the biological treatment.



438

Effect on Phytotoxicity. To assess the environmental impact of simple phenol

439

degradation and secoiridoid formation, the phytotoxic potential of the untreated

440

alkaline and treated with A. niger streams was evaluated. The results of the GI that

441

can be used as an indicator of phytotoxicity are shown in Table 4. In the case of

442

untreated L effluent, without and even after 50% dilution no seed germination was

443

observed for both L. sativum and L. sativa species. In contrast, the effect of 50, 75,

444

and the 100% WW resulted in higher (less toxic) GI values, despite the higher TPP

445

content (Table 1). This can be attributed to the higher concentration of sodium ions in

446

L which was reflected in the electrical conductivity differences between the two

447

streams (29 vs. 13 mS/cm for L and WW, respectively). The GI of aqueous solutions

448

of NaOH with electrical conductivity values equal to those of untreated L and WW

449

were also tested. At 13 mS/cm, the GI showed a reduction of 30 and 46% for L.

450

sativum and L. sativa, respectively, as compared to control water (100%), while at 29

451

mS/cm no seed germination was observed for both species. All the above indicate the

452

primary role of NaOH in the phytotoxicity of the untreated alkaline TOPWs. This is

453

related to the fact that high salinity causes osmotic stress and induces nutritional

454

imbalance, while alkalinity has a strong caustic effect and reduces the solubility of

455

important nutrients.47 The contribution of phenolic compounds in the phytotoxic

456

activity of the streams became clear by the fact that GI values of NaOH solutions at

457

13 mS/cm were 8 to 9 fold higher than those of the untreated WW. Ιn the case of L,

458

this was evident only after stream dilution to 20%, when the effect caused by sodium

459

ions was minimized. The suggested synergistic effect between NaOH and phenolics

460

on phytotoxicity is in accordance with similar studies reported for winery

461

wastewaters48 and TOPWs.8


Page 20 of 42

Page 21 of 42


462

Treatment with A. niger significantly reduced the phytotoxicity of both effluents.

463

This was confirmed by the results described in Table 4, which presented the GI values

464

for every diluted sample assessed on the two plant species, and the EC50 and EC30

465

values for each sample. Specifically, the respective EC50 and EC30 values of the

466

treated streams were 51–79% and 22–40% higher than those of the untreated ones

467

(Table 4). The positive effect of the biological treatment on the phytotoxic activity of

468

the streams was more pronounced in treated samples with effluent concentrations

469

lower than 20%, having GI values ≥ 80% that indicate zero or low concentrations of

470

phytotoxic substances.49 Concentrations up to 4% of treated streams seem not only to

471

reduce their phytotoxicity, but also to stimulate plant growth. The superiority of A.

472

niger to decrease the phytotoxicity of TOPWs, as compared to white−rot fungi,

473

because of the formation of the toxic phenoxy radicals and quinonoids by the latter8

474

highlight further the pros of the proposed process.

475

Effect on Color and COD. Color and COD levels in treated TOPWs can be used

476

to evaluate further their status.10,11 As shown in Table 5, after treatment the effluents

477

became lighter. Also, the treatment increased significantly b* values (moving to

478

yellowish tones) of WW, while decreased a* and b* values (faintly red and less

479

yellow) of L. In both cases, color intensity (Chroma) reached a final value of about 40

480

and color changes were above the “just noticeable difference” value (∆E ≥ 2.3)

481

proposed by Mahy et al.50 Decolorization can be due to different factors including

482

degradation and biosorption of phenolics on fungal mycelium, depolymerization of

483

complex polyphenols with dark color and other reaction enhanced in acidic

484

environment.11,20 The overall process efficiency was reflected to a COD reduction of

485

59% and 76% in L (4.0 g O2/L) and WW (7.2 g O2/L), respectively, in accordance to

486

previous studies that used A. niger for TOPW treatment.9–11 Noticeably, the



487

percentage reduction of COD was in line with the respective one of TPP content (62%

488

in L and 78% in WW), indicating that organic matter removal was mainly attributed

489

to the degradation of phenolics. Despite this significant reduction, the final values

490

were above the Greek limits for acceptance to the sewage (COD < 1.2 g O2/L).10

491

Overall, the results of the current study indicate for the first time that A. niger B60

492

is effective for the rapid degradation of HTyr and other simple phenols present in

493

Spanish−style Chalkidiki green olive processing wastewaters, in amounts ranging

494

from 0.2 to 1.5 g/L. These values are among the highest ones reported so far for this

495

type of wastewaters demonstrating the general applicability of the treatment. The

496

newly−formed compounds, EA and rEAM, are likely produced by OME and DOA.

497

The treated streams were less phytotoxic, with COD values of lower than 7.2 g O2/L.

498

Findings will provide useful information for the subsequent treatment of residual

499

contaminants.

500 501

ASSOCIATED CONTENT

502

Supporting Information

503

Figure S1, HPLC chromatograms of phenol−enriched washing water extracts

504

(recorded at 280 nm) before and after the biological treatment with A. niger B60;

505

Table S1, Data of β–glucosidase, esterase and laccase activity during L and WW

506

treatment with A. niger (PDF)

507 508

ABBREVIATIONS USED

509

ABTS, 2,2′−azino−bis(3−ethylbenzothiazoline−6−sulfonic acid) diammonium salt;

510

AHPin, acetylhydroxypinoresinol; ANOVA, analysis of variance; CA, caffeic acid;

511

CuA, p−coumaric acid; COD, chemical oxygen demand; DOA, decarboxymethyl


Page 22 of 42

Page 23 of 42


512

oleuropein aglycon; EA, elenolic acid; EAM, elenolic acid monoaldehyde; EC,

513

effective concentration; ESI, electrospray ionization source; FA, formic acid; F−C,

514

Folin−Ciocalteu; FL, fluorescence; GI, germination index; HCl, hydrogen chloride;

515

HPLC, high−performance liquid chromatography; HTyr, hydroxytyrosol; LC, liquid

516

chromatography; LuG, luteolin−7−O−glucoside; L, lye; M, molecular ion; MS, mass

517

spectrometry; NaOH, sodium hydroxide; Ole, oleuropein; OME, oleoside−11−methyl

518

ester; OMWs, olive mill wastewaters; PDA, potato dextrose agar; PNPA,

519

p−nitrophenyl acetate; PNPG, 4−nitrophenyl−β−D−glucopyranoside; rEAM, reduced

520

form of elenolic acid monoaldehyde; RP, reversed−phase; TDS, total dissolved solids;

521

TOPWs, table olive processing wastewaters; TPP, total polar phenol; TS, total solids;

522

TSM, time segment method; TSS, total suspended solids; Tyr, tyrosol; Un,

523

unidentified; UV, ultraviolet; WW, washing water.

524



525

REFERENCES 1. Şen, M.; Yentürk, A. Zero discharge of olive waste: Green energy application.

526

Olivae 2016, 123, 37–42.

527 528

2. Papadaki, E.; Mantzouridou, F. T. Current status and future challenges of table

529

olive processing wastewater valorization. Biochem. Eng. J. 2016, 112, 103–

530

113.

531

3. Brenes, M.; García, P.; Romero, C. N.; Garrido, A. Treatment of green table

532

olive waste waters by an activated−sludge process. J. Chem. Technol.

533

Biotechnol. 2000, 75, 459–463.

534

4. Gargouri, B.; Gargouri, O. D.; Khmakhem, I.; Ammar, S.; Abdelhèdi, R.;

535

Bouaziz, M. Chemical composition and direct electrochemical oxidation of

536

table olive processing wastewater using high oxidation power anodes.

537

Chemosphere 2017, 166, 363–371.

538

5. Fiorentino, A.; Gentili, A.; Isidori, M.; Monaco, P.; Nardelli, A.; Parrella, A.;

539

Temussi, F. Environmental effects caused by olive mill wastewaters: Toxicity

540

comparison of low−molecular−weight phenol components. J. Agric. Food

541

Chem. 2003, 51, 1005–1009.

542

6. Isidori, M.; Lavorgna, M.; Nardelli, A.; Parrella, A. Model study on the effect

543

of 15 phenolic olive mill wastewater constituents on seed germination and

544

Vibrio fischeri metabolism. J. Agric. Food Chem. 2005, 53, 8414–8417.

545

7. Beltran, J.; Gonzalez, T.; Garcia, J. Kinetics of the biodegradation of green

546

table olive wastewaters by aerobic and anaerobic treatments. J. Hazard.

547

Mater. 2008, 154, 839–845.

548

8. Aggelis, G.; Ehaliotis, C.; Nerud, F.; Stoychev, I.; Lyberatos, G.; Zervakis, G.

549

I. Evaluation of white–rot fungi for detoxification and decolorization of


Page 24 of 42

Page 25 of 42


550

effluents from the green olive debittering process. Appl. Microbiol.

551

Biotechnol. 2002, 59, 353–360.

552

9. Kotsou, M.; Kyriacou, A.; Lasaridi, K.; Pilidis, G. Integrated aerobic

553

biological treatment and chemical oxidation with Fenton’s reagent for the

554

processing of green table olive wastewater. Process Biochem. 2004, 39, 1653–

555

1660.

556

10. Kyriacou, A.; Lasaridi, K. E.; Kotsou, M.; Balis, C.; Pilidis, G. Combined

557

bioremediation and advanced oxidation of green table olive processing

558

wastewater. Process Biochem. 2005, 40, 1401–1408.

559

11. Ayed, L.; Chammam, N.; Asses, N.; Hamdi, M. Optimization of biological

560

pretreatment of green table olive processing wastewaters using Aspergillus

561

niger. J. Bioremed. Biodeg. 2013, 5 (1), 1–10.

562

12. Ayed, L.; Asses, N.; Chammam, N.; Hamdi, M. Improvement of green table

563

olive processing wastewater decolorization by Geotrichum candidum. Desalin.

564

Water Treat. 2016, 57 (37), 1–11.

565

13. Serrano, A.; Contreras, C.; Ruiz–Filippi, G.; Borja, R.; Fermoso, F. G.

566

Sequential adaptation of Nannochloropsis gaditana to table olive processing

567

water. J. Environ. Sci. Health A Tox. Hazard. Subst. Environ. Eng. 2017, 52,

568

986–991.

569

14. Segovia Bravo, K. A.; Arroyo López, F. N.; García García, P.; Durán

570

Quintana, M. C.; Garrido Fernández, A. Treatment of green table olive

571

solutions with ozone. Effect on their polyphenol content and on Lactobacillus

572

pentosus and Saccharomyces cerevisiae growth. Int. J. Food Microbiol. 2007,

573

114, 60–68.



Page 26 of 42

574

15. Brenes, M.; Concepción, R.; De Castro, A. Combined fermentation and

575

evaporation processes for treatment of washwaters from Spanish−style green

576

olive processing. J. Chem. Technol. Biotechnol. 2004, 79, 253–259. 16. Union of the processing and packaging exporters of table olives (PEMETE),

577

http:// http://www.pemete.gr

578 579

17. Eaton, A. D.; Clesceri, L. S.; Greenberg, A. E.; Franson, M. A. H. In Standard

580

methods for the examination for water and wastewater, 19th edition; American

581

Public Health Association (APHA): Washington, DC, 1995.

582

18. Hamza, M.; Khoufi, S.; Sayadi, S. Changes in the content of bioactive

583

polyphenolic compounds of olive mill wastewater by the action of exogenous

584

enzymes. J. Agric. Food Chem. 2012, 60, 66–73.

585

19. Lalou, S.; Ferentidou, M.; Mantzouridou, F.; Tsimidou, M. Balsamic type

586

varietal vinegar from cv. Xinomavro (Northen Greece). Optimization and

587

scale−up of the alcoholic fermentation step using indigenous multistarters.

588

Food Chem. 2018, 244, 266–274.

589

20. Salgado, J.; Abrunhosa, L.; Venâncio, A.; Domínguez, J.; Belo, I. Combined

590

bioremediation and enzyme production by Aspergillus sp. in olive mill and

591

winery wastewaters. Int. Biodeterior. Biodegradation. 2016, 110, 16−23. 21. Segovia−Bravo,

592

K.

A.;

Garcia−Garcia,

P.;

Arroyo−López,

F.

N.;

593

López−López, A.; Garrido−Fernández, A. Ozonation process for the

594

regeneration and recycling of Spanish green table olive fermentation brines.

595

Eur. Food Res. Technol. 2008, 227, 463–472.

596

22. El–Abbassi, A.; Khayet, M.; Hafidi, A. Micellar enhanced ultrafiltration

597

process for the treatment of olive mill wastewater. Water Res. 2011, 45, 4522–

598

4530.


Page 27 of 42


599

23. Nenadis, N.; Kyriakoudi, A.; Tsimidou, M. Z. Impact of alkaline or acid

600

digestion to antioxidant activity, phenolic content and composition of rice hull

601

extracts. LWT Food Sci. Technol. 2013, 54, 207–215. 24. Tasioula–Margari,

602

M.;

Tsabolatidou,

E.

Extraction,

separation,

and

603

identification of phenolic compounds in virgin olive oil by HPLC–DAD and

604

HPLC–MS. Antioxidants 2015, 4, 548–562.

605

25. Yousuf, A.; Sannino, F.; Addorisio, V.; Pirozzi, D. Microbial conversion of

606

olive oil mill wastewaters into lipids suitable for biodiesel production. J.

607

Agric. Food Chem. 2010, 58, 8630–8635.

608

26. Zwietering, M. H.; Jongenburger, I.; Rombouts, F. M.; Van't Riet, K.

609

Modeling of the bacterial growth curve. Appl. Environ. Microbiol. 1990, 56,

610

1875–1881.

611

27. Erkmen, O.; Alben, E. Mathematical modeling of citric acid production and

612

biomass formation by Aspergillus niger in undersized semolina. J. Food Eng.

613

2002, 52, 161–166. 28. Lee, S.; Kim, D. J.; Choi, J. W. Novel method for determination of phenol

614

degradation kinetics. Bioprocess Biosyst. Eng. 2013, 36, 1939–1945.

615

29. Council Directive 91/271/EEC of 21 May, 1991 concerning urban waste water

616

treatment. Off. J. Eur. Communities 1991, L135, 40−52.

617 618

30. Parinos, C. S.; Stalikas, C. D.; Giannopoulos, T. S.; Pilidis, G. A. Chemical

619

and physicochemical profile of wastewaters produced from the different stages

620

of Spanish–style green olives processing. J. Hazard. Mater. 2007, 145, 339–

621

343.



622

31. Villena, G. K.; Gutiérrez–Correa, M. Assessment of Aspergillus niger biofilm

623

growth kinetics in minibioreactors by carbon dioxide evolution. Afr. J.

624

Biotechnol. 2011, 10, 13495−13504.

625

32. Santos, V.; Linardi, V. Biodegradation of phenol by a filamentous fungi

626

isolated from industrial effluents—identification and degradation potential.

627

Process Biochem. 2004, 39, 1001–1006. 33. Supriya, M.; Neehar, D. Biodegradation of phenol by Aspergillus niger. IOSR

628

J. Pharm. 2014, 4, 11–17.

629 630

34. Tebbouche, L.; Hank, D.; Zeboudj, S.; Namane, A.; Hellal, A. Evaluation of

631

the phenol biodegradation by Aspergillus niger: application of full factorial

632

design methodology. Desalin. Water Treat. 2015, 57, 6124−6130. 35. Bilotta, G.; Brazier, R. Understanding the influence of suspended solids on

633

water quality and aquatic biota. Water Res. 2008, 42, 2849–2861.

634 635

36. Yemendzhiev, H.; Zlateva, P.; Alexieva, Z. Comparison of the biodegradation

636

capacity of two fungal strains toward a mixture of phenol and cresol by

637

mathematical modeling. Biotechnol. Biotechnol. Equip. 2012, 26, 3278–3281.

638

37. Aryal, M.; Liakopoulou–Kyriakides, M. Phenol degradation in aqueous

639

solutions by Pseudomonas sp. isolated from contaminated soil of mining

640

industry. J. Water Sustainability 2015, 5, 45–57.

641

38. Chakraborty, B.; Ray, L.; Basu, S. Study of phenol biodegradation by an

642

indigenous mixed consortium of bacteria. Indian J. Chem. Technol. 2015, 22,

643

227−233.

644

39. Obied, H. K.; Bedgood, D. R. Jr.; Prenzler, P. D.; Robards, K. Chemical

645

screening of olive biophenol extracts by hyphenated liquid chromatography.

646

Anal. Chim. Acta 2007, 603, 176−189.


Page 28 of 42

Page 29 of 42


647

40. Cecchi, L.; Migliorini, M.; Cherubini, C.; Innocenti, M.; Mulinacci, N. Whole

648

lyophilized olives as sources of unexpectedly high amounts of secoiridoids:

649

the case of three Tuscan cultivars. J. Agric. Food Chem. 2015, 63, 1175−1185.

650

41. Pinto, J.; Paiva−Martins, F.; Corona, G.; Debnam, E. S.; Oruna–Concha, M.

651

J.; Vauzour, D.; Gordon, M. H.; Spencer, J. P. E. Absorption and metabolism

652

of olive oil secoiridoids in the small intestine. Br. J. Nutr. 2011, 105, 1607–

653

1618.

654

42. Sato, A.; Shinozaki, N.; Tamura, H. Secoiridoid type of antiallergic substances

655

in olive waste materials of three japanese varieties of Olea europaea. J. Agric.

656

Food Chem. 2014, 62, 7787−7795. 43. Friedman, M.; Jürgens, H. S. Effect of pH on the stability of plant phenolic

657

compounds. J. Agric. Food Chem. 2000, 48, 2101−2110.

658 659

44. De la Torre–Carbot, K.; Jauregui, O.; Gimeno, E.; Castellote, A. I.; Lamuela–

660

Raventós, R. M.; López–Sabater, M. C. Characterization and quantification of

661

phenolic compounds in olive oils by solid−phase extraction, HPLC−DAD, and

662

HPLC−MS/MS. J. Agric. Food Chem. 2005, 53, 4331–4340.

663

45. Vougogiannopoulou, K.; Angelopoulou, M. T.; Pratsinis, H.; Grougnet, R.;

664

Halabalaki, M.; Kletsas, D.; Deguin, B.; Skaltsounis, L. A. Chemical and

665

biological investigation of olive mill wastewater – OMWW secoiridoid

666

lactones. Planta Med. 2015, 81, 1205–1212.

667

46. Medina, E.; Romero, C.; Brenes, M.; García, P.; De Castro, A.; García, A.

668

Profile of anti–lactic acid bacteria compounds during the storage of olives

669

which are not treated with alkali. Eur. Food Res. Technol. 2008, 228, 133–

670

138.



671

47. Day, A. D.; Ludeke, K. L. Soil Alkalinity. In Plant Nutrients in Desert

672

Environments. Adaptations of Desert Organisms; Cloudsley–Thompson, J. L.,

673

Punzo, F., Eds.; Springer: Berlin, Germany, 1993; pp. 35–37.

674

48. Mosse, K. P.; Patti, A. F.; Christen, E. W.; Cavagnaro, T. R. Winery

675

wastewater inhibits seed germination and vegetative growth of common crop

676

species. J. Hazard. Mater. 2010, 180, 63–70.

677

49. Mañas, P.; De las Heras, J. Phytotoxicity test applied to sewage sludge using

678

Lactuca sativa L. and Lepidium sativum L. seeds. Int. J. Environ. Sci. Technol.

679

2018, 15, 273–280.

680

50. Mahy, M.; Van Eycken, L.; Oosterlinck, A. Evaluation of uniform color

681

spaces developed after the adoption of CIELAB and CIELUV. Color Res.

682

Appl. 1994, 19, 105–121.

683


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684

FIGURE CAPTIONS

685

Figure 1. Evolution of biomass formation, sugar consumption and pH in lye (L) (A)

686

and washing water (WW) (B) effluents (2015−2016 processing period), and the

687

content of total polar phenol (TPP) and individual phenolic compounds

688

(hydroxytyrosol,

689

decarboxymethyl oleuropein aglycon, DOA; elenolic acid, EA; reduced form of

690

elenolic acid monoaldehyde, rEAM) in L (C) and WW (D) effluents treated with A.

691

niger B60. Error bars represent the standard deviation of the mean value (n = 3).

HTyr;

tyrosol,

Tyr;

oleoside−11−methyl

ester,

OME;

692 693

Figure 2. Changes in the phenolic profiles of the lye (L) extracts (2015−2016

694

processing period) during the biological treatment with A. niger B60 by using HPLC.

695

Detection was at (A) 240 nm, (B) 280 nm, (C) 320 nm (DAD), as well as (D) λexc 280

696

nm; λem 320 nm (FLD). Peaks: (1) Unidentified 1, Un1; (2) Hydroxytyrosol, HTyr;

697

(3) Unidentified 2, Un2; (4) Unidentified 3, Un3; (5) Tyrosol, Tyr; (7) Caffeic acid,

698

CA; (8) Luteolin−7−O−glucoside, LuG; (9) p−Coumaric acid, CuA; (10) Reduced

699

form of elenolic acid monoaldehyde, rEAM isomer 1; (11) rEAM isomer 2; (12)

700

Acetylhydroxypinoresinol, AHPin; (13) Decarboxymethyl oleuropein aglycon, DOA

701

isomer 1; (14) DOA isomer 2.

702 703

Figure 3. Changes in the phenolic profiles of the washing water (WW) extracts

704

(2015−2016 processing period) during the biological treatment with A. niger B60 by

705

using HPLC. Detection was at (A) 240 nm, (B) 280 nm, (C) 320 nm (DAD), as well

706

as (D) λexc 280 nm; λem 320 nm (FLD). Peaks: (1) Unidentified 1, Un1; (2)

707

Hydroxytyrosol,

708

oleoside−11−methyl ester, OME; (7) Caffeic acid, CA; (8) Luteolin−7−O−glucoside,

HTyr;

(4)

Unidentified

3,

Un3;


(5)

Tyrosol,

Tyr;

(6)


709

LuG; (9) p−Coumaric acid, CuA; (10) Reduced form of elenolic acid monoaldehyde,

710

rEAM isomer 1; (11) rEAM isomer 2; (15) Elenolic acid, EA isomer 1; (16) EA

711

isomer 2.

712 713

Figure 4. Proposed conversion mechanisms for the production of the newly–formed

714

compounds during treatment of TOPWs with A. niger B60.

715


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Table 1. Physicochemical Characteristics of Untreated Lye (L) and Washing Water (WW) Effluents for Three Consecutive Production Seasonsa 2014−2015

2015−2016

2016−2017

Characteristics

L

WW

L

WW

L

WW

pH

12.8 ± 0.2 a

11.2 ± 0.1 b

13.4 ± 0.0 c

11.1 ± 0.1 d

12.1 ± 0.0 e

10.1 ± 0.1 f

Electrical conductivity (mS/cm)

18.4 ± 0.0 a

7.9 ± 0.0 b

29.3 ± 0.1 c

13.0 ± 0.1 d

20.0 ± 0.0 e

9.9 ± 0.0 f

COD (g O2/L)

15.3 ± 0.5 a

10.7 ± 0.1 b

17.3 ± 0.1 c

15.5 ± 0.1 a

17.7 ± 0.3 c

15.7 ± 0.2 a

Total solids (g/L)

18.3 ± 1.0 a

9.8 ± 1.1 b

26.0 ± 0.4 c

13.2 ± 0.2 d

18.0 ± 0.5 a

12.6 ± 0.4 d

Total dissolved solids (g/L)

14.1 ± 0.3 a

8.6 ± 0.1 b

18.9 ± 0.4 c

11.6 ± 0.1 d

13.3 ± 0.2 e

9.2 ± 0.2 f

Total suspended solids (g/L)

4.2 ± 0.3 a

1.3 ± 0.1 b

7.1 ± 0.4 c

1.6 ± 0.1 b

4.8 ± 0.2 d

3.5 ± 0.2 e

Total polar phenol content (mg/L)

262.1 ± 5.7 a

220.9 ± 4.5 b

270.4 ± 5.2 a

561.3 ± 9.5 c

429.2 ± 10.0 d

720.1 ± 12.1 e

Total nitrogen (mg/L)

72.2 ± 0.3 a

119.2 ± 1.3 b

139.0 ± 7.9 c

176.5 ± 7.5 d

91.2 ± 3.5 e

124.5 ± 3.8 b

Total sugars (g/L)

5.1 ± 0.1 a

2.8 ± 0.0 b

4.9 ± 0.1 c

4.2 ± 0.1 d

6.0 ± 0.1 e

5.1 ± 0.1 a

Glucose (g/L)

2.4 ± 0.0 a

1.3 ± 0.0 b

2.3 ± 0.0 c

2.0 ± 0.0 d

2.9 ± 0.0 e

2.4 ± 0.1 f

Fructose (g/L)

2.7 ± 0.1 a

1.5 ± 0.0 b

2.6 ± 0.0 c

2.3 ± 0.1 d

3.1 ± 0.1 e

2.6 ± 0.1 f

a

Results are given as mean ± standard deviation (n = 3). Significant differences in the same row were marked with lowercase letters (p < 0.05).

33

ACS Paragon Plus Environment


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Table 2. Growth and Phenol Degradation Kinetic Parameters Obtained from the Treatment of Lye (L) and Washing Water (WW) Effluents (2015−2016 Processing Period) with A. niger B60a Effluent

L

Growth parametersb

WW

Enriched WW

Logistic model

A (g/L)

2.841 ± 0.059 a

3.243 ± 0.038 b

3.904 ± 0.011 c

µm (1/d)

1.470 ± 0.064 a

2.040 ± 0.063 b

2.626 ± 0.033 c

λ (d)

0.601 ± 0.043 a

0.933 ± 0.001 b

0.955 ± 0.012 b

R2

0.993 ± 0.002 a

0.995 ± 0.003 a

0.994 ± 0.001 a

Degradation parametersc

Edward model

qm (1/h)

0.129 ± 0.018 a

0.297 ± 0.016 b

0.690 ± 0.069 c

KI (mg/L)

113.3 ± 11.0 a

120.1 ± 3.6 a

300.1 ± 13.6 b

KS (mg/L)

10.7 ± 0.9 a

11.7 ± 0.2 a

46.2 ± 4.6 b

R2

0.800 ± 0.053 a

0.734 ± 0.037 ab 0.823 ± 0.022 ac

a

Each value was expressed as mean ± standard deviation of three replicates.

Significant differences in the same row marked with lowercase letters (p < 0.05). bA = upper asymptote value, µm = maximum specific biomass formation rate and λ = lag phase duration. cqm = maximum specific degradation rate, KI = total polar phenol (TPP) inhibition constant and KS = TPP affinity constant.


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Table 3. Main Compounds Detected in Lye (L) and Washing Water (WW) Phenolic Extracts (2015−2016 Processing Period) during Treatment with A. niger B60 by HPLC−DAD−FLD and LC−MS (ESI−, SIM) Data Peaksa Phenolic compoundsb Un1 1 HTyr 2 Un2 3 Un3 4 Tyr 5 OME 6 CA 7 LuG 8 CuA 9 rEAM 1 10 rEAM 2 11 AHPin 12 DOA 1 13 DOA 2 14 EA 1 15 EA 2 16 a

λmax (nm)c 239, 277, 330sh 239, 279 239, 267, 389sh 241 239, 275 241 241, 289, 322 245, 348 242, 307 243 243 244, 275 241, 286 241, 286 241 241

LC−MS (ESI−) (m/z) 155, 201 153 167

403, 449, 807 179 447, 493 163 243, 289 243, 289 431 319, 275, 365, 639 319, 275, 365, 639 241 241

FLd No Yes No No Yes No No No No No No No No No Yes No

Peak numbering as in Figures 2 and 3. bUn = unidentified, HTyr = hydroxytyrosol,

Tyr = tyrosol, OME = oleoside−11−methyl ester, CA = caffeic acid, LuG = luteolin−7−O−glucoside, CuA = p−coumaric acid, rEAM = reduced form of elenolic acid monoaldehyde, AHPin = acetylhydroxypinoresinol, DOA = decarboxymethyl oleuropein aglycon and EA = elenolic acid. cλmax = maximum absorbance (UV−Vis spectra). dFL = fluorescence.



Table 4. Germination index (%) of L. sativum and L. sativa Seeds on Untreated and Treated Lye (L) and Washing Water (WW) Effluents (2015−2016 processing period) with A. niger B60 and Determination of the Effective Concentration Expressed as EC50 and EC30 (% v/v) for Seed Germinationa Waste Concentration

Germination index (% of control) L WW Untreated Treated Untreated Treated L. sativum 0 ± 0 a, A 2 ± 0 b, A 3 ± 1 b, A 5 ± 1 c, A 100% 0 ± 0 a, A 4 ± 1 b, A 5 ± 1 b, A 7 ± 1 c, A 75% 0 ± 0 a, A 5 ± 0 b, A 40 ± 1 c, B 47 ± 1 d, B 50% 20 ± 1 a, B 30 ± 2 b, B 74 ± 3 c, C 91 ± 4 d, C 20% 63± 4 a, C 83 ± 2 b, C 82 ± 4 b, D 108 ± 3 c, D 10% 76 ± 3 a, D 94 ± 2 b, D 98 ± 5 b, E 135 ± 5 c, E 4% 98 ± 2 a, E 114 ± 5 b, E 108 ± 3 b, F 151 ± 5 c, F 2% 100 0% 11 ± 1 a 16 ± 0 b 24 ± 2 c 37 ± 1 d EC50 23 ± 1 a 31 ± 1 b 50 ± 2 c 62 ± 2 d EC30 L. sativa 0 ± 0 a, A 0 ± 0 a, A 5 ± 1 b, A 5 ± 1 b, A 100% 0 ± 0 a, A 0 ± 0 a, A 10 ± 2 b, B 12 ± 1 c, B 75% 1 ± 0 a, A 5 ± 1 b, B 42 ± 3 c, C 50 ± 2 d, C 50% 35 ± 2 a, B 47 ± 1 b, C 59 ± 2 c, D 73 ± 3 d, D 20% 62 ± 4 a, C 82 ± 3 b, D 76 ± 3 b, E 100 ± 2 c, E 10% 82 ± 3 a, D 119 ± 2 b, E 88 ± 2 c, F 134 ± 4 d, F 4% 90 ± 2 a, E 138 ± 4 b, F 98 ± 1 c, G 146 ± 2 d, G 2% 100 0% 12 ± 1 a 21 ± 1 b 21 ± 1 b 35 ± 1 c EC50 25 ± 1 a 35 ± 1 b 49 ± 1 c 60 ± 2 d EC30 a Data are the mean ± standard deviation (n = 3). Different lowercase letters in the same row or capital letters in the same column represent significant differences in values (p < 0.05).


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Table 5. CIELAB parameters (L*, a*, b*), Chroma Value, Hue Angle (h0) and Total Color Difference (∆E*) of Untreated and Treated Lye (L) and Washing Water (WW) Effluents (2015−2016 processing period) with A. niger B60 a Effluent Parameters

L

WW

0d

8d

L*

46.4 ± 0.2 a

74.6 ± 0.0 b

24.9 ± 0.3 c

78.7 ± 0.0 d

a*

26.9 ± 0.1 a

3.1 ± 0.1 b

1.5 ± 0.1 c

0.9 ± 0.0 d

b*

72.5 ± 0.1 a

38.2 ± 0.1 b

1.6 ± 0.1 c

39.5 ± 0.1 d

Chroma

77.3 ± 0.1 a

38.3 ± 0.1 b

2.2 ± 0.1 c

39.5 ± 0.1 d

h0

69.7 ± 0.1 a

85.4 ± 0.0 b

45.7 ± 0.5 c

88.8 ± 0.0 d

∆E*

9.3 ± 0.0 a

0d

6d

9.6 ± 0.0 b

a

Results were expressed as means ± standard deviation of three values. Significant differences in the same row marked with lowercase letters (p < 0.05).



Figure 1.


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Figure 2.



Figure 3.


Page 40 of 42

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Figure 4.



TOC Graphic


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Changes in Phenolic Compounds and Phytotoxicity of the Spanish

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