Antioxidant and biocide behaviour of lignin fractions from apple tree pruning residues

Industrial Crops & Products 104 (2017) 242–252

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Antioxidant and biocide behaviour of lignin fractions from apple tree pruning residues

MARK

⁎

Araceli Garcíaa,b, , Giorgia Spignoc, Jalel Labidib a b c

Department of Organic Chemistry, University of Cordoba UCO, Campus of Rabanales, Marie Curie (C-3) Building, Ctra Nnal IV km 396, 14014 Córdoba, Spain Department of Chemical and Environmental Engineering, University of the Basque Country UPV-EHU, Plaza Europa, 1, 20018 Donostia-San Sebastián, Spain Institute of Oenology and Agro-Food Engineering, Universitá Cattolica del Sacro Cuore, Via Emilia Parmense, 84, 29122 Piacenza, Italy

A R T I C L E I N F O

A B S T R A C T

Keywords: Lignin Biorefinery processes Antioxidant capacity Biocide effect Yeast Microbial growth

Different lignin fractions were obtained from apple tree pruning residues following different isolation processes (autohydrolysis, organosolv with ethanol and acetic acid, and soda processes) and acid hydrolysis purification to obtain the acid insoluble lignin fractions. All the obtained lignin samples were characterised for chemical composition, chemical structure (by ATR-IR, NMR and Pyr-GC/MS analyses), molecular weight distribution (by GPC), total phenolic content, antioxidant power and antimicrobial activity towards a typical food and environment contaminant (Aspergillus niger) and a food processing yeast (Saccharomyces cerevisiae). The results showed different chemical composition and structure for the lignin samples, in particular for the soda lignin characterised by a higher hemicellulose and inorganic matter content. The acid hydrolysis was effective in removing hemicellulose but partially modified the lignin structure. The total phenolic content was the lowest for the soda lignin, the highest for the autohydrolysis lignin and was generally reduced after acid hydrolysis. However, the specific antioxidant capacity (in relation to the phenolic content) was higher for the organosolv lignin samples and was not reduced by acid hydrolysis (with exception for the soda lignin). Addition of lignin samples in the culturing media of A. niger, delayed the growth and brought to colonies with different morphological aspects. On the contrary, the lignin samples showed a clear inhibitory effect on S. cerevisiae growth.

1. Introduction

external aggressions, such as weather, fungi and insects. The antioxidant activity of lignin, i.e. the ability to prevent or retard the oxidation processes induced by oxidizing species such as free radicals, makes it suitable as additive against oxidation and bio- or photo-degradation in polymer blends, cosmetics and pharmaceuticals (Bhat et al., 2009). Due to this, lignin is being widely investigated for new and value-added applications related to its bio-activity, such anti-inflammatory, antibacterial or anti-carcinogenic agent (Hollman, 2001). Nowadays, the research on the use of plant extracts or phytochemicals in food and feed industries for the development of the so-called functional food (Baurhoo et al., 2008; Chun et al., 2005) is becoming of great interest for health promotion through diet. In addition, the use of natural additives result in profitable outcomes for food processing industries, since consumers prefer natural supplements, fearing that synthetic ingredients may be toxic or unhealthy (Ayala-Zavala et al., 2011; Mikulásová and Kosíková, 2003). The use of natural compounds is not limited to ingredients for healthcare benefits, but also as additive to ensure food safety. This way, the use of plant extracts (organic acids,

Lignin is the second most predominant biopolymer present in nature. Together with cellulose and hemicelluloses, it forms the plant cell wall and, because of its renewable origin, lignin is becoming an interesting natural source of aromatic compounds. Obtained as main by-product from lignocellulose transformation industries, such as pulp and paper or bio-ethanol production processes, the actual applications of lignin represent low added value uses (energy, dispersing, binding or emulsifying applications). However, the increasing interest on the profitable exploitation of renewable resources is leading on the development of different processes, in order to comprehensively use the lignocellulosic biomass and revalue all its components, particularly the lignin (Amendola et al., 2012; García et al., 2012, 2010; Serrano et al., 2010). As many other natural phenolic compounds, lignin is biosynthesized by plants and contains one or more hydroxyl groups in its aromatic structure. This phenolic structure acts as plant protector against

⁎ Corresponding author at: Department of Organic Chemistry, University of Cordoba UCO, Campus of Rabanales, Marie Curie (C-3) Building, Ctra Nnal IV km 396, 14014 Córdoba, Spain E-mail address: [email protected] (A. García).

http://dx.doi.org/10.1016/j.indcrop.2017.04.063 Received 4 January 2017; Received in revised form 26 April 2017; Accepted 30 April 2017 0926-6690/ © 2017 Elsevier B.V. All rights reserved.

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essential oils, other plant extracts …) with known antimicrobial properties actually results of great significance in food preservation (Hollman, 2001; Liu et al., 2007; Negi, 2012; Wijekoon et al., 2011). The way that these phytochemicals interact with microorganisms is a very important issue because of their antibacterial effects. Natural compounds could prevent proliferation of undesired organisms in food and therefore avoid contamination due to harmful secondary metabolites or mycotoxins (Ghorai et al., 2009). On the other hand, this biocide effect could interfere in the growth of certain yeast and fungi, that have an important role in industrial food production and processing (brewing, wine making, and bread making industries), and even disappear during food/feed heat processing stages, such as sterilization, pasteurization, or dehydration (Negi, 2012). As non-digestible component of plants, lignin is the main constituent of the dietary fibre, with beneficial nutritive and human/ animal protective effects (Ayala-Zavala et al., 2011; Mikulásová and Kosíková, 2003). Furthermore, lignin from different sources has been widely studied because of its antioxidant behaviour (Amendola et al., 2012; García et al., 2010; Ugartondo et al., 2009) and Alcell (organosolv) and kraft lignins have been probed as prebiotic additive for farm animals (Baurhoo et al., 2008). Based on the above premises, in the present work, different lignin fractions were obtained from apple tree pruning wastes following four fractionation processes: autohydrolysis (HL), organosolv treatment with acetic acid-water (AL) or with ethanol-water (EL), and soda hydrolysis (SL). Purified lignin samples were also obtained by acid hydrolysis to remove hemicellulose residues (PHL, PAL, PEL and PSL) and obtain the acid insoluble lignin (AIL) fraction. All the lignin samples were characterized for chemical composition (content of inorganic matter, acid insoluble and soluble lignin, and hemicelluloses), chemical structure (by attenuated-total reflectance infrared spectroscopy ATR-IR, proton nuclear magnetic resonance analysis 1H NMR and pyrolysis gas chromatography/mass spectroscopy Pyr-GC/MS), total phenolic content (by the Folin-Ciocalteu assay) and antioxidant capacity (by the ABTS radical assay). Finally, the lignin fractions were also evaluated for their potential antimicrobial activity. In this case, Aspergillus niger was selected as a typical food and environment contaminant, therefore an inhibitory activity would make lignin interesting for application as natural preservative in food products or as natural biocide in paints and coatings. Saccharomyces cerevisiae was instead selected as a typical technological microorganism at the basis of many fermented food products (leavened bakery products, wine and beer), therefore an inhibitory activity would prevent lignin use in such products.

(Iberfluid Instruments S.A, Spain), for 30 min at 180 °C. The resulting liquid and solid fractions were separated by vacuum filtration. The liquid fraction was evaporated using a Rotavapor® R-215 (BÜCHI Labortechnik AG) up to approximately 50% w/w in total dissolved solids and then, the dissolved lignin was recovered by precipitation, by adding two volumes of cold water acidified at pH 2 with H2SO4). The precipitate was vacuum filtered, washed twice with 100 mL acidified water and dried at 60 °C for 24 h, obtaining the HL sample. The organosolv treatments were carried out in the same reactor used for autohydrolysis with the same 1:10 S/L ratio and using a solvent-water ratio of 60:40 v/v. The raw material was fractionated 75 min at 160 °C in the acetic acid-water treatment, and 90 min at 180 °C in the ethanol-water treatment. The lignin samples (AL and EL) were recovered as reported for HL. In the soda hydrolysis process, the raw material was treated with an aqueous solution of soda (7.5% w/w) with a S/L ratio of 1:10 in a glass atmospheric reactor of 25 L, at 90 °C for 90 min. The resulting liquid fraction (separated as reported for the HL) was submitted to an ultrafiltration process, similar to that described by Toledano et al. (Toledano et al., 2010) using ceramic membranes with two different cut–offs, 150 and 50 kDa (IBMEM – Industrial Biotech Membranes, Frankfurt, Germany). For this study the alkaline liquid fraction containing lignin molecules of size between 50 kDa and 150 kDa was collected and used. The lignin fraction (SL) was then isolated by acidification with sulphuric acid to pH 2. The resulting precipitate was subsequently recovered by vacuum-filtration, washed and dried at 60 °C, obtaining the SL sample. All the isolated lignin samples were submitted to a purification process in order to dissolve the lignin-linked hemicelluloses impurities and therefore to obtain only the acid insoluble lignin (AIL) fraction. A strong acid hydrolysis treatment (S/L ratio 1:10, 72% v/v H2SO4, 30 °C, 1 h) followed by a mild acid hydrolysis (dilution up to 4% v/v H2SO4, 100 °C, 4 h) was then carried out. After treatment, the acid insoluble lignin fraction was collected by vacuum filtration and oven-dried (50 °C for 24 h). These lignin samples are referred to as purified lignin: PHL, PAL, PEL and PSL. 2.2. Physicochemical characterization of lignin samples 2.2.1. Chemical composition The lignin samples were chemically characterized, determining moisture, ash, acid insoluble lignin (AIL), acid soluble lignin (ASL) and hemicellulosic sugars (glucose, xylose and arabinose) contents, according to the methodology described in a previous work (García et al., 2012) based on the work reported by Gosselink et al. (Gosselink et al., 2004). Moisture and inorganic matter contents were thermogravimetrically quantified by TGA analysis (TGA/SDTA RSI analyzer of Mettler Toledo) with a heating rate of 10 °C/min from 35 to 800 °C in air oxidizing atmosphere. The AIL was obtained following the same acid hydrolysis process followed to obtain the so called purified lignin samples and gravimetrically determined. The hydrolysate liquid fraction from the AIL was spectrophotometrically analyzed in a Jasco V-630 spectrophotometer measuring its absorbance at 205 nm for acid soluble lignin (ASL) determination according to the procedure described in literature (García et al., 2012; Gosselink et al., 2004). Glucose (GLU), xylose (XYL) and arabinose (ARA) concentrations in the hydrolysates were measured by high performance liquid chromatography (HPLC) in a Jasco LC-Net III-ADC chromatograph, equipped with a ROA Organic Acid column (00H-0138-K0, Phenomenex) and using 0.005 N H2SO4 as mobile phase (0.35 mL/min flow, 40 °C, injection volume 20 μL). High purity glucose, xylose and arabinose (≥99%, Fluka) were used for the calibration procedure.

2. Materials and methods Lignin samples studied in the present work were obtained from apple tree pruning wastes by means of different pre-treatments and fractionation methods. After an exhaustive characterization, several cultures were tested in order to evaluate the antimicrobial activity of the samples. In the following sections, extraction, characterization and testing methods are full described. 2.1. Lignin obtaining processes In this study apple tree pruning residues, kindly supplied by an independent farmer from Gurutze (Oiartzun, Spain), were used as raw material. After drying and grounding, the chemical composition of the raw material (moisture, ash, extractives, lignin, cellulose and hemicelluloses contents) was determined according to TAPPI standards (Serrano et al., 2010). The raw material was treated by four different fractionation processes (Amendola et al., 2012; García et al., 2010; Serrano et al., 2010; Toledano et al., 2010). In the auto-hydrolysis process, raw material was mixed with water (at a solid to liquid ratio (S/L) of 1:10) and treated in a reactor of 4 L

2.2.2. Chemical structure The chemical structure and the amount of functional groups in the lignin samples were evaluated by attenuated-total reflectance infrared spectroscopy (ATR-IR), performed by direct transmittance in a single243

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reflection ATR System (ATR top plate fixed to an optical beam condensing unit with ZnSe lens) with a MKII Golden Gate SPECAC instrument. Spectra were recorded over 32 scans with a resolution of 4 cm−1 in a wavelength range between 4000 and 600 cm−1. Significant changes in lignin structure due the different obtaining process were evaluated by proton nuclear magnetic resonance analyses (1H NMR). The spectra were recorded on a Bruker Avance 500 MHz at 25 °C. The spectral width and the acquisition time were 10,000 Hz and 3.1719 s respectively. For the analysis, lignin samples were dissolved in DMSO–d6 (Xu et al., 2007). 1H NMR was used for the evaluation of functional groups and regions, and comparison of the intensity of these clusters between lignin samples obtained by different methods. The signal intensity estimation was referred to protons in aromatic structures (δ 6.0–8.0 ppm), protons in methoxy groups (δ 3.0–4.5 ppm), to protons of acetyl groups in aromatic and aliphatic structures (δ 2.5–2.2 ppm and δ 2.2–1.9 ppm, respectively) and to protons in aliphatic structures (δ 0.5–1.5 ppm) (Serrano et al., 2010). The relative amount of each cluster was determined after normalization of its integral respect to that corresponding to the aromatic region (δ 6.0–8.0 ppm). Pyrolysis-Gas Chromatography-Mass Spectroscopy analyses (Pyr–GC/MS) were carried out using a commercial pyrolyzer (Pyroprobe model 5150, CDS Analytical Inc., Oxford, PA) (Erdocia et al., 2016). The sample (< 1 mg) was pyrolyzed in a quartz boat between two pieces of rock wool at 600 °C for 15 s, with a heating rate of 2 °C/ms and with the interface kept at 300 °C. The pyrolyzates were purged from the pyrolysis interface into the GC (7890A)-MS (5975C inert MSD with Triple-Axis Detector) Agilent instrument equipped with an Agilent 19091S-433 HP-5MS capillary column (30 m × 250 μm × 0.25 μm). Helium (3 mL/min) was used as the carrier gas, with a split ratio set to 1:50. The oven temperature program started at 50 °C, held for 2 min. Then, the temperature was raised to 120 °C (10 °C/min, held for 5 min), then to 280 °C (10 °C/min, held for 8 min) and finally to 300 °C (10 °C/min, held for 10 min). The identification of the products from the obtained pyrograms was accomplished comparing the obtained mass spectra with the mass spectra of National Institute of Standards Library (NIST). Compounds retained between 2 and 23 min and abundance higher than 0.50% were considered for identification.

CAT and TX. The specific AOP was calculated with reference both to lignin mass and TPC as:

AOPlignin = AOP/μglignin

AOPTPC = AOP/μgGAE

2.3. Antimicrobial activity 2.3.1. Inhibition of Aspergillus niger The antifungal activity of the samples was assessed based on the inhibition observed in the growth of a strain of A. niger. The fungal culture was pre-grown on malt agar (CM0059, Oxoid) and the spores were collected in sterile water, containing 0.3% v/v of Tween 80®. The spores were counted at the optical microscope by means of Burker chamber and, then, two fungi concentrations (103 and 102 spores/mL) were prepared. Lignin samples and antioxidant reference compounds (GA, CAT and TX) were respectively finely dispersed and dissolved in sterile water at two testing concentrations (5000 and 500 ppm). For each test, 1 mL of fungi solution and 1 mL of sample were mixed with 8 mL of malt extract agar in Petri plates (Ø 90 mm). Two control plates (only with the fungi solutions) were also tested. The plates were kept at 24 °C for 120 h and afterwards number of grown colonies and morphological aspect were evaluated.

2.3.2. Inhibition of Saccharomyces cerevisiae A spectrophotometric method (Serrano et al., 2010) was used to easily determine the concentration of colony forming units (CFU/mL) in liquid cultures of Saccharomyces cerevisiae. For this purpose, a calibration curve was constructed from different dissolutions of an initial solution of commercial baking yeast (Mastro Fornaio, Paneangeli, Italy) in malt agar extract broth previously activated for 24 h. For each yeast dissolution the CFU/mL (between 105 and 107 CFU/mL) was evaluated with optic microscope, and the corresponding absorbance or optical density at 660 nm (OD660) was read against agar broth in a UV-1601 Shimadzu spectrophotometer. The obtained linear correlation CFU/ mL = 1.169·107xOD660 had a R2 of 0.999. The lignin samples and the reference antioxidant compounds (GA, CAT and TX) were dissolved in DMSO at two concentrations (10,000 and 5000 ppm). For the test, 1 mL of sample was added to 20 mL of agar broth and the OD660 was measured against agar broth. Afterwards, 200 μL of yeast inoculum was added, with a final concentration of 2·104 CFU/mL in each culture. A blank (with 1 mL of DMSO) and a control (without the sample) were included. The samples were then incubated at 24 °C and the OD660 was read at different times (0, 2, 4, 6, 22, 24, 27, 30 and 48 h) always against agar broth According to literature, the microbial growth often shows a lagphase (λ) in which the specific growth rate begins at 0 and then increases up to a maximum value μmax. In a final phase, growth decreases and becomes 0 reaching an asymptote (A). When the growth curve is defined as the logarithm of number of microorganisms vs. time, a sigmoid curve is obtained (Franden et al., 2009; Toussaint et al., 2006) and the Gompertz equation modified by Zwietering et al. (1990) can be used to describe microbial growth data and determine the growth parameters:

2.2.3. Molecular weight distribution Gel permeation chromatography (GPC) was used to determine the average molecular weight (Mw) and polydispersity (Mw/Mn) of the obtained lignin samples. N,N-dimethylformamide (DMF) was used as mobile phase (Gosselink et al., 2004), with a flow rate of 0.7 mL/min at 40 °C, in a Jasco Inc. chromatograph provided with a LC-NetII/ACD interface, a column oven CO-2065Plus and a RI-2031Plus Intelligent Refractive Index Detector, a guard column and two columns PolarGel-M (Varian Inc.). Calibration was made using polystyrene standards (Mp from 250 to 70,000) purchased from Sigma Aldrich. 2.2.4. Total phenolic content and antioxidant power The total phenolic content (TPC) in the lignin samples was determined by the Folin-Ciocalteu (FC) method (Amendola et al., 2012) using gallic acid (> 98% HPLC, Fluka) as reference compound and dimethyl sulfoxide (DMSO > 99.5% GC, Sigma) as solvent (Bhat et al., 2009). For the analysis, samples were dissolved (2 g/L) in DMSO. The TPC was expressed as percentage of gallic acid equivalents (% GAE w/w). For comparison purposes, GAE content was also determined for Trolox® (TX) (97%, Aldrich) and (+)-catechin hydrate (CAT) (98% HPLC, Fluka). The antioxidant power (AOP) of the analyzed lignins was determined according to the ABTS (2,2′-azino-bis(3-ethylbenzothiazoline-6sulphonic acid)) spectrophotometric assay described in a previous work (García et al., 2012). The results were expressed as percentage of the ABTS radical reduction. AOP was determined also for gallic acid (GA),

log(ODt /ODo) = A⋅exp{ − exp[(μmaxe/A)(λ − t) + 1]} where ODt is the optical density at time t and OD0 the optical density at the beginning, e is a constant (2.72). The parameters A, λ and μm were graphically determined to evaluate the sensitivity of yeast to the compounds contained in the culture media. 244

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3.2.2. Chemical structure The ATR-IR spectra of the analysed lignin samples showed typical bands associated to functional groups in lignin (Boeriu et al., 2004; Negi, 2012; Vanderghem et al., 2011) (labelled with letters in Fig. 1). The signal related to hydroxyl group (a) appeared around 3300 cm−1, and those for methyl and methylene groups (b) at 2940, 2850 and 1460 cm−1. In the carbonyl region, the band related to C]O stretching (c) was found at 1700 cm−1. Typical aromatic skeleton vibrations (d) of lignin were appreciated at 1600, 1515 and 1420 cm−1, and the band arising from OH bending in phenolic structures (e) was found at 1370 cm−1. All spectra showed characteristic vibrations in lignin and associated to syringyl (f) (1325, 1215 and 830 cm−1) and guaiacyl units (g) (1265 and 1030 cm−1), as well as bands associated to OH vibrations in primary and secondary alcohols (h) (1150, 1040 cm−1), the later originated by carbohydrates, indicating contamination due to polysaccharides presence in the lignin samples. In Fig. 1 it can be observed that the intensity of the bands varied significantly between the spectra of the lignin samples obtained by different fractionation processes. The profiles of HL, EL and AL appeared rather similar regarding the intensity of the signals, whereas the SL spectrum presented noticeable differences compared to the others. HL, EL and AL showed high intensity of the aromatic ring vibration and a visible band associated to the phenolic group, while in SL the triplet associated to aromatic structure of lignin was much lower, the peak at 1370 cm−1 was unnoticeable, and the relative intensity of the peak around 1700 cm−1 resulted higher. This could be related to oxidation mechanisms of the lignin structure during the soda process (Boeriu et al., 2004; Łojewski et al., 2010) which could attack the aromatic ring and promote formation of carbonyl, carboxyl and quinone groups (Mancera et al., 2010). Moreover, the intensity of the band around 1040 cm−1 resulted more visible in SL than in the other samples, indicating that the alkaline lignin contained more carbohydrate impurities than the autohydrolysis or organosolv ones and confirming the results shown in Table 1. On the other hand, the isolation of the acid insoluble lignin fractions (the so called purified samples) had a marked effect on the alkaline sample, with an increase of the signals associated to the aromatic ring (at 1600, 1517 and 1420 cm−1) and a decrease in the carbohydrates influence in the PS spectrum (band at 1040 cm−1), which indicated polysaccharides removal. However, also the less hemicellulose contaminated lignin samples resulted affected by the acid-hydrolysis isolation process, showing a variation in the intensity of some bands when compared with the corresponding initial samples. For instance, a slight increase of the carbonyl signal (around 1700 cm−1) was observed, as well as a weak decrease of the aromatic ring triplet. The strong peak at 1215 cm−1 in HL, EL and AL spectra, associated to CeC plus CeO plus C]O stretching in the aromatic structure, resulted less intense after acid hydrolysis. This suggested that the applied acid treatment affected the lignin structure, increasing the amount of carbonyl groups and causing the modification of eOCH3 groups in the aromatic ring and lignin side chain. A rather similar behaviour, as well as a broadening of spectra below 1000 cm−1, was reported by

2.4. Statistics The lignin isolation processes were carried out in duplicate and the chemical analyses were carried out at least in duplicate. The results are reported as mean values ± s.d. The influence of the lignin type on the lignin properties was assessed by one-way analysis of variance (ANOVA) using the software STATGRAPHICS centurion XV, version 15.2.11 (StatPoint, Inc.). Differences at p < 0.05 were considered significant. In case of significant difference, the means were discriminated applying the post-hoc Tukey multiple range test.4.

3. Results 3.1. Lignin yield of the extraction processes After characterization, it was found that apple tree pruning residues was composed (in % w/w on dry basis) by 30.60 ± 2.40 of cellulose, 32.24 ± 0.60% of hemicelluloses, 17.00 ± 3.41% of lignin, 1.09 ± 0.13% of extractives and 9.28 ± 0.21% of inorganics. Each applied extraction process yielded differently in terms of isolated sample. After autohydrolysis 1.4 g of AH sample/100 g of raw material were obtained; from organosolv processes 10.1 g of AL and 7.9 g of EL were obtained per 100 g of raw material, and 9.1 g/ 100 g were obtained by alkaline pretreatment, from which only 1.3 g corresponded to the tested SL sample (from the ultrafiltration using 150 and 50 kDa UF membranes). The chemical composition of these samples (AH, AL, EL and SL) was later determined.

3.2. Physicochemical properties of lignin 3.2.1. Chemical composition The chemical composition of the analysed lignin samples resulted strongly influenced by the treatment applied for their isolation, as can be seen in Table 1. This effect has been previously reported by other authors (Ammar et al., 2014; Boeriu et al., 2004; Gosselink et al., 2004). In the present work, the lignin samples obtained by hydrolysis (HL), ethanol-water (EL) and acetic acid-water (AL) treatments contained low inorganic matter and relatively high AIL. In agreement with the literature (Vanderghem et al., 2011), different carbohydrates contents were found for the samples obtained by different fractionation processes. The AL, HL and EL were the less hemicelluloses contaminated sample (presence of xylose, arabinose and glucose due to the hydrolysis of hemicelluloses), whereas SL was the most contaminated sample. These results were expected, since the fractionation of lignocellulose under neutral-acidic media results in intensive hydrolysis of lignincarbohydrate bonds (Vanderghem et al., 2011) while alkaline fractionation mainly promotes lignin structural alterations with less solvation effect on hemicelluloses (Brodeur et al., 2011). The alkaline lignin (SL) was found to be the more contaminated by impurities. All the analysed lignin samples contained low amount of acid soluble lignin ranging from less than 1% for EL and AL, to almost 2.5% for HL and SL.

Table 1 Chemical composition (in % wt on dry matter.) of the different lignin samples obtained in this work. HL: autohydrolysis lignin; EL: ethanol-organosolv lignin; AL: acetic acid-organosolv lignin; SL: soda lignin. n/d: not detected. Same letter indicates means are not statistically different (according to ANOVA and Tukey HSD multiple range test, P < 0.05). Composition (% wt.)

Inorganic matter Acid insoluble lignin Acid soluble lignin Glucose Xylose Arabinose

Sample HL

EL

AL

SL

1.03 ± 0.12 a 87.37 ± 1.07 c 2.28 ± 0.13 b 2.47 ± 0.46 b 2.32 ± 0.35 b 0.70 ± 0.02 c

2.72 ± 0.19 b 64.50 ± 4.73 b 0.92 ± 0.04 a 2.74 ± 0.45 b 1.80 ± 0.29 ab 0.46 ± 0.02 b

2.97 ± 0.10 b 82.58 ± 1.03 c 0.96 ± 0.02 a 0.80 ± 0.02 a 1.20 ± 0.01 a n/d a

14.88 ± 0.51 c 56.48 ± 2.06 a 2.60 ± 0.06 c 1.30 ± 0.01 a 15.62 ± 0.33 c 1.11 ± 0.01 c

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Fig. 1. ATR-IR spectra, wide range on the left (from 4000 to 800 cm−1) and magnified on the right (from 1800 to 800 cm−1), of autohydrolysis lignin (HL), purified autohydrolysis lignin (PHL), ethanol-water lignin (EL), purified ethanol-water lignin (PEL), acetic acid-water lignin (AL), purified acetic acid-water lignin (PAL), soda lignin (SL) and purified soda lignin (PSL). Letters refer to different functional groups: (a) hydroxyl, (b) methyl and methylene, (c) C]O stretching, (d) aromatic skeleton, (e) OH bending in phenolic structures, (f) syringyl, (g) guaiacyl and (h) OH in primary/secondary alcohols.

Thus, the relative contents of aromatic and aliphatic acetyl groups (δ 2.2–2.5 and 1.9–2.2 ppm, respectively) resulted 0.42%, 0.55%, 0.33% and 0.51%, and 0.08%, 0.26%, 0.37% and 0.86% for HL, EL, AL and SL, respectively. The amount of methoxy groups (δ 3.0–4.5 ppm) resulted higher for SL (17.98%), whereas EL, HL and AL showed lower relative content of eOCH3 moiety (11.56%, 5.86% and 4.71%, respectively). Similarly, the SL was that with the highest aliphatic/aromatic ratio compared to EL, AL and HL (8.49% against 2.29%, 0.78% and 0.43%, respectively). These relative abundances, of both methoxy groups and aliphatic moieties (δ 0.5–1.5 ppm), were highly related with the carbohydrate content, increasing with the presence of hemicellulose impurities. After the acid hydrolysis treatment the relative intensities of the aromatic cluster significantly decreased for PHL and PAL (5.35% and 4.81%, respectively), while increased for PEL and PSL (12.69% and 5.41%, respectively). Furthermore, the aliphatic/aromatic ratios resulted higher for autohydrolysis and acetosolv lignins after purification (0.55% and 2.31%, respectively) but considerably lower for ethanol and soda extracted samples (1.20% and 5.40%, respectively). These results confirmed that the acid treatment was effective in the removal of aliphatic moieties (hemicellulose impurities) but also caused certain degradation of the aromatic structure. All the analyzed samples, in fact, showed a decrease of the relative intensity of the signal between 3.0 and 4.0 ppm, associated to the protons in methoxy group (3.86%, 1.58%, 4.63% and 1.97% for PHL, PEL, PAL and PSL respectively), and a sharp peak in the range from 4.0 to 4.5 ppm, more specifically at 4.05, 4.19, 4.28 and 4.45 ppm for PHL, PEL, PAL and PSL, respectively. Several authors reported that, in the region from 4.0 to approximately 5.0 ppm, the signals related to protons in β-O-4 inter-lignin units

Mancera et al. (Mancera et al., 2010) during the oxidation of lignin with hydrogen peroxide, which was associated with depolymerization and repolymerization mechanisms of lignin in strong basic or acid conditions. Through 1H NMR analysis, the chemical structure of the lignin samples was roughly compared by the variation observed for the intensity of the different spectral regions (spectra shown in Fig. 2). After estimation of relative intensity of the aromatic cluster (δ between 6.0 and 8.0 ppm) with respect to the integral of the whole analysed region of each lignin spectra (δ 0.5–8.0 ppm), the highest aromatic content was found for HL, AL and EL (12.24%, 12.19% and 6.07%, respectively), whereas SL sample showed a low relative intensity of 3.00% related to the presence of aromatic protons (see Table 2). The less intensity of the aromatic ring in the alkaline sample corroborated the results from the IR spectroscopy analyses and the chemical composition results related to its higher impurity content. Regarding the acid insoluble lignin fractions, the PEL showed the highest aromatic relative intensity (12.69%), followed by PSL, PHL and PAL (5.41%, 5.35% and 4.81%, respectively). These results indicated that the acid hydrolysis was effective in terms of hemicelluloses removal since the relative aromatic content increased almost twice for the alkaline sample. The less relative intensity of the aromatic region in the other samples again suggests partial modification of lignin structure due to the severe conditions of the treatment, which might have led to degradation of the aromatic ring. On the other hand, other functional groups (Fig. 2) were evaluated as the relative intensity of their corresponding proton signal respect to those determined for the aromatic group (δ between 6.0 and 8.0 ppm). 246

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Fig. 2. 1H NMR spectra of the analyzed lignin samples: autohydrolysis lignin (HL), purified autohydrolysis lignin (PHL), ethanol-water lignin (EL), purified ethanol-water lignin (PEL), acetic acid-water lignin (AL), purified acetic acid-water lignin (PAL), soda lignin (SL) and purified soda lignin (PSL).

2014; Prado et al., 2016). A total of 36 structures were identified (results shown in Table S1). Besides others, compounds 2 (furfural) and 25 (levoglucosan) clearly indicate the presence of polysaccharide impurities (xylose and glucose) (Ralph and Hatfield, 1991). These compounds were detected for HL and SL, supporting the chemical characterization results reported above, and disappeared after the acid hydrolysis of the samples, thus confirming the effectiveness of the applied purification. The main pyrolysis phenolics detected for each sample were: 29 (1-(3,4-dimethoxyphenyl)- ethanone) for HL (9.93%),

linkages are displayed (Sun et al., 2011; Vanderghem et al., 2011; Xu et al., 2007). Thus, according to the obtained 1H NMR chromatograms, it can be assumed that under the strong acidic conditions used for the isolation of acid insoluble lignin, not only carbohydrate dissolution should occur, but also self-condensation between lignin units. Pyr-GC mass spectra of the obtained lignin samples are shown in Fig. 3. This technique allowed evaluating not only the presence of carbohydrates in the samples but also the predominance of certain lignin structures due to the detection of pyrolysis products (Kim et al., 247

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Table 2 Relative abundance of aromatic protons (H Ar in %), methoxy groups (eOCH3 in %), proton Aliph/Ar ratio (H Aliph/Ar), S, G and H-units (%), syringyl/guaiacyl/hydroxyphenyl ratio (S/ G/H), average molecular weight (Mw), polydispersity (Mw/Mn), total phenolic content (TPC in % wt. gallic acid equivalents, GAE), specific antioxidant power per sample mass (AOP/μg lignin) and per total phenolic content (AOP/μgGAE) of the isolated lignin samples HL: autohydrolysis lignin; EL: ethanol-organosolv lignin; AL: acetic acid-organosolv lignin; SL: soda lignin. P: purified sample. Same letter indicates means are not statistically different (according to ANOVA and Tukey HSD multiple range test, P < 0.05). Lignin Sample

1

H Ar eOCH3 2 H Aliph/Ar 2 S-units 2 G-units 2 H-units 2 S/G/H Mw Mw/Mn TPC AOP/μglignin AOP/μgGAE 1

1 2

HL

PHL

EL

PEL

AL

PAL

SL

PSL

12.24 5.86 0.43 20.2 27.5 7.11 3:4:1 4263 ± 33a 2.09 ± 0.13b 38.13 ± 1.73d 2.28 ± 0.25bc 5.97 ± 0.17ab

5.35 3.86 0.55 9.58 16.1 3.31 3:5:1 24606 ± 150e 5.11 ± 0.16d 23.33 ± 1.30b 1.27 ± 0.30abc 5.44 ± 0.31ab

6.07 11.56 2.29 19.6 25.08 9.51 2:3:1 8810 ± 85c 3.15 ± 0.21c 30.33 ± 0.88c 2.23 ± 0.34bc 7.34 ± 0.53b

12.69 1.58 1.20 8.04 9.66 2.72 3:4:1 16305 ± 148d 5.20 ± 0.14d 13.39 ± 1.82a 0.99 ± 0.41ab 7.38 ± 0.86b

12.19 4.71 0.78 22.41 30.8 8.91 3:3:1 7175 ± 78b 3.56 ± 0.08c 33.99 ± 1.56 cd 2.44 ± 0.31c 7.17 ± 0.38b

4.81 4.63 2.31 13.57 20.65 5.26 3:4:1 7205 ± 28b 6.64 ± 0.14e 12.64 ± 0.93a 0.91 ± 0.56ab 7.20 ± 0.28b

3.00 17.98 8.49 6.68 6.98 3.4 2:2:1 16305 ± 148d 1.53 ± 0.04a 15.13 ± 0.74a 0.97 ± 0.35ab 6.38 ± 0.54bc

5.41 1.97 5.40 2.33 5.74 1.65 1:3:1 4192 ± 59a 2.35 ± 0.07b 12.43 ± 1.93a 0.55 ± 0.10a 4.40 ± 0.57a

determined as relative content by clusters integration of proton 1H NMR spectra. determined as relative content of S, G and H compounds identified by Pyr-GC/MS (see Table S1).

PHL (5.01%), AL (8.80%) and PAL (5.68%), 23 (3-Methoxy-5-methylphenol) for EL (7.46%), 20 (2,6-dimethoxyphenol or syringol) for PEL (2.75%) and 11 (2-methoxy-5-methylphenol or methylguaiacol) for PSL (1.62%). The principal product detected for SL sample (5.27%) was (6) 2,3-dimethylpentanal, an aliphatic chain resulted from the thermal degradation of carbohydrates (Ralph and Hatfield, 1991). The great difference found on the pyrolysis compounds abundance places on record that pyrolysis products depend strongly on the lignin origin but also on the isolation process (Lin et al., 2015). The classification of detected pyrolysis phenolic products according to S, G or H-type units (Kim et al., 2014) allowed to determine the primary structures (Table 2 and Supplementary data, Table S1). All the analyzed lignins presented high abundance of G-type units (30.8–6.98% for the original lignins and 20.65–5.74% for the purified ones),

followed by S-type units (between 22.41% and 2.33%). The ratio S:G:H varied between samples (HL 3:4:1, PHL 3:5:1, EL 2:3:1, PEL 3:4:1, AL 3:3:1, PAL 3:4:1, SL 2:2:1 and PSL). In general, the applied purification method led to a decrease in the abundance of phenolic structures as the severity of the acid hydrolysis promoted partial oxidation of the aromatic structure of lignin. Interestingly, the purification method affected markedly the decline of H and S-units (up to 71% and 65%, respectively) whereas G-units resulted less degraded (between 18% and 60%). The higher abundance of phenolic moieties corresponded to those lignins obtained under more acid conditions. It has been reported in literature that fractionation with organic solvents allows obtaining fractions with higher content of these essential functional groups (Ponomarenko et al., 2015).

Fig. 3. Pyr-GC/MS chromatograms of the different analyzed lignin samples and structure of some identified compounds (see Supplementary materials, Table S1).

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3–6 times increase of average molecular weight and 1.5–2.5 times increase in polydispersity values found for the HL and EL after the purification stage. The fact that average molecular weight neither polydispersity of SL increase, could be due to the high hemicellulose contamination, suggesting that the acid hydrolysis in this case caused only the polysaccharide dissolution. According to this assumption and taking into account the high Mw that hemicelluloses generally have (Sun et al., 2011), it could be expected a decrease of both average molecular weight and polydispersity after the carbohydrate impurities removal, as Fig. 4d shows. Probably, a longer purification treatment might promote condensation mechanisms also in SL. 3.2.4. Total phenolic content and antioxidant power Total phenolic content (TPC) and antioxidant power of the lignin samples are reported in Table 2. HL, EL and AL showed higher and comparable TPC, even slightly higher than values reported in previous works for lignin obtained by similar procedures (Amendola et al., 2012; García et al., 2012). The lower value found for SL is in agreement with the TPC previously found for alkaline samples (García et al., 2010). The acid hydrolysis promoted on average a decrease of TCP, particularly for EL and AL for which it was reduced by more than 50%. The lowest reduction was found in SL. The specific antioxidant capacity of lignin (AOPlignin) almost followed the same trend of the TPC, suggesting a correlation between the two properties. Calculating the specific AOP of total phenols (AOPGAE), it was found that the organosolv lignin samples contained the most powerful phenolic compounds, followed by soda lignin and last by autohydrolysis lignin. Except for SL, the acid insoluble lignin fraction showed the same AOPGAE than the initial corresponding lignin, suggesting that the treatment reduced the total amount of phenolic compounds but not their antioxidant capacity. The reference antioxidant compounds (gallic acid, catechin and Trolox®) showed higher specific AOP than the lignin samples (33.03, 5.54 and 5.03 AOP/μg of reference and 33.03, 19.70 and 19.11 AOP/μgGAE, respectively). It is well known that the antioxidant activity does not only depend on the TPC but also on the presence of other compounds and on the chemical structure (Li et al., 2005), therefore it was expected that the isolation process would have influenced on AOP values. The high decrease (approx. 31.0%) of the specific phenolic AOP between SL and PSL, confirms that modification of the phenolic structure occurred for this sample, as previously commented. Ponomarenko et al. (2015) reported a clear influence of lignin structure on its antioxidant properties, asserting that not only phenolic groups are responsible of this antiradical feature. They concluded that the influence of o-methoxyl groups and the CH2 groups in the αposition of the aliphatic chain is positive, while the influence of the carbonyl groups in the aliphatic chains and of the large size of πconjugated systems results negative. In this sense, HL sample presented larger abundance of phenolic structures with a double bound or a carbonyl group in the side-chain of phenylpropane units (29, 31, 33 or 34 structures in Table S1). Thus, even if the highest TPC content (38.13% GAE) corresponded to HL sample, the effectiveness of its antiradical activity diminished due to its less scavenging structure. On the other hand, PLS sample with a lower than 3-times phenolic content (12.43% GAE) presented a quite similar AOP/μgGAE value to HL (4.40 and 5.97, respectively). This could be due to the larger relative amounts of phenolic structures with o-methoxy groups, as found after Pyr-GC/ MS analyses (e.g. structure 11 in Table S1).

Fig. 4. Gel permeation chromatograms of original and purified lignin samples obtained by (a) autohydrolysis, (b) ethanol-organosolv process, (c) acetic acid-organosolv process and (d) soda process. Table 3 Effect of lignin sample addition on the growth of A. niger. Results are expressed as difference in the number of colony forming units (CFU) per plate in comparison with control after 120 h incubation. (+): increment < 25 CFU/plate; (++) increment of 25–75 CFU/plate; (+++) increment > 75 CFU/plate; (−)reduction < 25 CFU/plate; (−−) reduction of 25–75 CFU/plate. HL: autohydrolysis lignin; EL: ethanol-organosolv lignin; AL: acetic acid-organosolv lignin; SL: soda lignin. P: purified sample; n/t: not tested. Lignin sample concentration

5000 ppm 500 ppm

Effect on A. niger growth HL

PHL

EL

PEL

AL

PAL

SL

PSL

– +++

n/t +++

+ ++

+ ++

++ ++

++ ++

– +

n/t ++

3.2.3. Molecular weight distributions The molecular weight distributions of the isolated lignin samples showed significant differences according to the used fractionation process and the subsequent acid hydrolysis step. As shown in Table 2, original soda lignin had the highest Mw value, followed by the organosolv lignin samples and finally by the autohydrolysis lignin. These results agreed with the Mw range reported in literature for lignin of different sources (Gosselink et al., 2004; Mancera et al., 2010; Serrano et al., 2010). The acid treatment markedly increased the Mw of HL and EL, confirming the previous hypothesis based on 1H NMR analysis, that self-condensation between lignin units might have occurred. However the Mw was not influenced by this step in the case of AL and, on the opposite, decreased for SL. In Fig. 4a–c, a broadening of the molecular profile and an attenuation of chromatogram intensity can be observed in the chromatograms of the AIL samples in agreement with the variation in the average molecular weight and polydispersity values summarized in Table 2 for HL, EL and AL. Several authors have claimed that under strong acid conditions, lignin could suffer first a depolymerization and subsequently self-condensation/repolymerization mechanisms after the formation of highly reactive and unstable carbocations (Funaoka, 2013; Mancera et al., 2010; Vanderghem et al., 2011). This would explain the

3.3. Antimicrobial activity 3.3.1. Inhibition of Aspergillus niger The lignin samples did not show a pronounced biocide effect against A. niger (Table 3). Indeed, at low concentration (500 ppm), lignin sample addition enhanced fungi growth probably due to the presence of 249

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Fig. 5. Visual appearance (after 120 h incubation at 24 °C) of malt agar plates inoculated with two initial Aspergillus niger inoculum (102 or 103 spores), without (CONTROL) and with the addition of 1 mL of antioxidant compound (TX: Trolox®; CAT: catechin; GA: gallic acid) or lignin sample (HL: autohydrolysis lignin; EL: ethanol-organosolv lignin; AL: acetic acidorganosolv lignin; SL: soda lignin; P: purified sample) at 5000 or 500 ppm.

presence decreases effectiveness against some bacteria. Interestingly, growth was not totally inhibited by the reference antioxidant compounds as well, as can be observed in Fig. 5. It was observed that the fungus growth resulted affected by the presence of lignin, in terms of morphological characters (ripeness and colouring of the colonies) while the reference antioxidant compounds (Trolox®, catechin and gallic acid) did not influence these aspects (Fig. 5). In particular, most of the cultures added with 1 mL of 5000 ppm lignin sample appeared less ripened and slightly coloured (with pale blue, green or yellow pigmentation), whereas the cultures added with 1 mL of 500 ppm lignin sample showed a normal growing with reference to control test samples. The inhibiting capacity of lignin

hemicelluloses and minerals, even though SL, the richest in hemicelluloses, caused the lower growth promotion. When a higher concentration was used, generally the positive effect on growth was reduced (except for AL) and some samples (HL and SL) exhibited an inhibitory effect. Unexpectedly, the acid insoluble lignin fractions (purified samples) did not show any different behaviour compared to the initial lignin and PHL and PSL could not tested at the highest concentration due to solubilisation problems. Hemicelluloses absence did not greatly affect colonies forming, even though some authors (Negi, 2012) reported a lower effectiveness for crude vegetal extracts compared to pure compounds, probably due to the fact that crude extracts generally contain phenolic compounds in glycosidic form, where the sugar

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insoluble lignin fraction led to a decrease in total phenolic content of the sample, but also to polymerization of lignin structure with reduction of the methoxy moieties. Therefore, it could be said that the isolated acid insoluble lignin samples were more easily tolerated by the A. niger strain, at least based on morphological aspects, since CFU number did not change (Table 3). 3.3.2. Inhibition of Saccharomyces cerevisiae The sensitivity of yeast to the isolated lignin samples and reference compounds was evaluated by analysing the growth parameters (Fig. 6) obtained from the growth curves elaborated according to Gompertz equation to allow a better interpretation and comparison of the results (see Supplementary Fig. S1). For these tests, DMSO was used as solvent for the lignin samples at a concentration of 5000 and 10,000 ppm, therefore the inhibition effect of DMSO was also assessed. It was observed in Fig. 6 that DMSO inhibited the yeast leading to a slightly lower growth, compared with the control. A similar effect was observed for, the reference antioxidants (Trolox®, catechin and gallic acid), that induced a slight inhibition on the yeast in terms of maximum growth rate (μmax) (Fig. 6c), while the final maximum growth was not significantly different (Fig. 6a) and the lag time was actually reduced (Fig. 6b). The addition of different lignin samples induced both a lower and slower growth. All the tested samples induced a decrease of yeast population (from 25.8 to 78.7% lower value of maximum growth in Fig. 6a) and a marked delay of the beginning of the exponential growth phase (1–4 times longer lag time than control in Fig. 6b). Moreover, the calculated μmax was definitely lower (by 72.8–94.3%) in the presence of lignin samples than in control culture, probably due to lignin structure and size (not nutritionally accessible by the yeast) or the formation of inhibitory compounds (Ghorai et al., 2009; Liu et al., 2007). In this sense, clear inhibitory differences were observed between lignin and reference compounds. Baurhoo et al. (Baurhoo et al., 2008) reported that, in general, phenolic components with functional groups containing oxygen (eOH, eCO, eCOOH) in the side chain are less inhibitory, whereas the presence of double bonds and methyl groups increases the biocide effect of phenolics. Thus, as lignin structure results more complex than that of the tested reference compounds (higher molecular weight and functionality), the isolated lignin samples exhibited higher biocide effect than the reference compounds.

Fig. 6. Growth parameters for Saccharomyces cerevisiae grown in 20 mL liquid malt agar mixed with 1 mL of DMSO or DMSO containing 5000 ppm (white) and 10,000 ppm (grey) of lignin. HL: autohydrolysis lignin; EL: ethanol-organosolv lignin; AL: acetic acidorganosolv lignin; SL: soda lignin; P: purified lignin sample. (a) Maximum growth, (b) lag time and (c) maximum specific growth rate.

against several microorganisms growth has been widely reported (Vermaas et al., 2015). This type of alteration was also observed by Rahouti et al. (Rahouti et al., 1999), who asserted that when some strains were grown in presence of phenolic substrates some physiological changes occurred (fructification changes, abnormal production of pigments or of viscous compounds). In this sense, the chemical structure of the phenolic compound (functionality and substitution degree in the aromatic ring) highly affected strains pigmentation and growth (Baurhoo et al., 2008; Ghorai et al., 2009; Rahouti et al., 1999). Fig. 5 illustrates the growth of A. niger cultures containing 5000 ppm of original lignin samples (HL, EL, AL and SL), and shows a clearer ripening delay and colour alterations than those containing only the acid insoluble lignin fractions (PHL, PEL, PAL and PSL). This behaviour could be related not only to the hemicelluloses removal but also to structural changes occurred during the process. In the work described by (Rahouti et al., 1999), microbial degradation of ligninmodel compounds was highly affected by their toxicity and microorganism tolerance, resulting easier as more hydrophilic the compound was, and with more fructification in the presence of free hydroxyl groups than in the presence of methoxy groups. Some authors (Baurhoo et al., 2008; Qin et al., 2016) reported phenolic components of lignin are capable to inhibit some enzymes behaviour and the growth of microorganisms such as Escherichia coli, Saccharomyces cerevisiae, Bacillus licheniformis and Aspergillus niger. These authors also commented that side chain structure and nature of the functional groups of the phenolic compounds are major determinants of the antimicrobial effects of lignin. In the present work, the analysis of the tested lignin samples revealed that isolation of the acid

4. Conclusions In the present work, different isolation processes (autohydrolysis, ethanol or acetic acid organosolv process, and alkaline hydrolysis) were applied to obtain lignin from apple tree pruning residues. Chemical characterisation showed a higher content in organic matter and hemicelluloses for the sample obtained with an alkaline hydrolysis. ATR-IR, NMR and Pyr-GC/MS analyses confirmed notable structural differences of the samples depending on the obtaining method as well as lower purity (carbohydrate content) in soda and autohydrolysis lignins. The subsequent acid hydrolysis purification applied to isolate the acid insoluble lignin fractions, was effective in removing the carbohydrate impurities but also caused a partial degradation (condensation mechanism) of the aromatic structure. Evaluation of total phenolic content showed the highest content for the autohydrolysis lignin and the lowest content for the alkaline lignin. In general the acid purification process caused also a reduction in the phenolic content. However, based on the antioxidant capacity (estimated as antiradical activity against the ABTS radical), all the samples revealed statistically comparable specific power per mass of sample. Antimicrobial activity against an environmental contaminant fungus (A. niger) and a food-technology yeast (S. cerevisiae) was finally assessed. If tested at low concentration, all the samples actually showed a positive effect on the growth of A. niger. Increasing the tested concentration, inhibitory properties were observed only for the auto251

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reactfunctpolym.2012.05.010. Network Polymers towards High Performance Materials. García, A., Toledano, A., Andrés, M.Á., Labidi, J., 2010. Study of the antioxidant capacity of Miscanthus sinensis lignins. Process Biochem. 45, 935–940. http://dx.doi.org/10. 1016/j.procbio.2010.02.015. García, A., González Alriols, M., Labidi, J., 2012. Evaluation of the effect of ultrasound on organosolv black liquor from olive tree pruning residues. Bioresour. Technol. 108, 155–161. http://dx.doi.org/10.1016/j.biortech.2012.01.010. Ghorai, S., Banik, S.P., Verma, D., Chowdhury, S., Mukherjee, S., Khowala, S., 2009. Fungal biotechnology in food and feed processing. Food Res. Int. 42, 577–587. http://dx.doi.org/10.1016/j.foodres.2009.02.019. Gosselink, R.J.A., Abächerli, A., Semke, H., Malherbe, R., Käuper, P., Nadif, A., van Dam, J.E.G., 2004. Analytical protocols for characterisation of sulphur-free lignin. Ind. Crops Prod. 19, 271–281. http://dx.doi.org/10.1016/j.indcrop.2003.10.008. Hollman, P.C.H., 2001. Evidence for health benefits of plant phenols: local or systemic effects? J. Sci. Food Agric. 81, 842–852. http://dx.doi.org/10.1002/jsfa.900. Kim, J.-Y., Hwang, H., Park, J., Oh, S., Choi, J.W., 2014. Predicting structural change of lignin macromolecules before and after heat treatment using the pyrolysis-GC/MS technique. J. Anal. Appl. Pyrolysis 110, 305–312. http://dx.doi.org/10.1016/j.jaap. 2014.09.020. Li, J., Ding, S., Ding, X., 2005. Comparison of antioxidant capacities of extracts from five cultivars of Chinese jujube. Process Biochem. 40, 3607–3613. http://dx.doi.org/10. 1016/j.procbio.2005.03.005. Lin, X., Sui, S., Tan, S., Pittman, C.U., Sun, J., Zhang, Z., 2015. Fast pyrolysis of four lignins from different isolation processes using Py-GC/MS. Energies 8, 5107–5121. http://dx.doi.org/10.3390/en8065107. Liu, M., Seidel, V., Katerere, D.R., Gray, A.I., 2007. Colorimetric broth microdilution method for the antifungal screening of plant extracts against yeasts. Methods San Diego Calif. 42, 325–329. http://dx.doi.org/10.1016/j.ymeth.2007.02.013. Mancera, A., Fierro, V., Pizzi, A., Dumarçay, S., Gérardin, P., Velásquez, J., Quintana, G., Celzard, A., 2010. Physicochemical characterisation of sugar cane bagasse lignin oxidized by hydrogen peroxide. Polym. Degrad. Stab. 95, 470–476. http://dx.doi. org/10.1016/j.polymdegradstab.2010.01.012. Mikulásová, M., Kosíková, B., 2003. Modulation of mutagenicity of various mutagens by lignin derivatives. Mutat. Res. 535, 171–180. Negi, P.S., 2012. Plant extracts for the control of bacterial growth: efficacy, stability and safety issues for food application. Int. J. Food Microbiol. 156, 7–17. http://dx.doi. org/10.1016/j.ijfoodmicro.2012.03.006. Ponomarenko, J., Dizhbite, T., Lauberts, M., Volperts, A., Dobele, G., Telysheva, G., 2015. Analytical pyrolysis – a tool for revealing of lignin structure-antioxidant activity relationship. J. Anal. Appl. Pyrolysis 113, 360–369. http://dx.doi.org/10.1016/j. jaap.2015.02.027. Prado, R., Brandt, A., Erdocia, X., Hallet, J., Welton, T., Labidi, J., 2016. Lignin oxidation and depolymerisation in ionic liquids. Green Chem. 18, 834–841. http://dx.doi.org/ 10.1039/C5GC01950H. Qin, L., Li, W.-C., Liu, L., Zhu, J.-Q., Li, X., Li, B.-Z., Yuan, Y.-J., 2016. Inhibition of ligninderived phenolic compounds to cellulase. Biotechnol. Biofuels 9, 70. http://dx.doi. org/10.1186/s13068-016-0485-2. Rahouti, M., Steiman, R., Seigle-Murandi, F., Christov, L.P., 1999. Growth of 1044 strains and species of fungi on 7 phenolic lignin model compounds. Chemosphere 38, 2549–2559. http://dx.doi.org/10.1016/S0045-6535(98)00462-7. Ralph, J., Hatfield, R.D., 1991. Pyrolysis-GC–MS characterization of forage materials. J. Agric. Food Chem. 39, 1426–1437. http://dx.doi.org/10.1021/jf00008a014. Serrano, L., Egües, I., Alriols, M.G., Llano-Ponte, R., Labidi, J., 2010. Miscanthus sinensis fractionation by different reagents. Chem. Eng. J. 156, 49–55. http://dx.doi.org/10. 1016/j.cej.2009.09.032. Sun, X.-F., Jing, Z., Fowler, P., Wu, Y., Rajaratnam, M., 2011. Structural characterization and isolation of lignin and hemicelluloses from barley straw. Ind. Crops Prod. 33, 588–598. http://dx.doi.org/10.1016/j.indcrop.2010.12.005. Toledano, A., García, A., Mondragon, I., Labidi, J., 2010. Lignin separation and fractionation by ultrafiltration. Sep. Purif. Technol. 71, 38–43. http://dx.doi.org/10. 1016/j.seppur.2009.10.024. Toussaint, M., Levasseur, G., Gervais-Bird, J., Wellinger, R.J., Elela, S.A., Conconi, A., 2006. A high-throughput method to measure the sensitivity of yeast cells to genotoxic agents in liquid cultures. Mutat. Res. 606, 92–105. http://dx.doi.org/10.1016/j. mrgentox.2006.03.006. Ugartondo, V., Mitjans, M., Vinardell, M.P., 2009. Applicability of lignins from different sources as antioxidants based on the protective effects on lipid peroxidation induced by oxygen radicals. Ind. Crops Prod. 30, 184–187. http://dx.doi.org/10.1016/j. indcrop.2009.03.001. Vanderghem, C., Richel, A., Jacquet, N., Blecker, C., Paquot, M., 2011. Impact of formic/ acetic acid and ammonia pre-treatments on chemical structure and physico-chemical properties of Miscanthus x giganteus lignins. Polym. Degrad. Stab. 96, 1761–1770. http://dx.doi.org/10.1016/j.polymdegradstab.2011.07.022. Vermaas, J.V., Petridis, L., Qi, X., Schulz, R., Lindner, B., Smith, J.C., 2015. Mechanism of lignin inhibition of enzymatic biomass deconstruction. Biotechnol. Biofuels 8, 217. http://dx.doi.org/10.1186/s13068-015-0379-8. Wijekoon, M.M.J.O., Bhat, R., Karim, A.A., 2011. Effect of extraction solvents on the phenolic compounds and antioxidant activities of bunga kantan (Etlingera elatior Jack.) inflorescence. J. Food Compos. Anal. 24, 615–619. http://dx.doi.org/10.1016/ j.jfca.2010.09.018. 8th International Food Data Conference: Quality Food Composition Data, Key for Health and Trade. Xu, F., Zhou, Q.-A., Sun, J.-X., Liu, C.-F., Ren, J.-L., Sun, R.-C., Curling, S., Fowler, P., Baird, M.S., 2007. Fractionation and characterization of chlorophyll and lignin from de-juiced Italian ryegrass (Lolium multifolrum) and timothy grass (Phleum pratense). Process Biochem. 42, 913–918. http://dx.doi.org/10.1016/j.procbio.2007.02.001. Zwietering, M.H., Jongenburger, I., Rombouts, F.M., van’t Riet, K., 1990. Modeling of the bacterial growth curve. Appl. Environ. Microbiol. 56, 1875–1881.

hydrolysis and the soda lignin. The purified lignin samples exhibited the same behaviour than the initial ones. The presence of lignin induced changes in the morphological characters of the fungus but, anyway, the results suggest it could not be exploited as natural biocide. On the other hand, all the lignin samples showed antimicrobial activity against S. cerevisiae, resulting in lower maximum specific growth, longer lag time and lower maximum final growth level, with an apparent correlation of the measured effect lignin size and functionality, since the samples with higher aromatic moiety and phenolic content were more effective against yeast growth. Therefore the isolated lignin samples could be used as food/feed antioxidant or functional ingredients, but not in products obtained through a fermentation process based on S. cerevisiae. Further analyses considering other microorganism (yeast and bacteria) should be carried out in order to better evaluate the biocide or fungistatic effect of lignins obtained from different sources and processes. Acknowledgments The authors would like to thank the Spanish Ministry of Economy, Industry and Competitiveness (contract Juan de la Cierva Incorporacion IJCI-2015-23168) and the Spanish Ministry of Science and Innovation together with the Italian Ministry of Instruction, University and Research (Italy-Spain Integrated Actions Program, project IT20090054), for financial support of this work. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.indcrop.2017.04.063. References Łojewski, T., Miśkowiec, P., Missori, M., Lubańska, A., Proniewicz, L.M., Łojewska, J., 2010. FTIR and UV/vis as methods for evaluation of oxidative degradation of model paper: DFT approach for carbonyl vibrations. Carbohydr. Polym. 82, 370–375. http://dx.doi.org/10.1016/j.carbpol.2010.04.087. Amendola, D., De Faveri, D.M., Egües, I., Serrano, L., Labidi, J., Spigno, G., 2012. Autohydrolysis and organosolv process for recovery of hemicelluloses, phenolic compounds and lignin from grape stalks. Bioresour. Technol. 107, 267–274. http:// dx.doi.org/10.1016/j.biortech.2011.12.108. Ammar, M., Khiari, R., Berrima, B., Belgacem, M.N., Elaloui, E., 2014. Isolation and characterization of lignin from stipa tenacissima l and phoenix dactylifera. Cellul. Chem. Technol. 48, 255–263. Ayala-Zavala, J.F., Vega-Vega, V., Rosas-Domínguez, C., Palafox-Carlos, H., VillaRodriguez, J.A., Siddiqui, M.W., Dávila-Aviña, J.E., González-Aguilar, G.A., 2011. Agro-industrial potential of exotic fruit byproducts as a source of food additives. Food Res. Int. 44, 1866–1874. http://dx.doi.org/10.1016/j.foodres.2011.02.021. Exotic Fruits: their Composition, Nutraceutical and Agroindustrial Potential. Baurhoo, B., Ruiz-Feria, C.A., Zhao, X., 2008. Purified lignin: nutritional and health impacts on farm animals—a review. Anim. Feed Sci. Technol. 144, 175–184. http:// dx.doi.org/10.1016/j.anifeedsci.2007.10.016. Bhat, R., Khalil, H.P.S.A., Karim, A.A., 2009. Exploring the antioxidant potential of lignin isolated from black liquor of oil palm waste. C. R. Biol. 332, 827–831. http://dx.doi. org/10.1016/j.crvi.2009.05.004. Boeriu, C.G., Bravo, D., Gosselink, R.J.A., van Dam, J.E.G., 2004. Characterisation of structure-dependent functional properties of lignin with infrared spectroscopy. Ind. Crops Prod. 20, 205–218. http://dx.doi.org/10.1016/j.indcrop.2004.04.022. 6th International Lignin Institute Conference. Brodeur, G., Yau, E., Badal, K., Collier, J., Ramachandran, K.B., Ramakrishnan, S., Brodeur, G., Yau, E., Badal, K., Collier, J., Ramachandran, K.B., Ramakrishnan, S., 2011. Chemical and physicochemical pretreatment of lignocellulosic biomass: a review. Enzyme Res. 2011, e787532. http://dx.doi.org/10.4061/2011/787532. Chun, S.-S., Vattem, D.A., Lin, Y.-T., Shetty, K., 2005. Phenolic antioxidants from clonal oregano (Origanum vulgare) with antimicrobial activity against Helicobacter pylori. Process Biochem. 40, 809–816. http://dx.doi.org/10.1016/j.procbio.2004.02.018. Erdocia, X., Prado, R., Fernández-Rodríguez, J., Labidi, J., 2016. Depolymerization of different organosolv lignins in supercritical methanol ethanol, and acetone to produce phenolic monomers. ACS Sustain. Chem. Eng. 4, 1373–1380. http://dx.doi. org/10.1021/acssuschemeng.5b01377. Franden, M.A., Pienkos, P.T., Zhang, M., 2009. Development of a high-throughput method to evaluate the impact of inhibitory compounds from lignocellulosic hydrolysates on the growth of Zymomonas mobilis. J. Biotechnol. 144, 259–267. http://dx.doi.org/10.1016/j.jbiotec.2009.08.006. Funaoka, M., 2013. Sequential transformation and utilization of natural network polymer LIGNIN. React. Funct. Polym. 73, 396–404. http://dx.doi.org/10.1016/j.

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