Cynara cardunculus a novel substrate for solid-state production of Aspergillus tubingensis cellulases and sugar hydrolysates

Biomass and Bioenergy 127 (2019) 105276

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Research paper

Cynara cardunculus a novel substrate for solid-state production of Aspergillus tubingensis cellulases and sugar hydrolysates

T

Silvia Crognalea, Federico Liuzzib, Alessandro D'Annibalea,∗, Isabella de Barib, Maurizio Petrucciolia a

Department for Innovation of Biological, Agro-food and Forestry Systems, University of Tuscia, Viterbo, 01100, Italy ENEA - Italian National Agency for New Technologies, Energy and Sustainable Economic Development, Trisaia Research Centre, S.S. 106 Jonica, 75026, Policoro, (MT), Italy

b

A R T I C LE I N FO

A B S T R A C T

Keywords: Cynara cardunculus Solid-state fermentation Cellulase production Steam explosion biomass saccharification Aspergillus tubingensis

The production of seed oils from Cynara cardunculus generates huge amounts of lignocellulosic residues which can be exploited according to a cascade approach. In this paper, residual cardoon biomass (RCB) was tested as a growth substrate for the solid-state production of cellulolytic cocktails by species known to produce glucosetolerant β-glucosidase isoenzymes. Best productions were obtained with 10-d-old Aspergillus tubingensis cultures on RCB supplemented with wheat bran (200 g kg−1) yielding β-glucosidase and endo-β-1,4-glucanase activities as high as (25 and 4) IU g−1, respectively, and 4 FPU g−1. The saccharification performance of the obtained cocktail tested on acid-catalysed steam-exploded RCB at low solid loading (25 g dm−3) was around 53% at 20 FPU g−1 cellulose. These performance were significantly enhanced by adding the xylanase-rich NS 22083 commercial formulation, reaching glucose yields higher than 80% after 72 h incubation. The use of the catalytic additive was optimized by a statistical approach, based on factorial analysis. A comparison of the performance of the A. tubingensis reinforced cocktail with the Cellic®CTec2 taken as benchmark formulation was done at the same enzyme load and performed at industrially relevant solid loadings, namely at (100 and 200) g dm−3. This comparison showed that Cellic®CTec2 led to only slightly higher glucose yields while an opposite outcome was observed for xylose yields, irrespective of the solid loading conditions. Thus, this study shows that an in-house enzyme production, based on the solid-state conversion of an industrial byproduct, able of yielding cellulolytic cocktails with substantial saccharification performance is feasible.

1. Introduction Low cost sugars from non-edible lignocellulosic biomass represent a versatile platform for the production of biofuels and chemicals. Enzymatic saccharification of biomass is an important cost item to make the process feasible. Despite the decreasing trends of the commercial blends prices in the last decade due to recombinant DNA-based production techniques both costs and company-dependent availability of cellulase preparations are among the major constraints yet. Reduction in the cost of cellulases can be achieved only by concerted efforts which address several aspects of enzyme production such as use of cheaper raw materials, selection and improvement of microbial strains and cost-effective fermentation strategies like solid state fermentation [1]. Solid-state fermentation (SSF), referred to bioconversion processes carried out in the absence of free water, mimics the natural habitat of

∗

lignocellulolytic microorganisms. This fermentation approach is less energy-demanding and equipment-oriented than submerged liquid fermentation and, thus, it is compatible with an in-house and cost-effective production of cellulolytic cocktails [2]. Although cellulases are produced by a wide array of microorganisms [3]; among them, the most exploited species for commercial glycosyl hydrolases preparations is the ascomycete Trichoderma reesei. Its cellulolytic cocktails, however, are reportedly lacking of sufficient β-glucosidase. As a consequence, T. reesei cellulolytic preparations have to be integrated by β-glucosidase from other microbial sources. Thus enzymeprospecting research continues to identify new opportunities to enhance the saccharification efficiency through the supplementation of T. reseei preparations with novel enzymatic extracts produced by other strains [4]. Cynara cardunculus L. var. Altilis (cardoon) is a perennial crop well adapted to the Mediterranean climate with a variety of commercial

Corresponding author. E-mail address: [email protected] (A. D'Annibale).

https://doi.org/10.1016/j.biombioe.2019.105276 Received 14 November 2018; Received in revised form 6 June 2019; Accepted 13 June 2019 Available online 17 June 2019 0961-9534/ © 2019 Elsevier Ltd. All rights reserved.

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(Milan, Italy), Cellic®CTec2, commercial xylanases (NS 22083) and βglucosidase (NS 22118) were from Novozymes (Bagsvćrd, Denmark).

applications; some spine-less cultivars of the aforementioned variety provide edible fleshy stems and leaf stems which are mainly consumed in France and Spain. Moreover, cardoons are used as sources of vegetable rennet for cheese production in Portugal. Due to its above-ground biomass productivity, ranging from (14–20) Mg ha−1 on a dry matter basis, and chemical composition, C. cardunculus is a very attractive lignocellulosic renewable feedstock for biorefinery purposes also due to its ability to be grown in marginal lands and to its low irrigation and fertilization requirements [5]. Extractives of this plant species include bioactive compounds with application in food and pharmaceutical industries [6,7]. Most importantly, cardoon seeds are exploited for oil production that can be used for making biodegradable plastics [8], while hairs and pappi are used for the production of paper [9,10] and bioethanol [11,12]. Several research efforts have been devoted to the optimization of cardoon pre-treatment strategies [11,13–18]. The framework of the present study is the innovative cardoon-based biorefinery located in Porto Torres (Italy) and developed by Novamont that manages around 4200 Mg of this lignocellulosic feedstock per year. The main biorefinery product is represented by seed oil from which mono-, dicarboxylic acid and esters are derived to be used as precursors of plasticizers and lubricants. The lignocellulosic residue after harvesting cardoon seed, from here onwards referred to as residual cardoon biomass (RCB), can be exploited in compliance with the principles of circular economy. Thus, the main objective of the present work was to develop a SSF process, relying on RCB as the growth substrate, able to be used for “in-house” cellulase production and, hence, either as an alternative or integrative source of enzyme supply to that represented by commercial formulations. To this aim, some fungal species were selected based on their reported ability to produce β-glucosidase isoenzymes with very low susceptibility to glucose inhibition [19,20]. The saccharification efficiencies of SSF-derived enzymatic preparations, used alone or in combination with commercial glycosyl hydrolase preparations, were then tested by using steam pretreated RCB as the substrate and its saccharification conditions were optimized with the most promising cocktail by a statistical approach relying on response surface methodology. The feasibility of the saccharification process of RCB was finally investigated at industrially relevant solid loadings and the glucose and xylose yields obtained with the in house-produced cocktail were benchmarked with a widely used commercial cellulase preparation.

2.3. Solid state fermentation (SSF) Oven-dried (60 °C) RCB (14 g) was transferred into 500-cm3 Erlenmeyer flasks and supplemented with either deionized water or Mandels and Sternburg's medium (MSM) that had the following composition (g dm−3): peptone, 1.0; (NH4)2SO4, 1.4; KH2PO4, 2.0; urea, 0.3; CaCl2, 0.3; MgSO4·• 7 H2O, 0.3 and trace elements (mg dm−3): FeSO4 • 7 H2O, 5.0; MnSO4·• H2O, 1.6; ZnSO4 •·7 H2O, 1.4; CoCl2, 2.0. The moistening solutions were then added to RCB in order to reach an initial water content of 700 cm3 kg−1. Flasks were autoclaved (121 °C for 20 min) and, after cooling, inoculated with 5 cm3 of 72 h–old precultures obtained by adding spore suspension (1 × 106 spores cm−3) to potato dextrose broth (PDB) or malt extract broth (MEB). Then, flasks were incubated under stationary conditions at a temperature depending on the strain. In particular, A. tubingensis NRRL4700 and A. caespitosus NRRL1929 were grown at 28 °C while H. insolens at 40 °C. All the experiments were carried out in triplicate and whole flasks were sacrificed after 7 and 10 d of incubation in order to perform the recovery of the enzymatic extracts and their subsequent assay. Some SSF experiments were also conducted by supplementing the growth substrate with wheat bran (200 g kg−1). 2.4. Enzyme extraction At each set incubation time, the recovery of the crude enzymatic extract from the harvested SSF cultures was performed with the aid of a hydraulic press generating a pressure of 2.0 × 107 Pa as described elsewhere [22]. To enhance the recovery, the pressed cake was rinsed in 0.1 mol dm−3 citrate buffer pH 5.0 (100 cm3), incubated under orbital shaking (2.5 Hz) for 30 min at 4 °C and subjected again to hydraulic pressing as above. The extracts were pooled together and centrifuged (8000 x g, 20 min). 2.5. Enzyme assays Filter Paper Units (FPU) and endo-β-1,4-glucanase activity were determined according to the method of Ghose [23]. At the end of the incubation period (60 and 30 min, respectively), reducing sugars released were determined by the 2, 5-dinitrosalicyclic acid (DNS) method [23]. β-glucosidase activity was determined by monitoring the formation of glucose from cellobiose, using an analysis kit (D-glucose) based on the enzymes glucose oxidase and peroxidase (RBiopharm AG, Darmstadt, Germany). Activities were expressed in International Units (IU), defined as the amount of enzyme releasing 1 μmole of glucose reducing-sugar equivalents per cm3 of the sample per min under the assay conditions.

2. Materials and methods 2.1. Microorganisms Humicola insolens CBS.147.64, A. tubingensis NRRL4700 and A. caespitosus NRRL1929, selected for this study, were maintained and sub-cultured every month on Malt Extract Agar (Difco) slants. 2.2. Materials 2.2.1. Substrate Cardoon (Cynara cardunculus L. var. Altilis) residual biomass (RCB) was supplied by Novamont (Milan, Italy) as part of the REBIOCHEM project consortium. RCB, delivered as two bales of 200–300 kg each one, was divided in stocks of roughly 30 kg, stored indoor and periodically analyzed for its composition throughout the project. The drymatter content of the raw material was around 830 g kg−1. Oven-dried RCB, analyzed for its contents in carbohydrates, lignin and ash by the NREL standard methods [21], contained (g kg−1): 350 ± 15 glucan; 140 ± 7 xylan; 22 ± 1 arabinan; 16 ± 1 galactan; 198 ± 3 insoluble lignin; 81 ± 9 ashes; 131 ± 25 soluble lignin and 2.4 ± 0.6 organic extractives.

2.6. Substrate pretreatment Before the pretreatment, RCB was mechanically ground to yield particles with dimensions ranging from (1.7–5.6) mm and pretreated under previously optimized process conditions [24]. The acid catalysed steam-explosion (ACSE) pretreatment was carried out by acid impregnation of the biomass prior to feeding the 10-dm3 batch reactor. In particular, RCB was soaked for 10 min in a H2SO4 (0.03 mol dm−3) cold-diluted solution and then squeezed. The acid adsorption on the biomass amounted to 6 g kg−1, as estimated both on the basis of the volume and concentration of the retained acid solution and by titration of the residual acid after impregnation. Then, the steam explosion pretreatment was conducted at 195 °C for 7.5 min. The pretreatment yielded two fractions, namely a water soluble fraction (WSF), mainly containing hemicellulose-related oligomers and monomers, and a water

2.2.2. Additives and commercial enzymes Polyethylene glycol 4000 and Tween 80 were from Sigma-Aldrich 2

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insoluble fraction (WIF). The saccharification trials were performed on the WIF which contained (g kg−1): cellulose, 540; xylan, 50; arabinans, 0.6; galactans, 3.8; insoluble lignin, 320.

Table 1 -D-optimal design showing the actual values and, between square brackets, the coded values of the independent variables A. tubingensis crude extract (ATCE) enzyme load (X1) and NS22083 enzyme load (X2) and observed and predicted values of percent saccharification yield. The load of the latter variable is expressed in terms of endo-β-1,4-xylanase (EX) IU g−1 cellulose.

2.7. Batch saccharification tests Enzyme crude extracts were centrifuged at 4 °C (11000 x g, 30 min) and the supernatant clarified by polyethyleneimine (PEI) addition [25]. Proteins were precipitated by adding slowly 56.1 g (NH4)2SO4 to 100 cm3 PEI-clarified extracts and centrifuging as above. The precipitate was dissolved in 0.05 mol dm−3 acetate buffer pH 4.8 (buffer A), concentrated on a stirred cell, equipped with a Diaflo PM-10 membrane and, finally, subjected to diafiltration against the same buffer. Oven-dried (60 °C) WIF was mechanically ground with a lab-scale MF109 miller (IKA, Staufen, Germany) and sieved through a 1.0-mm screen. Standard saccharification tests of WIF were performed in sterile tubes (15 cm3) with 5.0 cm3 working volume, at a solid loading of 25 g dm−3 in buffer A. All the crude extracts, derived from SSF cultures, were tested at an enzyme load of 5 FPU g−1 cellulose. At the beginning of the incubation, sodium azide was added to the reaction mixture (final concentration, 2 x 10−4 mol dm−3) to inhibit microbial contamination. The reaction was carried out in a reciprocal shaker 50 °C for 72 h. Samples (0.25 cm3) were taken from the reaction mixture at different time points for process monitoring. Amount of released glucose was determined using D-Glucose Assay Kit (GOPOD Format, Novozyme). Saccharification yield (SY%) was calculated by equation (1):

SY % =

Gtx − Gto ⋅0.9⋅100 ICC

Run number

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22

NS22083 load (X2) (EX, IU g−1 cellulose)

Saccharification yield (%) Observed

Predicted

5 [-1] 20 [+1] 5 [-1] 20 [+1] 5 [-1] 20 [+1] 15 [0.333] 10 [-0.333] 12.5 [0] 12.5 [0] 12.5 [0] 5 [-1] 20 [+1] 5 [-1] 20 [+1] 5 [-1] 20 [+1] 15 [0.333] 10 [-0.333] 12.5 [0] 12.5 [0] 12.5 [0]

1.1 [-1] 1.1 [-1] 5.5 [+1] 5.5 [+1] 4.03 [+0.333] 2.57 [-0.333] 1.1 [-1] 5.5 [+1] 3.3 [0] 3.3 [0] 3.3 [0] 1.1 [-1] 1.1 [-1] 5.5 [+1] 5.5 [+1] 4.03 [+0.333] 2.57 [-0.333] 1.1 [-1] 5.5 [+1] 3.3 [0] 3.3 [0] 3.3 [0]

42 70 60 83 56 68 66.1 75 61.1 71.3 68.8 44 70 59 84.9 57 72 66.4 72 69.3 67.6 69.2

43.9 67.0 60.3 83.4 54.8 72.5 63.9 72.6 68.8 68.8 68.8 43.9 67.0 60.3 83.4 54.8 72.5 63.9 72.6 68.8 68.8 68.8

fraction of glucan. A similar equation was used for xylose by replacing the concentrations and the corresponding coefficients (rXx = 132.11/ 150.13).

(1)

Where Gto and Gtx are glucose concentrations at the beginning and at time t, respectively, and ICC is the initial cellulose content of the solid ACSE residue. Additional experiments were also conducted by adding Tween 80 (0.5 g dm−3) and polyethylene glycol 4000 (PEG) (1 g dm−3) Alternatively, commercial formulations rich in β-glucosidase (NS22118) (5.0 IU g−1 cellulose) or xylanase (NS22083) (5.5 IU g−1 cellulose) were also added to standard saccharification tests. The saccharification performance of the most promising crude extract was compared with the commercial formulation Cellic®CTec2 at the same enzyme load, based on FPU g−1 cellulose at solid loadings closer to industrially relevant conditions. Experiments were carried-out in 100 cm3 Erlenmeyer flasks with 50 cm3 working volume, at increased biomass loading, from (50–200) g dm−3 in buffer A. The enzymatic load was 20 FPU g−1 cellulose. Incubation was performed in an orbital shaker at 3 Hz at 50 °C for 72 h. In order to improve the mixing condition at high solids loadings (i.e, 200 g dm−3), a gravimetric mixing of WIF was used enabling the biomass slurry to be lifted and dropped inside the flasks. Stirring was kept at 0.5 Hz. Samples were regularly withdrawn and analyzed. High performance liquid chromatography analysis of carbohydrates in the hydrolysates was performed by using a modified HPIC DX 300 system equipped with a working Nucleogel 300 column. The mobile phase was 0.01 mol dm−3 H2SO4 at a flow rate 0.6 cm3 min−1 and the eluate was monitored by refractive index detection using a Shodex RI101 detector. At solids loadings equal to or higher than 100 g dm−3, saccharification yields (Yg) were calculated according to the equation proposed by Roche and collaborators [26], (also known as Roche hydrolysis yield), which ensures more reliable calculation of yields since it considers the variation of the density of the liquid in the system:

Yg =

ATCE load (X1) (FPU g−1 cellulose)

2.8. Statistical optimization of the saccharification cocktail A variant of a fractional factorial design, namely a D-optimal design [27], was used to optimize the saccharification performance of the A. tubingensis crude extract (ATCE) in association with the commercial formulation which had given the best results. Thus, the impact of two quantitative variables, namely the ATCE enzyme load (X1), based on FPU g−1 cellulose and that of the NS22083 commercial formulation (X2), based on endo-β-1,4-xylanase IU g−1 cellulose, were studied by the D-optimal design. The response variable was the saccharification yield obtained after 72 h incubation. This experimental design included a set of 8 variable combinations including three center points and each combination was replicated twice thus yielding a total number of 22 runs. Table 1 shows the actual and coded values of the two independent variables in the design. Data were subjected to the analysis of variance (ANOVA) and fitted according to a second-order polynomial model as shown in Equation (2):

RVi = βo +

∑ βi⋅Xi + ∑ βii⋅Xi2+ ∑ βij⋅Xi ⋅Xj

(2)

Where RVi is the response variable, βo is the intercept, βi and βii are linear and quadratic coefficients respectively, βij is the interaction coefficient and Xi and Xj are the coded forms of the input variables. Statistical analysis of the results and generation of response surfaces were performed by the software package Modde 5.0 (Umetrics AB, Umea Sweden).

rGC Δfg

2.9. Effect of pH and temperature on activity and stability of A. tubingensis enzymatic extract

fiso (x G,0)

where rGc is the molecular weight ratio of a glucan monomer to glucose [162.16/180.18], fg is glucose mass fraction of the total slurry, fis,0 is the initial mass fraction of insoluble solids and XG,0 is initial mass

The effect of pH on A. tubingensis endo-β-1,4-glucanase (CMC-ase), FP-ase and β-glucosidase activities was assessed in the pH range from 3

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3.0 to 7.0. Sodium acetate buffer (0.05 mol dm−3) was used in the (3.0–5.0) pH range while Mc Illvaine's citrate-phosphate buffer (0.05 mol dm−3) in the (6.0–7.0) pH range. The impact of temperature on the aforementioned activities was assayed at their respective pH optima in the (30–80) °C range. Stability was determined in (4.0–7.0) pH range using the aforementioned buffer systems at 40 °C. Thermal stability was determined by incubating enzyme in buffer A at (50, 55 and 60) °C for variable incubation periods. Samples were withdrawn periodically, immediately chilled on ice and then assayed for residual activity. The inactivation constants (k) related to pH and temperature stability experiments were calculated from the linearization (4) of the first order exponential decay equation (3):

At = e−k ⋅ t Ao

(3)

A ln ⎛ o ⎞ = −k⋅t ⎝ At ⎠

(4)

⎜

cultures. In general, similar FPase, endo-β-1,4-glucanase and β-glucosidase production levels were found in H. insolens and A. caespitosus solid-state cultures irrespective of the incubation time. Conversely, in 7and 10-d-old A. tubingensins cultures, β-glucosidase activity amounted to (3.0 and 12.8) IU g−1, respectively, and was significantly higher than that observed in H. insolens and A. caespitosus cultures. The highest FPase activities were detected in 10-d-old H. insolens and A. tubingensis cultures which yielded around 2.5 IU g−1. A further improvement of A. tubingensis glycosyl hydrolase production was observed when cardoon was mixed with wheat bran (WB). β-glucosidase production in 7- and 10-d-old A. tubingensis cultures was most affected by the substrate supplementation reaching (24.8 and 24.3) IU g−1, respectively; these production levels were around 8- and 2-fold higher than those obtained in coeval non-supplemented ones. An 8-fold increase in β-glucosidase activity was also observed in H. insolens cultures at both harvests. With regard to FPase, its production levels were improved by 52% in A. tubingensis supplemented cultures (3.8 IU g−1) as compared to non-supplemented ones (2.5 IU g−1). Thus, best production results were obtained with 10-d-old A. tubingensis cultures on WB-supplemented cardoon.

⎟

where Ao and At are initial and residual activity, respectively, and t is the incubation time. The half-lives (t1/2) were calculated from Equation (5):

t1/2 =

0.694 k

3.2. Saccharification tests

(5)

The crude extracts from H. insolens, A. caespitosus and A. tubingensis solid-state cultures were compared for their saccharification abilities of the ACSE-pretreated RCB. At an enzyme load of 5.0 FPU g−1 cellulose and irrespective of the presence or the absence of additives, namely Tween 80 and PEG 4000, the A. caespitosus crude extract led to negligible saccharification yields which were invariably lower than 2.5% (Fig. 2A). A significant improvement of the saccharification yield, which was, however, lower than 10% after 72 h, was only observed when the enzymatic load was increased to 20 FPU g−1 cellulose (Fig. 2A). The H. insolens crude extracts led to even worse saccharification performance than those observed with A. caespitosus. As a matter of fact, and irrespective of both additives and enzyme loads, the percent saccharification was always lower than 5% (Fig. 2B). When saccharification tests were performed with A. tubingensis extract, at an enzyme load of 5.0 FPU g−1 cellulose, the saccharification yield reached a maximal value of 9.3% after 72 h and was not affected by the presence of additives. The increase of enzymatic load from (5.0–20) FPU g−1 cellulose enabled the attainment of a saccharification yield as high as 53% (Fig. 2C). On the basis of these results, further saccharification tests were carried out with A. tubingensis enzymatic extract.

2.10. Statistical analysis The data were analyzed by one-way or two-way analysis of variance (ANOVA test) and the means compared by a post-hoc Tukey's test using the SigmaStat Software package version 3.5 (Jandel Scientific, Germany). 3. Results 3.1. Glycosyl hydrolases production in solid-state cultures All the three selected strains grew profusely and colonized fully the cardoon biomass already within 96 h from the inoculation and this was observed for both water- and MSM-moistened cultures. However, the use of the MSM as the initial moistening agent of the solid substrate led to at least doubled glycosyl hydrolase activities as compared to cultures added with deionized water (data not shown). As a consequence, Fig. 1, reporting enzyme production data on cardoon and the same substrate supplemented with wheat bran, are referred only to MSM-moistened

Fig. 1. Filter paper units (FPU) (A), endo-β-1,4-glucanase (B) and β-glucosidase (C) activities in 7- and 10-d-old solid-state cultures of A. caespitosus (black). H. insolens (light grey) and A. tubingensis (dark grey) on Cynara cardunculus (CC) biomass as such or supplemented with wheat bran (CC + WB). All the activities are referred to unit dry mass of colonized substrate. Data are the means ± standard deviation of triplicate cultures. Same lowercase and uppercase letters denote absence of statistical significance between same production conditions with different fungal strains and between presence and absence of wheat bran within each strain and at the same time of incubation, respectively. 4

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Fig. 2. Time course of saccharification reaction of acid-catalysed steam explosion-pretreated residual cardoon biomass with crude enzymatic extract from: A. caespitosus (A), H. insolens (B); A. tubingensis (C) at an enzyme load of 5.0 FPU g−1 cellulose in the presence and in the absence of Tween 20 and Polyethylene glycol (PEG) and at 20 FPU g−1 cellulose.

high relative activity (93%) at pH 5.0. Fig. 3B shows that both endo-β-1-4-glucanase and FPase exhibited highest activity in the (55-60) °C range. The temperature optimum of βglucosidase was observed at 60 °C and its relative activity at 50 °C amounted to 80%. Noteworthy, all the glycosyl hydrolase exhibited relative activities higher than 70% at 70 °C. pH and thermal denaturation profiles of A. tubingensis FPase exhibited first order exponential decay in the time range considered; thus, from the semi-logarithmic plot of residual activity vs. incubation time, it was possible to calculate inactivation constant and half-lives (Fig. 4).

3.3. Effect of pH and temperature on activity and stability of A. tubingensis glycosyl hydrolases On the basis of the higher saccharification performance of the A. tubingensis crude extract (ATCE), the influence of temperature and pH on the activity and stability of its cellulolytic system was investigated. The pH-activity profile shows that FPase and β-glucosidase showed an identical optimum at pH 5.0 (Fig. 3A). However, the relative activity of the latter dropped at pH 6.0 exhibiting a relative activity of 42%. Although endo-β-1-4-glucanase exhibited a pH optimum at 4.0, it had a 5

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Fig. 3. Effect of pH (A) and Temperature (B) on Filter paper-ase (FPase, empty circles), endo-β-1,4-glucanase (filled circles) and β-glucosidase (filled triangles) activities of A. tubingensis. Data are the mean ± standard deviation of triplicate assays.

Fig. 4. Semi-logarithmic plot of residual activity of A. tubingensis FPase vs. incubation time at pH 4.0, 5.0, 6.0 and 7.0 (A) and at different temperatures (50, 55 and 60 °C) (B).

At pH 4.0 and 5.0, A. tubingensis FPase exhibited nearly superimposable pH stability profiles and exhibited similar inactivation constants and half-lives were (147 and 142) h, respectively (Fig. 4A). At pH 6.0 and 7.0, FPase was less stable with half-lives of (104 and 57) h, respectively. Noteworthy, the pH stability of β-glucosidase was higher at acid pHs than at neutral pH. In particular, residual activities of 48 and 67% were found at pH 4.0 and 5.0 after 96 h of incubation, while a nearly complete loss of activity was observed at pH 7.0. FPase proved to rather thermostable at (50, 55 and 60) °C with inactivation constants amounting to (7.70 • 10−3, 9.15 • 10−3 and 1.23 • 10−2) h−1, respectively, and half-lives of (90, 76 and 56) h, respectively (Fig. 4B). Regarding the thermal stability of the other components, β-glucosidase and endo-β-1-4-glucanase were highly stable at 50 and 55 °C retaining more than 90% residual activity after 96 h of incubation (data not shown).

saccharification condition (ATCE, 5 FPU g−1; T, 50 °C; solid loading, 25 g dm−3). The addition of the former formulation, mainly containing β-glucosidase, did not lead to significant differences from the reference condition (Table 2). Conversely, the supplementation with the latter formulation, which mainly contained endo-β-1,4-xylanase, gave rise to around a 4-fold increase of the saccharification (Table 2).

3.5. Statistical optimization of cardoon biomass saccharification The previous experiments showed clearly that neither the two chemical additives nor the addition of the β-glucosidase-based commercial formulation significantly improved the saccharification performance of ATCE. Conversely, since both the enzyme loads of ATCE (X1) and of the endo-β-1,4-xylanase-based commercial formulation (X2) were found to be relevant factors, their impact on saccharification was investigated by a D-optimal design, a variant of fractional factorial designs. Table 1 shows the structure of the experimental design and the response, namely the saccharification yield after 72 h, obtained for each combination of the two factors. The response varied over a range from (42–84.9)% depending on the selected factor combination. Although data were initially fitted by a second order polynomial equation, leading to a high fraction of variation explained by the model (R2adj = 0.933), both X1•X2 interaction and second order coefficient of the X2 variable were not found to be statistically significant (P equal to 0.17 and 0.43, respectively) (Table 3). Consequently, data were refitted by excluding the insignificant terms from the model and resulting in

3.4. Supplementation of A. tubingensis enzymatic extract with commercial enzymes The addition of catalytic additives, represented by some commercial glycosyl hydrolase formulations, was evaluated as a possible means of improving the saccharification performance of the A. tubingensis crude extract and, above all, to gain insights into which glycosyl hydrolase component impaired the performance of the tested crude extract. To this aim, β-glucosidase- and xylanase-rich commercial formulations (i.e, NS22118 and NS22083, respectively) were added to the A. tubingensis crude extract (ATCE) and compared with the reference 6

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Table 2 -Effect of commercial catalytic additives on percent saccharification of ACSE-pretreated C. cardunculus biomass. Saccharification was conducted at a solid loading of 25 g dm−3 at 50 °C for 72 h with A. tubingensis crude extract (5.0 FPU g−1 cellulose) alone (ATCE) and combined with different enzyme formulations. Same lowercase and uppercase letters denote absence of statistical significance between row and column means, respectively (P ≤ 0.05). ATCE (FPU g−1 cellulose)

– 5.0 – 5.0 –

5.0 5.0 5.0 – – a b

NS22118 (IU g−1 cellulose)a

NS22083 (IU g−1 cellulose)b

– – 5.5 – 5.5

Percent saccharification after 24 h

48 h

72 h

4.2 ± 0.4 aA 8.4 ± 1.5 aA 23.4 ± 2.6 aB 0.7 ± 0.2 aA 5.1 ± 0.4 aA

7.8 ± 1.6 aA 10.9 ± 2.3 aA 27.9 ± 4.3 aB 1.1 ± 0.2 abA 7.3 ± 1.1 aA

9.3 ± 3.2 aA 11.7 ± 3.4 aA 38.1 ± 1.9bB 1.9 ± 0.3bA 10.1 ± 1.8 aA

β-glucosidase activity. Endo-β-1,4-xylanase.

high values of both R2adj and Q2 (0.930 and 0.917, respectively). Moreover, the FME value, calculated from the ratio between mean squares of model error and replicate error, showed that the probability for lack of fit of the model was not statistically significant (P = 0.187) (Table 3). Fig. 5, reporting the contour diagram for saccharification yield as a function of different combinations of the two factors, shows that the minimal amount of ATCE load to reach a saccharification as high as 80%, namely 15.3 FPU g−1 cellulose, requires the presence of 5.5 IU endo-β-1,4-xylanase g−1; the same target can be obtained at the highest ATCE load by a combination with 4.6 IU endo-β-xylanase g−1.

3.6. Comparative saccharification tests at high solid loadings To pursue an advancement step towards the combined use of the ATCE and NS22083 formulation in saccharification of ACSE-pretreated CRB, further experiments were conducted at solid loadings equal to (50, 100 and 200) g dm−3 and results compared with those obtained with the Cellic®CTec2 formulation, at an equal enzyme dose of 20 FPU g−1 cellulose, and added with an equal dose of NS22083. To obtain insights into the fate of the residual xylan, xylose yield was also determined (Table 4). Cellic®CTec2-based cocktails led to higher glucose yields than those obtained with ATCE-based ones while an opposite outcome was found for xylose yields. With the sole exceptions of experiments conducted at (50 and 100) g dm−3 with the Cellic®CTec2-based cocktails, both glucose and xylose yields tended to decrease as the solid loading was increased from (50–200) g dm−3. The highest glucose yield, observed at the lowest solid loading, was around 71% by using Cellic®CTec2. The use of the ATCE, instead, led to a xylose yield of (61.8 ± 3.0)% which was higher than that obtained with Cellic®CTec2. This indicated a more effective synergy between the A. tubingensis crude extract and the added xylanase.

Fig. 5. Contour diagram for saccharification of pretreated biomass of cardoon as a function of the enzyme loads of A. tubingensis crude extract (X1) and NS22083 (X2).

4. Discussion To the best of our knowledge, this is the first study that evaluates cardoon biomass as the growth substrate for the solid-state production of cellulolytic enzymes to be applied to the saccharification of cardoon lignocellulose residues. Although the feedstock enabled a profuse growth of the fungal species under study, cellulases production required a mineral supplementation. The mineral composition of cardoon biomass is known [28] and some essential elements, such as Fe, Mg Si, and S, are known to be present in the leaves rather than in the stalks, thus possibly explaining the positive effect on cellulase production observed

Table 3 Least squares estimates of coefficients of input variables ATCE (X1) and NS22083 (X2) enzyme loads. Statistical parameters measuring fitting power (R2), predictive power (Q2) and probability of lack of fit are also shown. Model's termsa and parameters

Original model

Refined model

Constant (βo) First order coefficient of X1 (β1) First order coefficient of X2 (β2) Second order coefficient of X1 (β1•β1) Second order coefficient of X1 (β2•β2) Interaction coefficient X1X2 (β1•β2) R2 R2adj Q2 FME

67.05 ± 1.00** 11.83 ± 0.79** 6.68 ± 0.78** −5.62 ± 1.29* 1.51 ± 1.06 n.s. −0.68 ± 0.83 n.s. 0.949 0.933 (DF = 16) 0.914 1.913 (DF = 3; P = 0.177)

67.76 ± 0.91** 11.73 ± 0.79** 7.14 ± 0.72** −4.94 ± 1.20* n.i. n.i. 0.941 0.930 (DF = 18) 0.917 1.778 (DF = 5; P = 0.187)

a Estimated coefficient ± standard error and significance level: ∗ P < 0.001; ∗∗ P < 0.0001; n.s., not significant; n.i., not included; R2, coefficient of determination; R2adj, coefficient of determination adjusted for degrees of freedom (DF); FME, ratio between mean squares of model error and replicate error.

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Table 4 Percent glucose and xylose yields obtained after 72 h incubation with different enzyme loads of A. tubingensis crude extract (ATCE) and combinations with the commercial enzyme formulations Cellic®CTec2 (CTec2) and NS22083. The enzyme load of the last formulation was expressed in terms of International Units (IU) of endo-β-1,4-xylanase activity. Data are the mean ± standard deviation of triplicate experiments. Solid loading (g dm−3)

ATCE FPU g−1 cellulose

CTec2 FPU g−1 cellulose

NS22083 IU g−1 cellulose

Glucose yield (%)a

Xylose yield (%)a

50

20 0 20 0 20 0

0 20 0 20 0 20

5.5 5.5 5.5 5.5 5.5 5.5

65.4 71.2 62.3 69.8 51.3 57.6

61.8 45.5 55.4 46.8 41.5 33.8

100 200

± ± ± ± ± ±

1.3aD 1.1bE 0.7 aC 1.0bE 0.8 aA 0.5bB

± ± ± ± ± ±

3.0bD 0.3 aB 1.2bC 0.5 aB 2.3bB 2.6 aA

a Multiple pair-wise comparisons were done by the Tukey test at the confidence level of 0.95: same lowercase and uppercase letters denote absence of statistical significance between ATCE- and CTec2-based cocktails at the same solid loading and between same cocktails at different solid loadings, respectively.

(50 g kg−1). Although the residual mass fraction of xylan in the treated fiber represented only 5% of the total, it represents the recalcitrant part of hemicellulose that, due to its tight binding with the other biomass components, is less susceptible to ACSE-promoted hydrolysis. It is recognized that even small concentrations of xylooligomers can exert significant inhibitory effect on the hydrolysis of cellulose itself [38]; therefore the addition of commercial xylanases endowed with a variety of auxiliary enzyme activities, such as, for instance, β-xylosidase and arabinoxylan-arabinofuranohydrolase (AX-AFH), improves cellulose accessibility as a consequence of xylan solubilization [39]. In this respect, a recent study reported the presence in the NS22083 preparation of significant levels of AX-AFH activity [40]. Consequently, due to its relevance as a the supplement, the use of the NS22083was optimized statistically by means of a fractional factorial design which, besides leading to a robust model with high predictive power, also showed that saccharification yields higher than 80% could be obtained by moderate additions of that supplement. Albeit significant and explanatory, all these results were obtained at a low solid loading. To be industrially feasible, saccharification of a given lignocellulosic material must be conducted at a high dry matter concentration to reduce costs associated with concentration of the obtained syrups [41]. Experiments, conducted at higher solid loadings, showed that the ATCE maintained good saccharification performance; at the highest loading tested (200 g dm−3), ATCE gave slightly lower glucose yields than those of the Cellic®CTec2 and performed better than the latter in the removal of residual xylan.

after the supplementation of CRB with MSM. Another reason underlying the improved glycosyl hydrolase production derived from the use of the MSM might be due to its observed ability to counteract the acidification of the solid substrate at least in A. tubingensis and A. caespitosus cultures. Noteworthy, this mineral medium was also added as the moistening agent for the solid state production of endo-β-1,4-xylanase by another strain of A. tubingensis [29]. The supplement of wheat bran has been found to boost cellulase production in co-colture of T. reseei and A. oryzae solid state fermentation [30]. Due to its high protein content (around 170 g kg−1), wheat bran is a good source of nitrogen able of increasing the C/N ratio of the substrate; moreover, due to its high content in arabinans, it can provide arabinose units which can be assimilated as a carbon source by A. tubingensis [31]. Among the tested species, the cellulolytic cocktails derived from solid-state cultures of A. tubingensis stood out for their high levels of the β–glycosidase (BG) component. In this respect, several papers reported the attractive biochemical properties of some BG isoenzymes of this species mainly due to their high tolerance to glucose, with two isoenzymes exhibiting inhibition constants for glucose as high as 0.45 and 0.60 mol dm−3 [19]. This is very important since the product inhibition exerted by glucose on several fungal BGs leads to the accumulation of cellobiose, which, in turn, might result in the inhibition of other cellulolytic components, such as endo-β-1,4-glucanase and cellobiohydrolase, leading ultimately to low saccharification yields [32]. Thus, the use fungal species endowed with glucose-tolerant BG isoenzymes might be appropriate for producing cellulolytic cocktails. In addition to the aforementioned inhibition phenomena exerted by hydrolysis products, the non-productive binding of cellulases to lignin has been reported to significantly reduce the saccharification efficiency, thus requiring higher enzyme loads [33]. In the present study, the ACSE pretreatment of CRB led to a significant removal of the hemicelluloses associated with a relative increase in its lignin content. This was expected since the ACSE pretreatment is known to enhance the susceptibility of cellulose to enzymatic hydrolysis and to be substantially ineffective in lignin removal [34–36]. Non-ionic surfactants, such as Tweens, and PEG, known to reduce the non-productive binding of cellulolytic enzymes onto lignin [37] were unable of enhancing significantly the saccharification levels of pretreated cardoon biomass. Among the cocktails under study, the ATCE was by far the most effective leading to a saccharification yield of 54% of CRB at an enzyme load of 20 FPU g−1 cellulose. To further improve the saccharification performance of ATCE, the addition of catalytic supplements, represented by commercial enzyme formulations, was assessed. The use of the β-glucosidase-rich NS22118 formulation failed to improve the saccharification of CRB, likely due to the already high β-glucosidase levels (50 IU) in ATCE associated with an enzyme load of 5.0 FPU g−1 cellulose. Conversely, the supplementation of ATCE with the NS22083, a commercial formulation rich in endo-β-1,4-xylanase, markedly improved the saccharification of CRB. Noteworthy, this effect was relevant even though the hemicellulose solubilization obtained by the ACSE pretreatment was high and the solid residue had a low xylan content

5. Conclusions This study shows that it is possible to use the residual cardoon biomass with negligible handling operations to obtain cellulolytic cocktails through solid-state bioprocesses able to deliver significant levels of activity. The glycosyl hydrolase enzyme titers in A. tubingensis cocktails produced by ‘‘in-house’’ fermentation, albeit not relevant for stand-alone commercial enzyme production, enabled the achievement of high saccharification yields of RCB which were not far from those achieved with a widely used commercial preparation. Noteworthy, the assessment of the saccharification performance of the enzyme cocktail in question was also assessed at a consistency level (200 g dm−3) which has not been taken so far into consideration in other saccharification studies conducted with the same residue [15–18]. The establishment of an enzyme supply source either alternative or integrative to that based on commercial formulations might exert a positive impact on the currently existing cardoon-based biorefinery.

Declaration of interest None.

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[19] C. Decker, J. Visser, P. Schreier, β-glucosidase multiplicity from Aspergillus tubingensis CBS 643.92: purification and characterization of four β-glucosidases and their differentiation with respect to substrate specificity, glucose inhibition and acid tolerance, Appl. Microbiol. Biotechnol. 55 (2001) 157–163. [20] K.G. Sonia, B.S. Chadha, A.K. Badhan, H.S. Saini, M.K. Bhat, Identification of glucose tolerant acid active β-glucosidases from thermophilic and thermotolerant fungi, World J. Microbiol. Biotechnol. 24 (2008) 599–604. [21] A. Sluiter, B. Hames, R. Ruiz, C. Scarlata, J. Sluiter, D. Templeton, D. Crocker, Determination of Structural Carbohydrates and Lignin in Biomass, National Renewable Energy Laboratory, Golden, Colorado, 2010 Jul,17, Report No.TP-51042618. [22] A. D'Annibale, C. Crestini, E. Di Mattia, G. Giovanozzi Sermanni, Veratryl alcohol oxidation by manganese-dependent peroxidase from Lentinus edodes, J. Biotechnol. 48 (1996) 231–239. [23] T.K. Ghose, Measurement of cellulase activities, Pure Appl. Chem. 59 (1987) 257–268. [24] F. Liuzzi, I. De Bari, Z. Kaouther, G. Braccio, Optimization of catalyzed pretreatment and fractionation of cardoon, Proceedings of the 24th European Biomass Conference & Exibition. Amsterdam, 2016, pp. 1217–1220. [25] C. Mishra, G.F. Leatham, Recovery and fractionation of the extracellular degradative enzymes from Lentinula edodes cultures cultivated on a solid lignocellulosic substrate, J. Ferment. Bioeng. 69 (1990) 8–15. [26] C.M. Roche, C.J. Dibble, J.S. Knutsen, J.J. Stickel, M.W. Liberatore, Particle concentration and yield stress of biomass slurries during enzymatic hydrolysis at highsolids loadings, Biotechnol. Bioeng. 104 (2009) 290–300. [27] L. Eriksson, E. Johansson, N. Kettaneh-Wold, C. Wikström, S. Wold, Design of Experiments: Principles and Applications, Umetrics AB, Umea Sweden, 2008. [28] A. Monti, N. Di Virgilio, G. Venturi, Mineral composition and ash content of six major energy crops, Biomass Bioenergy 32 (2008) 216–223. [29] J.J. Pandya, A. Gupte, Production of xylanase under solid-state fermentation by Aspergillus tubingensis JP-1 and its application, Bioproc. Biosyst. Eng. 35 (2012) 769–779. [30] K. Brijwani, H.S. Oberoi, P.V. Vadlani, Production of a cellulolytic enzyme system in mixed-culture solid-state fermentation of soybean hulls supplemented with wheat bran, Process Biochem. 45 (2010) 120–128. [31] I. Nikolaev, S.F. Hansen, S. Madrid, R.P. de Vries, Disruption of the L-arabitol dehydrogenase encoding gene in Aspergillus tubingensis results in increased xylanase production, Biotechnol. J. 8 (2013) 905–911. [32] P. Andrić, A.S. Meyer, P.A. Jensen, K. Dam-Johansen, Reactor design for minimizing product inhibition during enzymatic lignocellulose hydrolysis: I. Significance and mechanism of cellobiose and glucose inhibition on cellulolytic enzymes, Biotechnol. Adv. 28 (2010) 308–324. [33] M.G. Mithra, G. Padmaja, Comparative alterations in the compositional profile of selected root and vegetable peels subjected to three pretreatments for enhanced saccharification, Int. J. Environ. Agric. Biotechnol. 2 (2017) 1732–1744. [34] W.H. Chen, B.L. Pen, C.T. Yu, W.S. Hwang, Pretreatment efficiency and structural characterization of rice straw by an integrated process of dilute-acid and steam explosion for bioethanol production, Bioresour. Technol. 102 (2011) 2916–2924. [35] G. Santi, A. D'Annibale, A. Eshel, A. Zilberstein, S. Crognale, M. Ruzzi, R. Valentini, M. Moresi, M. Petruccioli, Ethanol production from xerophilic and salt-resistant Tamarix jordanis biomass, Biomass Bioenergy 61 (2014) 73–81. [36] G. Santi, S. Crognale, A. D'Annibale, M. Petruccioli, M. Ruzzi, R. Valentini, M. Moresi, Orange peel pretreatment in a novel lab-scale direct steam-injection apparatus for ethanol production, Biomass Bioenergy 61 (2014) 146–156. [37] T. Eriksson, J. Börjesson, F. Tjerneld, Mechanism of surfactant effect in enzymatic hydrolysis of lignocellulose, Enzym. Microb. Technol. 31 (2002) 353–364. [38] Q. Qing, B. Yang, C.E. Wyman, Xylooligomers are strong inhibitors of cellulose hydrolysis by enzymes, Bioresour. Technol. 101 (2010) 9624–9630. [39] J. Hu, V. Arantes, J.N. Saddler, The enhancement of enzymatic hydrolysis of lignocellulosic substrates by the addition of accessory enzymes such as xylanase: is it an additive or synergistic effect? Biotechnol. Biofuels 4 (2011) 36. [40] C. Fehér, B. Gál, A. Fehér, Z. Barta, K. Réczey, 2015. Investigation of commercial enzyme preparations for selective release of arabinose from corn fibre, J. Chem. Technol. Biotechnol. 90 (2015) 1329–1337. [41] I. De Bari, F. Liuzzi, A. Villone, G. Braccio, Hydrolysis of concentrated suspensions of steam pretreated Arundo donax, Appl. Energy 102 (2013) 179–189.

Acknowledgements This work was carried out within the REBIOCHEM project as part of the Italian cluster on green chemistry (SPRING) that was funded by Ministero dell'Istruzione, dell'Università e della Ricerca (MIUR) (grant n. CTN01-00063-49393). References [1] R.K. Sukumaran, R.R. Singhania, G.M. Pandey, A Cellulase production using biomass feed stock and its application in lignocellulose saccharification for bio-ethanol production, Renew. Energy 34 (2009) 421–424. [2] C.R. Soccol, E.S.F. da Costa, L.A.J. Letti, S.G. Karp, A.L. Woiciechowski, L.P. de Souza Vandenberghe, Recent developments and innovations in solid state fermentation, Biotechnol. Res. Inn. 1 (2017) 52–71. [3] S.S. Behera, R.C. Ray, Solid state fermentation for production of microbial cellulases: recent advances and improvement strategies, Int. J. Biol. Macromol. 86 (2016) 656–669. [4] C.S. Farinas, Developments in solid state fermentation for the production of biomass-degrading enzymes for the bioenergy sector, Renew. Sustain. Energy Rev. 52 (2015) 179–188. [5] L. Sulas, A. Ventura, L. Murgia, Phytomass production from Silybum marianum for bioenergy, Options Méditerranéennes, Series A 79 (2008) 487–490. [6] P.A.B. Ramos, Â.R. Guerra, O. Guerreiro, Antiproliferative effects of Cynara cardunculus L. var. altilis (DC) lipophilic extracts, Int. J. Mol. Sci. 18 (2017) 63. [7] Z. Velez, M. Campinho, A. Guerra, L. Garcia, P. Ramos, O. Guerreiro, L. Felicio, F. Schmitt, M. Duarte, Biological Characterization of Cynara cardunculs L.methanolic extract: antioxidant, anti-proliferative, anti-migratory, and anti-angiogenic activities, Agriculture 2 (2012) 472–492. [8] A. Bouriazos, E. Ikonomakou, G. Papadogianakis, Aqueous-phase catalytic hydrogenation of methyl esters of Cynara cardunculus alternative low-cost non-edible oil: a useful concept to resolve the food, fuel and environment issue of sustainable biodiesel, Ind. Crops Prod. 52 (2014) 205–210. [9] J. Gominho, A. Lourenço, M. Curt, J. Fernández, H. Pereira, Characterization of hairs and pappi from Cynara cardunculus capitula and their suitability for paper production, Ind. Crops Prod. 29 (2009) 116–125. [10] J. Gominho, M.D. Curt, A. Lourenço, J. Fernández, H. Pereira, Cynara cardunculus L. as a biomass and multi-purpose crop: a review of 30 years of research, Biomass Bioenergy 109 (2018) 257–275. [11] M.C. Fernandes, M.D. Ferro, A.F.C. Paulino, A.S.J. Menedes, J. Gravitis, D.V. Evtuguin, A.M.R. B Xavier, Enzymatic saccharification and bioethanol production from Cynara cardunculus pretreated by steam explosion, Bioresour. Technol. 186 (2015) 309–315. [12] G. Cavalaglio Cotana, M. Gelosia, V. Coccia, A. Petrozzi, D. Ingles, E. Pompili, A comparison between SHF and SSSF processes from cardoon for ethanol production, Ind. Crops Prod. 69 (2015) 424–432. [13] I. Ballesteros, M. Ballesteros, P. Manzanares, M.J. Negro, J.M. Oliva, F. Sáez, Dilute sulfuric acid pretreatment of cardoon for ethanol production, Biochem. Eng. J. 42 (2008) 84–91. [14] A.A. Shatalov, H. Pereira, Biorefinery of energy crop cardoon (Cynara cardunculus L.)-hydrolytic xylose production as entry point to complex fractionation scheme, J. Chem. Eng. Process Technol. 2 (2011) 8. [15] J. Martinez, M.J. Negro, F. Saez, J. Manero, R. Saez, C. Martin, Effect of acid steam explosion on enzymatic hydrolysis of O. nervosum and C. cardunculus, Appl. Biochem. Biotechnol. 24 (1990) 127–134. [16] M.C. Fernandes, M.D. Ferro, A.F. Paulino, H.T. Chaves, D.V. Evtuguin, A.M. Xavier, Comparative study on hydrolysis and bioethanol production from cardoon and rockrose pretreated by dilute acid hydrolysis, Ind. Crops Prod. 111 (2018) 633–641. [17] P. Vergara, M. Ladero, F. García-Ochoa, J.C. Villar, Valorization of Cynara cardunculus crops by ethanol-water treatment: optimization of operating conditions, Ind. Crops Prod. 124 (2018) 856–862. [18] P. Vergara, M. Ladero, F. García-Ochoa, J.C. Villar, Pre-treatment of corn stover, Cynara cardunculus L. stems and wheat straw by ethanol-water and diluted sulfuric acid: comparison under different energy input conditions, Bioresour. Technol. 270 (2018) 449–456.

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