Sugarcane bagasse fly ash as a no-cost adsorbent for removal of

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Sugarcane bagasse fly ash as a no-cost adsorbent for removal of phenolic inhibitors and improvement of biomass saccharification Juliana V Freitas, and Cristiane Farinas ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/ acssuschemeng.7b03214 • Publication Date (Web): 26 Oct 2017 Downloaded from http://pubs.acs.org on October 27, 2017

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ACS Sustainable Chemistry & Engineering

Sugarcane bagasse fly ash as a no-cost adsorbent for removal of phenolic inhibitors and improvement of biomass saccharification

Juliana V. Freitas1,2 and Cristiane S. Farinas1,2*

1

Embrapa Instrumentation, Rua XV de Novembro 1452, 13561-206, São Carlos – SP, Brazil

2

Graduate Program of Chemical Engineering, Federal University of São Carlos, 13565-905, São Carlos – SP, Brazil

*Corresponding author: Cristiane S. Farinas [email protected] Tel.: +55 16 2107 2908 Fax: +55 16 2107 2902

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Abstract The phenolic compounds generated during the pretreatment of lignocellulosic biomass have inhibitory effects on the enzymatic hydrolysis and fermentation steps in biorefineries employing the biochemical platform. This work proposes the use of sugarcane bagasse fly ash as a no-cost adsorbent for removal of the phenolics generated by the liquid hot water (LHW) pretreatment of sugarcane bagasse. Physical-chemical characterization revealed that the sugarcane bagasse fly ash was mesoporous and possessed a surface charge that promoted the adsorption of monomeric (vanillin) as well as oligomeric (tannic acid) phenolic compounds, under different conditions of pH and temperature. Adsorption isotherms for the fly ash revealed similar maximum capacities for both types of phenolic compound. The potential use of the fly ash as an adsorbent for biomass pretreatment inhibitors was demonstrated by the removal of 80% of the phenolics from the LHW liquor, which resulted in a remarkable 52% increase in the glucose released in the enzymatic hydrolysis of sugarcane bagasse. The findings demonstrated that the use of sugarcane bagasse fly ash to remove phenolic compounds could effectively increase the efficiency of enzymatic hydrolysis of lignocellulosic biomass, with performance similar to that of commercial activated carbon. In addition, there is no cost for the acquisition of the fly ash, making the process highly economically attractive for implementation in future large-scale biorefineries.

Keywords: Adsorption; sugarcane bagasse; fly ash; phenolic compounds, enzymatic hydrolysis; biorefinery.

Introduction In a biorefinery, the conversion of lignocellulosic biomass into simple sugars by means of the biochemical pathway involves a pretreatment step in order to facilitate the 2 ACS Paragon Plus Environment

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enzymatic hydrolysis of cellulose. However, the pretreatment process also results in the release of varying amounts of inhibitory compounds such as phenolics, soluble sugars, furan aldehydes, and weak acids 1, 2. Lignin-derived phenolic compounds are especially harmful to the subsequent hydrolysis and fermentation steps, because they can inhibit and/or deactivate enzymes 3-9, as well as impair the ethanolic fermentation by yeasts and bacteria 10-12. Economic constraints on the implementation of future biorefineries make it necessary to identify process options able to reduce the levels of phenolic inhibitors generated during the pretreatment step, since the enzymatic cocktails contribute significantly to the overall cost of the conversion process 13, 14. Strategies that have been reported for removal of the phenolic compounds include the use of chemical additives

15-17

, enzymatic treatment

18

, microbial

treatment 19-21, liquid-liquid extraction 22, and liquid-solid extraction 5, 23, 24. In the liquid-solid extraction process, activated carbon has been used as an efficient adsorbent for removing phenolics from liquid hot water pretreated maple 5, steam pretreated mixed hardwood 24, and steam pretreated poplar and lodgepole pine 23, among others. Although commercial activated carbon adsorbents provide efficient removal, their acquisition costs could make the processes economically unviable. Alternative lower cost adsorbents prepared from waste materials have been reported for the adsorption of phenolic compounds from different sources

25-31

. However, in these

cases, treatments (thermal or chemical) were required for preparation of the adsorbents, implying additional energy and/or reagent costs. In contrast, fly ash is a residual waste material generated in sugarcane mill boilers following incomplete carbonization of the sugarcane bagasse used as fuel 32. The fly ash has a high content of unburned carbon 33, and it has been previously used as an effective adsorbent for the removal of phenol from aqueous solution

34-38

. Therefore, these suggest that it might be feasible to use fly ash as a cost-



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effective adsorbent for removal of the inhibitors generated during the pretreatment of lignocellulosic material. Here, we evaluate the use of sugarcane bagasse fly ash, without any additional treatment, as a no-cost adsorbent for removal of the phenolic compounds generated during the liquid hot water (LHW) pretreatment of sugarcane bagasse. The fly ash was characterized by N2 adsorption-desorption (BET), Fourier transform infrared spectroscopy (FTIR), X-ray diffraction (XRD), and zeta potential measurements. A preliminary set of adsorption experiments was carried out using vanillin and tannic acid (monomeric and oligomeric phenolic compounds, respectively) as models in order to evaluate the effects of pH and temperature, with determination of the equilibrium times and construction of adsorption isotherms. The positive effect of phenolics removal from LHW pretreatment liquor was then demonstrated during enzymatic hydrolysis of sugarcane bagasse.

Materials and Methods Materials Sugarcane bagasse fly ash was kindly provided by the Biorigin/Zilor sugarcane mill (Lençóis Paulista, Brazil). The dried material was sieved to 1 mm to remove impurities, milled, and sieved to 300 µm. Particles smaller than 300 µm were used for adsorption. Activated carbon, vanillin, and tannic acid were used as received from Sigma-Aldrich (Brazil). Cellic® Ctec2 (231 FPU/mL) was provided by Novozymes America Latina (Araucaria, Brazil). Sugarcane bagasse, obtained from the Nardini sugarcane mill (Vista Alegre do Alto, Brazil), was pretreated using the liquid hot water (LHW) method in a 5.5-L PARR reactor (Model 4580, Parr Instruments, USA).


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The LHW pretreatment was carried out using a 10% solids loading and a temperature of 195 ºC for 10 min39. After pretreatment, the liquor was separated from the slurry by vacuum filtration using a Whatman® N°1 filter paper. The vacuum filtrate was stored frozen until use in the adsorption experiments. The solid fraction of the pretreated bagasse was dried at 50 °C up to a final moisture of around 10%. The hydrothermally-pretreated bagasse was subsequently milled and sieved, and the particle size selected was 1.0≤dp≤2.0 mm for use in the hydrolysis experiments, without any washing procedure.

Characterization of the sugarcane bagasse fly ash N2 adsorption-desorption isotherms were obtained at 77 K and relative pressures up to 0.01, using an ASAP 2020 instrument (Micrometrics, USA). The specific surface area (SBET) of the adsorbent was calculated from the adsorption data using the Brünauer-Emmett-Teller (BET) method. The micropore area and volume (Smic and Vmic) were obtained by the t-plot method. The mesopore volume (Vext) was calculated as the difference between the total pore volume (VT) and the micropore volume (Vmic), obtained at P/P0 = 0.95. The pore diameter distributions of the adsorbent were obtained using density functional theory (DFT) calculations applied to the nitrogen adsorption isotherms, performed with the ASAP 2020 software. X-ray diffraction (XRD) patterns were obtained using a Shimadzu XRD-6000 instrument (Japan) operated with Cu Kα radiation (λ = 0.1546 nm) in the 2θ range from 5 to 80°, with voltage and current of 30 kV and 30 mA, respectively. Fourier transform infrared spectroscopy (FTIR) was carried out with the adsorbent mixed with KBr (sample/KBr ratio of 1:100) and prepared in the form of a disk by compressing the powder using a hydraulic press. The FTIR spectra were recorded from 4000 to 400 cm-1 using a VERTEX 70 spectrometer (Bruker, Germany). 5 ACS Paragon Plus Environment


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The point of zero charge (PHpzc) was determined by a titration method, using a Zetasizer Nano ZS system (Malvern Instruments, UK). A 10 mg portion of the adsorbent was added to 10 mL of aqueous solution. The solution pH was adjusted in the range from pH 1 to pH 10 using 1 M HCl and 1 M NaOH. All the characterizations were also carried out for a commercial activated carbon adsorbent, for comparison with the sugarcane bagasse fly ash.

Effect of adsorption temperature and pH of the synthetic solutions Batch adsorption experiments were performed in an incubator (Ethik Technology, Brazil) with rotating mixing at 30 inversions per min. Preliminary tests were carried out using 2, 5, and 8% (w/v) of adsorbent in order to select a suitable adsorbent loading. In the subsequent experiments, the effects of adsorption temperature (from 30 to 50 °C) and pH (from 2 to 6) were evaluated using 2% (w/v) of adsorbent. A temperature of 30 °C was used for assays in which the pH was varied, and pH 5 was used for assays in which the temperature was varied. Both of these procedures were carried out for synthetic solutions containing 3 g/L of vanillin or tannic acid prepared in 50 mM citrate buffer. The samples were placed in 5 mL tubes and were mixed for 24 h, followed by centrifugation and removal of the supernatant for quantification of the concentrations of phenolics. The tests were performed in triplicate and the results were presented as averages and standard deviations.

Adsorption isotherms and equilibrium times for the synthetic solutions The adsorption equilibrium time was obtained by quantification of the non-adsorbed solutes in the supernatants after incubation of the synthetic solutions (vanillin and tannic acid at initial concentrations of 3 g/L) with 5% (w/v) of adsorbent in 50 mM citrate buffer (pH 5), in 5 mL tubes, at 30 rpm and 30 °C, until reaching equilibrium. After each incubation period, the adsorbent was removed from the solution by centrifugation at 4 °C and 12000 rpm, and 6 ACS Paragon Plus Environment

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the residual adsorbate concentration was determined. In order to obtain the adsorption isotherms, solutions of vanillin and tannic acid at concentrations of up to 9.5 and 6.8 g/L, respectively, in 50 mM citrate buffer (pH 5), were placed in contact with 1% (w/v) of adsorbent in 5 mL flasks, at 30 rpm and 30 °C for 24 h. A 1% (w/v) concentration of adsorbent was used, due to the low solubility of vanillin. The equilibrium time and adsorption isotherm procedures were also carried out using the commercial activated carbon (CAC), for the purposes of comparison. The tests were performed in duplicate. At equilibrium, the amount of adsorbed compound, representing the adsorption capacity (Q), could be obtained using Eq. 1: Q = V x

Co - Ce

(1)

m

where Co and Ce (g/L) are the initial and equilibrium adsorbate concentrations, respectively (Ce is equivalent to Ct when it is used to obtain Q at time t), V is the solution volume (L), and m is the mass of adsorbent (g). The experimental data were fitted using the empirical isotherm models of Freundlich, Langmuir, and Redlich-Peterson. The constants of the models were determined by nonlinear regression of the experimental data, performed using OriginPro 8.0 software. The Langmuir model is represented by: Q =

Qm x K x Ce

(2)

1 + K x Ce

where K is the Langmuir constant (L/g) and Qm is the amount of adsorption corresponding to complete coverage (g/g). The Freundlich model is given by: 1

(3)

Q = K x Ce n

where K (L/g) and n are the Freundlich constants. The Redlich-Peterson isotherm model is a fit between the Langmuir and Freundlich models and can be applied to homogeneous or heterogeneous systems 40. It is described by: 7 ACS Paragon Plus Environment


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K x Ce

Q = 1 + a x Ce b

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(4)

where K (L/g), a (L/g), and b are the Redlich-Peterson constants. The value of b lies between zero and unity: when b is close to unity, the isotherm reduces to the form of the Langmuir isotherm; when b is close to zero, the isotherm reduces to the Freundlich form.

Removal of phenolics from LHW pretreatment liquor followed by enzymatic hydrolysis Removal of phenolics generated from LHW pretreatment of sugarcane bagasse liquor was carried out using the fly ash as an adsorbent, prior to the enzymatic hydrolysis step. The adsorption was carried out for 24 h at 30 °C, pH 3.3±0.3 (original pH of the liquor), and 30 rpm, using 5% (w/v) sugarcane bagasse fly ash. The enzymatic hydrolysis was carried out using the treated liquor (adjusted to pH 5 with 1 M NaOH solution) and 10% (w/v) solids loading of LHW pretreated sugarcane bagasse, with a Cellic® Ctec 2 enzyme concentration of 3 FPU/g solids. These tests were carried out for 24 h in 5 mL flasks with a liquid volume of 2 mL, at 50 °C and 30 rpm. The experiments were performed in triplicate and the results were presented as averages and standard deviations.

Filter Paper Assay The Filter Paper Assay (FPA) was performed as described by Ghose 41, with some modifications in order to reduce the volume of reaction. Briefly, the assay was performed in 96 well microplates by adding a 20 µL aliquot of diluted enzyme into wells containing a 7mm diameter Whatman® N°1 filter paper disk and 80 µL of 50 mM sodium citrate buffer, pH 4.8. After 60 min of incubation at 50 °C under 300 rpm in a ThermoMixer (Eppendorf, Germany), 100 µL of DNS reagent was added and the mixture was incubated at 100 °C for 10 min. The reaction was stopped by incubation at ice cold temperature for another 10 min.


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Absorbance was then measured at 540 nm in a BioTek Epoch 2 Microplate Spectrophotometer (Fisher Scientific, Canada). The assays were performed in triplicate and the results were presented as averages and standard deviations.

Analytical methods The concentrations of the phenolic compounds as single constituents were determined by spectrophotometry using the Beer-Lambert relation (at wavelengths of 232 and 279 nm for vanillin and tannic acid, respectively), while the Folin-Ciocalteu reaction was used for the LHW pretreatment liquor sample. The total phenolic compound concentrations were expressed as gallic acid equivalents (g GAE/L)

42

. Glucose concentrations were determined

by an enzymatic reaction (GOD-POD), using a Liquiform kit (Labtest, Brazil). Reducing sugars were measured by the DNS method 43, using glucose as standard.

Results and Discussion Characterization of the sugarcane bagasse fly ash Table 1 shows the data for the pore structure properties of the sugarcane bagasse fly ash and the commercial activated carbon used for comparison purposes. The sugarcane bagasse fly ash surface area was 431 m2/g and the values of the micropore properties (area and volume) were higher than the external properties. The micropore values for the fly ash were lower than for the activated carbon, while the values of the external properties of the adsorbents were similar. The Sext/SBET and Vext/VT ratios showed that the fly ash was more mesoporous than the activated carbon. Figure 1 compares the differential pore volumes of the fly ash and the activated carbon. According to IUPAC

44

, pores are classified as micropores

(up to 2 nm), mesopores (between 2 and 50 nm), and macropores (larger than 50 nm). The 9 ACS Paragon Plus Environment


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predominant pore dimensions for the fly ash were at around 1.7 and 3 nm (Figure 1), while for the activated carbon they were at around 2 nm and in a larger size range from about 9 to 16 nm. The presence of mesopores is important for accelerating the diffusion of molecules into micropores and increasing the equilibrium coverage of the micropore surface

45

. The

pore structure properties of the fly ash were indicative of similar availability as activated carbon for the adsorption of large molecules (such as tannic acid). As expected, the pore development of the fly ash was less intense, compared to activated carbon (Table 1). This was probably because commercial activated carbon is usually prepared in two steps, with carbonization of the material under an inert atmosphere, followed by activation of the charred product. In contrast, the fly ash was produced in an industrial boiler, which only involved carbonization of the bagasse. Properties similar to those of the sugarcane bagasse fly ash used here were reported for activated carbon prepared by heating sugarcane bagasse at 750 °C for 2 h (with 80.6% burn-off), resulting in a material with surface area of 446 m2/g and total pore volume of 0.28 cm3/g 31. Figure 2 shows the FTIR, XRD, and zeta potential results for the sugarcane bagasse fly ash and the activated carbon. FTIR was used to identify the surface functional groups (Figure 2a). According to László

46

, the chemical surfaces of adsorbents are determined by

heteroatoms (oxygen, nitrogen, hydrogen, and phosphorus) whose concentrations and chemical forms depend on the origin of the precursor material and the preparation conditions. As expected, the spectrum of the fly ash was characteristic of carbonaceous materials. A band at 471 cm-1 reflected the presence of aliphatic C-H groups 47. A band at 1095 cm-1 could be assigned to stretching vibrations of carboxylate C-O and bending modes of O-H in phenolic structures

47, 48

, while a band at 1389 cm-1 was characteristic of O-H vibrations (phenolic

structures). A band at 1573 cm-1 was indicative of vibrations of C=O moieties in i) carboxylics, esters, and lactones, ii) quinones and/or ion-radical structures, and iii)


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conjugated systems such as diketones, keto-esters, and keto-enols

47

. These bands were

similar to the activated carbon bands, while bands at 3432 and 2905 cm-1 were also obtained for the activated carbon. These bands could be assigned to hydroxylic O-H groups and stretching of aliphatic -CH2- groups, respectively 47. The carboxyl and lactone acid groups (carbon-oxygen structures) are responsible for the acidity of the carbon surface 49. The adsorbents presented intense bands assigned to acid groups (at 1095 cm-1 for both adsorbents and at 3432 cm-1 for the activated carbon). It has been reported that the presence of basic groups on the surfaces of adsorbents is desirable for the adsorption of phenolic compounds

50-54

. Bands at 1573 cm-1 in the spectra of both

adsorbents were indicative of the presence of basic groups such as diketones and quinones, although the intensities were low, compared to the bands of acid groups. Electrostatic attraction considerations therefore suggest that phenolic compounds should be more easily adsorbed onto fly ash than onto activated carbon due to the presence of an additional band assigned to acid groups in the latter. X-ray diffractograms were used for comparative analysis of the structures of the sugarcane bagasse fly ash and activated carbon (Figure 2b). Broad peaks between 5° and 40° (fly ash) and between 15° and 30° (activated carbon) reflected the presence of disordered microcrystalline carbonized material, coexisting with small amounts of amorphous phase silica and/or alumina. Peaks at 26° revealed the presence of quartz on the surfaces of the adsorbents

55

. Similar amorphous structures have been observed in the XRD patterns of

activated carbon from sugarcane bagasse

30, 56, 57

and pyrolytic tire char

29

. These

characteristics were expected, since the formation of crystalline structures in activated carbons is unusual 56, 57. The zeta potential results for the sugarcane bagasse fly ash and the activated carbon (Figure 2c) revealed isoelectric points of 2.7 and 2.0, respectively. At pH lower than the



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pHpcz, the adsorbent is positively charged and adsorption of anions is favored, while at pH higher than the pHpcz, the adsorbent is negatively charged, favoring the adsorption of cations.

Effects of adsorption pH and temperature The effects of pH (from 2 to 6) and temperature (from 30 to 50 °C) on the adsorption of vanillin and tannic acid onto the sugarcane bagasse fly ash are shown in Figure 3. The choice of these phenolic compounds (vanillin as a monomeric phenolic, and tannic acid as an oligomeric phenolic) at initial concentrations of 3 g/L was based on previous work concerning enzymatic inhibition and deactivation

3, 4, 6

. The adsorption of vanillin was only

slightly affected by the pH, while tannic acid showed an abrupt reduction in adsorption with increasing pH, from 95% (pH 2) to 16% removal (pH 6) (Figure 3a). The adsorption temperature only slightly affected the removal of vanillin, while the adsorption of tannic acid decreased with increasing temperature (Figure 3b). According to Xiao et al.

58

the proportion of vanillin present in the molecular form

does not change up to pH 5.31 (indicating an absence of ionization of the compound), which could explain the behavior observed for adsorption of vanillin onto the fly ash. In contrast, tannic acid is a weak organic acid and its ionization is strongly dependent on pH. It occurs in a molecular form at pH ≤4 (with charge close to zero), with initiation of ionization of the molecules above this value. Tannin is almost completely dissociated at pH 7 59. As shown by the zeta potentials (Figure 2c), the charge on the fly ash tended to become negative as the pH increased. Higher concentrations of OH- ions in the solution resulted in competition with the fly ash surface charge, hence decreasing the adsorption. Therefore, the adsorption of tannic acid onto the fly ash was negatively affected by dissociation of the molecules. Increasing the adsorption temperature did not reduce the adsorption of vanillin, possibly due to its relatively low solubility (2.5 g/L) at room temperature. Tannic acid is 12 ACS Paragon Plus Environment

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highly soluble (250 g/L), so its affinity for the aqueous solution favors desorption processes. With increasing temperature, the solubility of the adsorbate tends to increase. The affinity between the solute and the adsorbent is usually the main factor that controls the adsorption. However, the strength of the interaction between the solute and the solvent is an important contributing factor, and can vary widely 60. According to Moreno-Castilla and Rivera-Utrilla 61

, the adsorption capacity of activated carbon is dependent on the degree of activation, the

hydrophobicity of the phenolic compounds, and their solubility. The same adsorption characteristics are expected for the fly ash. Similar adsorption behavior at different pH values has been reported for the adsorption of vanillin onto resin modified by acetamine and hydroxyl groups, at pH ranging from 0 to 14

58

, and for adsorption of tannic acid onto

coconut activated carbon, at pH ranging from 2 to 10 62. The choice of pH and temperature for the following adsorption experiments was made based on these results and also taking into consideration that the main idea of this work was to evaluate the feasibility of carrying out the adsorption at the original conditions in which the liquor would be generated at the LHW pretreatment process (pH around 3.0) and/or at the conditons in which the cellulolytic enzymes are mostly active (pH around 5.0 and temperature of 50 °C). However, considering that the higher temperature values were not favored for adsorption of tannic acid, the preferred choice of temperature would be 30 °C, justified also as a condition closer to room temperature, thus contributing to energy savings.

Adsorption isotherms for vanillin and tannic acid The equilibrium adsorptions of vanillin and tannic acid onto the sugarcane bagasse fly ash and the commercial activated carbon are illustrated in Figure 4. Table 2 provides the parameters of the three isotherm models (Langmuir, Freundlich, and Redlich-Peterson), and



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correlation coefficients (R2). The adsorption isotherms describe the relation, at equilibrium, between the solid phase concentration (Q) and the liquid phase concentration (Ce). All the isotherms presented concave forms (Figure 4), indicative of favorable adsorption, with the capacity to adsorb high amounts of solute at low concentrations of adsorbent. The fitted models showed different correlations for the conditions evaluated (Table 2). The Freundlich model provided a good description of the adsorption of vanillin by the fly ash (Figure 4a), suggesting that the stronger binding sites were occupied first. The Redlich-Peterson model also satisfactorily described these experimental data. All the models provided good fits to the experimental data for adsorption of tannic acid by the fly ash (Figure 4b), with R2 values between 0.98 and 0.99. The correlation coefficients obtained for the isotherm models applied to the adsorption of vanillin onto activated carbon were lower than 0.90, although the Freundlich and Redlich-Peterson models showed good fits to the experimental data (Figure 4c). The Langmuir isotherm model provided the best fit to the experimental data for adsorption of tannic acid onto activated carbon (Figure 4d), indicative of significant monolayer coverage of the adsorbate on the outer surface of the adsorbent. The linearity of the isotherm decreases as the n value of the Freundlich model increases. Here, the linearity of the isotherms decreased in the order: tannic acid onto fly ash > vanillin onto fly ash > tannic acid onto activated carbon ≈ vanillin onto activated carbon. In the case of the Langmuir model, the affinity of the isotherms was the inverse, as can be seen from the increasing K values of the model (Table 2). Although the Langmuir model did not provide a good fit for the adsorption of vanillin onto activated carbon, the K value was consistent, since vanillin showed a high affinity for this adsorbent (Figure 4c). All the adsorption data could be described by the Redlich-Peterson isotherm model. This model reduces to the Langmuir isotherm when the value of the b parameter is close to unity. The highest b value (0.91±0.04) was obtained for the adsorption of tannic acid onto


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activated carbon. The adsorption of tannic acid onto fly ash also showed the potential for reduction to the Langmuir isotherm (with a b value of 0.82±0.21). The maximum adsorption capacities of the fly ash for vanillin and tannic acid were similar (Figures 4a and 4b). These results suggested that the pore structure properties of the fly ash (Table 2) favored the adsorption of both oligomers and monomers. Interestingly, the maximum adsorption capacity of the fly ash for tannic acid (Figure 4b) was higher than the value found for activated carbon (Figure 4d). As expected, the highest maximum adsorption capacity (0.35 g/g) was obtained for the adsorption of vanillin onto activated carbon (Figure 4c). This could be explained by the high surface area and micropore area of this adsorbent, since vanillin is a monomeric molecule with a size of 0.72 x 0.52 nm 58. The external properties of the fly ash were very similar to those of the activated carbon, while the values of the micropore properties of the fly ash were lower, compared to the activated carbon (Table 1). Therefore, it was not expected that the maximum adsorption capacity of the fly ash for tannic acid would be higher than that of the activated carbon. There are two possible explanations for this finding. The first is that the size of the oligomeric tannic acid molecule (1.6 nm)

45

could have resulted in an exclusion effect, given that the

dimensions of most of the pores of the activated carbon were close to the micropore range (Figure 1), which could have limited the area available for adsorption of this molecule. A second possible explanation, either alone or in conjunction with the pore size effect, is that the smaller quantity of acid functional groups in the fly ash assisted the adsorption of tannic acid. It is possible that the acid functional groups of the activated carbon could have hindered tannic acid adsorption, since the acid groups produce “clusters” that can block the transport of molecules towards the micropores

54

. This effect was not noticed in the case of vanillin,

which could be explained by the high surface area of activated carbon available for the adsorption of small molecules. The size exclusion effect has been observed previously for the



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adsorption of tannic acid onto activated carbon primarily constituted of micropores with sizes smaller than 1.5 nm 63.

Time profiles of vanillin and tannic acid adsorption onto sugarcane bagasse fly ash Figure 5 shows the time profiles for adsorption of vanillin and tannic acid onto the fly ash and activated carbon. The assays were carried out at 30 °C, pH 5, 30 rpm, using 5% (w/v) of adsorbent and 3 g/L initial solute concentration. The adsorption of vanillin onto the fly ash was only slightly slower than onto the activated carbon. It took 5 min to remove 80% of the vanillin onto the fly ash, with equilibrium reached after 120 min, corresponding to an adsorption capacity of 52 mg/g (92% removal). The adsorption of vanillin onto activated carbon was practically instantaneous, with almost complete adsorption (97%) in only 5 min, corresponding to an adsorption capacity of 57 mg/g. For tannic acid, adsorption using the fly ash reached equilibrium within 60 min, with an adsorption capacity of 41 mg/g (65% removal), while adsorption onto the activated carbon reached equilibrium after 120 min, with the same adsorption capacity as the fly ash. Interestingly, the adsorption of tannic acid onto the fly ash was faster than onto activated carbon, possibly due to the greater availability of mesopores that could facilitate diffusion of the molecules within the pores of the adsorbent. The high rate of adsorption of vanillin onto the activated carbon could be explained by the high specific surface area of this adsorbent, which provided greater probability of contact with the small vanillin molecules, with no influence of pore size. Similar behavior has been reported for the adsorption of phenol and tannic acid on mesoporous carbon (CMK-3), activated CMK-3, and activated carbon 63.


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Effect of removal of phenolics from LHW pretreatment liquor on the enzymatic hydrolysis of sugarcane bagasse In order to minimize the negative effects of the phenolic compounds in the enzymatic hydrolysis, the sugarcane bagasse LHW liquor was first treated using the fly ash as an adsorbent. A preliminary set of adsorption experiments using the LHW liquor was carried out at pH 3 and 5, under both 30 and 50 °C, since these conditions would be preferred choices to apply industrially, as previously mentioned. No significant differences were observed for phenolics removal under this set of conditions and therefore the rest of adsorption experiments were performed using the original pH of the LHW liquor (pH 3) at 30 °C. After the adsorption, the liquor was separated from the fly ash and the pH was adjusted to pH 5 prior to initiation of the enzymatic hydrolysis. Adsorption was also performed using the commercial activated carbon, followed by enzymatic hydrolysis using the treated liquor, for comparison purposes. The results obtained for the adsorption of phenolic compounds and the enzymatic hydrolysis using the treated liquors are shown in Figure 6. The initial total phenolic compounds concentration in the liquor was 2.08±0.06 GAE g/L. Removal rates of around 80% and 90% were obtained using the sugarcane bagasse fly ash and the commercial activated carbon, respectively. Enzymatic hydrolysis of the LHW pretreated sugarcane bagasse using a buffer solution instead was carried out, resulting in a glucose concentration of 10.08±0.91 g/L after 24 h of hydrolysis. Hence, the presence of the enzyme inhibitors in the liquor resulted in a 57% lower glucose concentration. When the phenolics concentration in the liquor was reduced by adsorption onto the sugarcane bagasse fly ash, the enzymatic hydrolysis started with a lower phenolics concentration of 0.42 GAE g/L, resulting in a substantial increase of 52% in the glucose released. This improvement was comparable to the 65% increase achieved using the commercial activated carbon. 17 ACS Paragon Plus Environment


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In order to confirm that the major positive effect observed in the enzymatic hydrolysis was due to the removal of phenolic compounds by adsorption onto the fly ash, a control experiment was carried out using vanillin and tannic acid solutions, before and after treatment with fly ash, in the enzymatic hydrolysis of a cellulose source (filter paper), performed as described for the FPA protocol (Figure 7). Consistent to previous studies

3, 4, 6

, we observed

that tannic acid, an oligomeric phenolic, exerted more inhibition on enzymatic hydrolysis than vanillin, a monomeric phenolic. There was no sugar released in the assays performed in the presence of tannic acid alone and in the mixture with vanillin, showing a complete inhibition of the hydrolysis. Interestingly, a major positive effect was observed when the mixture of vanillin and tannic acid was treated by adsorption onto the fly ash, achieving the same values of sugars as the hydrolysis using buffer solution only. This result suggests that there is a favored synergy during the adsorption process. Considering that the sizes and charges of these two compounds differs, it is plausible to suggest that they are occupying different sites of the adsorbent and making a more efficient use of it than when tannic acid is used alone at a higher concentration. Such explanation would also be in agreement with the significant percent removal of phenolics from LHW liquor obtained when using the adsorption onto fly ash (around 80%), since in such complex medium a great variety of phenolic compounds is found 24, 64, 65. Among the identified phenolic compounds originated from lignin degradation, tannic, gallic, caffeic, vanillic, p-coumaric, ferulic and cinnamic acids were detected in the washate of steam pretreated mixed hardwood 24. Besides the wide range of phenolic compounds, the concentration of total phenolics will also vary depending on the source of biomass as well as the pretreatment type and severity

64

. For instance, the concentration of phenolics in the

liquor of LHW pretreated sugarcane varied from 1.4 to 2.4 g/L as the temperature of the pretreatment process increased from 180 to 200 °C 66.


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Another important aspect to consider during selection of an adsorbent for the removal of inhibitors in the biomass conversion process is that the sugar loss should be either avoided and/or minimized during the adsorption. In order to investigate the possible loss of sugars, a set of adsorption experiments was carried out with glucose, xylose as well as the LHW liquor using fly ash and commercial activated carbon for comparison (Figure 8). The results showed that there was no adsorption of the pure sugars glucose and xylose when fly ash was used as adsorbent. Also for the adsorption of the LHW liquor onto fly ash, only a small difference of around 8% was observed when comparing to the initial sugar concentration. In contrast, for the commercial activated carbon the loss of sugars in the liquor due to adsorption was 71%. Such results further support the advantages of using fly ash as an effective adsorbent for the removal of phenolics compounds generated in the pretreatment step of the biomass conversion process. Other studies have reported improvements in enzymatic hydrolysis following the removal of phenolic compounds from the pretreatment liquor. Kim et al. 5 found that the use of activated carbon to remove phenolics from maple liquid pretreatment liquor increased the hydrolysis yield by 20%. Kim et al.

24

removed phenolics from steam pretreated mixed

hardwood using XAD-7, which increased the hydrolysis yield by 25%. Zhai et al.

23

also

reported increased yields in the hydrolysis of steam pretreated poplar and lodgepole pine, when phenolics were removed using activated carbon. The improvements in the enzymatic hydrolysis following phenolics removal can be explained by the reduction in the inhibitory effects of the compounds on the activity of cellulolytic enzymes

3, 4, 6

. Michelin et al.

66

found that the inhibition and deactivation of

cellulases and hemicellulases by water-soluble and acetone-extracted phenolics from LHW pretreated sugarcane bagasse depended on the type of enzyme and the method used to extract the phenolics from the bagasse. González-Bautista et al.

67

evaluated the activities of



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endoglucanase and xylanase in the presence of different concentrations of phenolic compounds released during saccharification of sugarcane bagasse. The endoglucanase activity showed 64% inhibition at 50 µg/mL of phenolic compounds, while the xylanase activity showed 77% inhibition at 100 µg/mL of phenolics. In order to reduce the inhibitory effects of the phenolic compounds, PVP and PEG (polymers) were added to the enzyme production process, with positive effects on the release of reducing sugars, because the binding of the polymers to the phenolic compounds acted to protect the enzymes. Despite the consensus in the literature about the need to reduce the inhibitory effects of phenolics in order to improve the enzymatic hydrolysis reactions of lignocellulosic biomass, implementation of most of the technological options that have been proposed would not be economically feasible in large-scale industrial processes. Therefore, it is clearly necessary to find more cost-effective strategies to mitigate the effects of the inhibitors of the biochemical reactions involved in lignocellulosic biomass conversion. In this work, the removal of phenolic compounds from LHW pretreatment liquor using sugarcane bagasse fly ash was found to be very efficient and provided a significant increase in the glucose released during the enzymatic hydrolysis reaction. The fly ash is a residual waste material generated in the sugarcane mill boilers after the incomplete carbonization of sugarcane bagasse, which is used as fuel in the boilers for energy production 32

. We show here that the sugarcane bagasse fly ash is a no-cost adsorbent with efficiency

comparable to that of a commercial activated carbon for the removal of phenolic compounds from sugarcane bagasse LHW pretreatment liquor. The proposed use of fly ash for removal of the inhibitors generated in the pretreatment of lignocellulosic biomass could also contribute to the implementation of different process configurations within the biorefinery. The improved efficiency of the enzymatic hydrolysis reactions to produce simple sugars could enable a better efficiency of the subsequent


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fermentation process to obtain ethanol or other bioproducts as well. Moreover, the removal of phenolics using the fly ash could also be applied to improve the use of the pentose sugars present in the liquor in a separated unit operation. This configuration would contribute to the use of the sugars from the hemicellulosic fraction, which is crucial to the efficient use of plant biomass within the biorefinery 68. Therefore, the findings reported here could be applied to support the use of sugars as a platform to obtain biofuels and different bioproducts by employing green and sustainable chemistry, which is being considered as the most promising route towards achieving the current demands for a future bio-based economy 69, 70.

Conclusions Removal of the phenolic compounds generated during the lignocellulosic biomass pretreatment process is required in order to minimize enzyme inhibition and improve the efficiency of the subsequent enzymatic hydrolysis. We have demonstrated here that sugarcane bagasse fly ash can provide highly effective removal of phenolic compounds from sugarcane bagasse LHW pretreatment liquor, resulting in an increase of 52% in the glucose concentration, comparable to the results obtained using a commercial activated carbon. Furthermore, the sugarcane bagasse fly ash was able to adsorb monomeric as well as oligomeric phenolic compounds. This characteristic is essential, because the liquors derived from different types of lignocellulosic feedstock and different pretreatment processes will contain a variety of phenolic compounds with different sizes and charges. Therefore, the use of sugarcane bagasse fly ash as an adsorbent for phenolics removal is highly attractive, especially since the fly ash is a waste material produced in significant amounts in sugarcane mills, so there is no cost involved in acquisition of the adsorbent. The findings reported here can potentially contribute to the implementation of more efficient large-scale industrial



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processing of lignocellulose biomass to produce sugars or other bioproducts, in compliance with the biorefinery concept.

Acknowledgements The authors would like to thank Embrapa, CNPq (Process 401182/2014-2), CAPES, and FAPESP (Processes 2014/19000-3 and 2016/10636-8) (all from Brazil) for their financial support. We are also grateful to the sugarcane mill Biorigin/Zilor (Lençóis Paulista, Brazil) for kindly providing the sugarcane bagasse fly ash.

References 1. Ko, J. K.; Kim, Y.; Ximenes, E.; Ladisch, M. R., Effect of liquid hot water pretreatment severity on properties of hardwood lignin and enzymatic hydrolysis of cellulose. Biotechnology Bioengineering 2015, 112 (2), 252-62. 2. Mosier, N.; Wyman, C.; Dale, B.; Elander, R.; Lee, Y. Y.; Holtzapple, M.; Ladisch, M., Features of promising technologies for pretreatment of lignocellulosic biomass. Bioresource Technology 2005, 96 (6), 673-686. 3. Ximenes, E.; Kim, Y.; Mosier, N.; Dien, B.; Ladisch, M., Inhibition of cellulases by phenols. Enzyme and Microbial Technology 2010, 46 (3–4), 170-176. 4. Ximenes, E.; Kim, Y.; Mosier, N.; Dien, B.; Ladisch, M. R., Deactivation of cellulases by phenols. Enzyme and Microbial Technology 2011, 48 (1), 54-60. 5. Kim, Y.; Ximenes, E.; Mosier, N. S.; Ladisch, M. R., Soluble inhibitors/desactivators of cellulase enzymes from lignocellulosic biomass. Enzyme and Microbial Technology 2011, 48 (4-5), 408-415. 6. Tejirian, A.; Xu, F., Inhibition of enzymatic cellulolysis by phenolic compounds. Enzyme and Microbial Technology 2011, 48 (3), 239-247. 7. Mhlongo, S. I.; Haan, R. d.; Viljoen-Bloom, M.; Zyl, W. H. v., Lignocellulosic hydrolysate inhibitors selectively inhibit/deactivate cellulase performance. Enzyme and Microbial Technology 2015, 81, 16–22. 8. Malgas, S.; Dyk, S. v.; Sagaran, A.; Pletschke, B. I., The inhibitory effects of various substrate pre-treatment by-products and wash liquors on mannanolytic enzymes. Journal of Molecular Catalysis B: Enzymatic 2016, 123, 132–140.


Page 23 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


9. Qin, L.; Li, W.-C.; Liu, L.; Zhu, J.-Q.; Li, X.; Li, B.-Z.; Yuan, Y.-J., Inhibition of lignin-derived phenolic compounds to cellulase. Biotechnology for Biofuels 2016, 9 (70). 10. Palmqvist, E.; Hahn-Hägerdal, B., Fermentation of lignocellulosic hydrolysates. II: inhibitors and mechanisms of inhibition. Bioresource Technology 2000, 74 (1), 25-33. 11. Olsson, L.; Hahn-Hägerdal, B., Fermentation of lignocellulosic hydrolysates for ethanol production. Enzyme and Microbial Technology 1996, 18 (5), 312-331. 12. Palmqvist, E.; Hahn-Hägerdal, B.; Galbe, M.; Zacchi, G., The effect of water-soluble inhibitors from steam-pretreated willow on enzymatic hydrolysis and ethanol fermentation. Enzyme and Microbial Technology 1996, 19 (6), 470-476. 13. Johnson, E., Integrated enzyme production lowers the cost of cellulosic ethanol. Biofuels, Bioproducts and Biorefining 2016, 10, 164-174. 14. Klein-Marcuschamer, D.; Oleskowicz-Popiel, P.; Simmons, B. A.; Blanch, H. W., The challenge of enzyme cost in the production of lignocellulosic biofuels. Biotechnology Bioengineering 2012, 109 (4), 1083-7. 15. Alriksson, B.; Horvath, I. S.; Sjöde, A.; Nilvebrant, N.; Jönsson, L. J., Ammonium Hydroxide Detoxification of Spruce Acid Hydrolysates. Aplied Biochemistry and Biotechnology 2005, 121-124, 911-922. 16. Alriksson, B.; Sjode, A.; Nilvebrant, N. O.; Jonsson, L. J., Optimal conditions for alkaline detoxification of dilute-acid lignocellulose hydrolysates. Applied Biochemistry Biotechnology 2006, 129-132, 599-611. 17. Alriksson, B.; Cavka, A.; Jonsson, L. J., Improving the fermentability of enzymatic hydrolysates of lignocellulose through chemical in-situ detoxification with reducing agents. Bioresource Technolology 2011, 102 (2), 1254-63. 18. Jurado, M.; Prieto, A.; Martinez-Alcala, A.; Martinez, A. T.; Martinez, M. J., Laccase detoxification of steam-exploded wheat straw for second generation bioethanol. Bioresource Technolology 2009, 100 (24), 6378-84. 19. Cao, G.; Ximenes, E.; Nichols, N. N.; Zhang, L.; Ladisch, M., Biological abatement of cellulase inhibitors. Bioresource Technolology 2013, 146, 604-10. 20. Nichols, N. M.; Sharma, L. N.; Mowery, R. A.; Chambliss, C. K.; Walsum, G. P. v.; Dien, B. S.; Iten, L. B., Fungal metabolism of fermentation inhibitors present in corn stover dilute acid hydrolysate. Enzyme and Microbial Technology 2008, 42 (7), 624-630. 21. Okuda, N.; Soneura, M.; Ninomiya, K.; Katakura, Y.; Shioya, S., Biological detoxification of waste house wood hydrolysate using Ureibacillus thermosphaericus for bioethanol production. Journal of Bioscience Bioengineering 2008, 106 (2), 128-33. 22. Zhu, J.; Yong, Q.; Xu, Y.; Yu, S., Detoxification of corn stover prehydrolyzate by trialkylamine extraction to improve the ethanol production with Pichia stipitis CBS 5776. Bioresource Technolology 2011, 102 (2), 1663-8.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 40

23. Zhai, R.; Hu, J.; Saddler, J. N., What Are the Major Components in Steam Pretreated Lignocellulosic Biomass That Inhibit the Efficacy of Cellulase Enzyme Mixtures? American Chemical Society 2016, 4 (6), 3429-3436. 24. Kim, Y.; Kreke, T.; Hendrickson, R.; Parenti, J.; Ladisch, M. R., Fractionation of cellulase and fermentation inhibitors from steam pretreated mixed hardwood. Bioresource Technology 2013, 135, 30–38. 25. Rengaraj, S.; Moon, S.-H.; Sivabalan, R.; Arabindoo, B.; Murugesan, V., Agricultural solid waste for the removal of organics: adsorption of phenol from water and wastewater by palm seed coat activated carbon. Waste Management 2002, 22 (5), 543-548. 26. Ahmaruzzaman, M.; Sharma, D. K., Adsorption of phenols from wastewater. Journal of Colloid and Interface Science 2005, 287 (1), 14-24. 27. Arellano-Cárdenas, S.; Gallardo-Velázquez, T.; Osorio-Revilla, G.; López-Cortéz, M. d. S.; Gómez-Perea, B., Adsorption of Phenol and Dichlorophenols from Aqueous Solutions by Porous Clay Heterostructure (PCH). Sociedad Química de México 2005, 49 (3), 287-291. 28. Mukherjee, S.; Kumar, S.; Misra, A. K.; Fan, M., Removal of phenols from water environment by activated carbon, bagasse ash and wood charcoal. Chemical Engineering Journal 2007, 129 (1–3), 133-142. 29. Makrigianni, V.; Giannakas, A.; Deligiannakis, Y.; Konstantinou, I., Adsorption of phenol and methylene blue from aqueous solutions by pyrolytic tire char: Equilibrium and kinetic studies. Journal of Environmental Chemical Engineering 2015, 3 (1), 574–582. 30. Purnomo, C. W.; Salim, C.; Hinode, H., Preparation and characterization of activated carbon from bagasse fly ash. Journal of Analytical and Applied Pyrolysis 2011, 91 (1), 257– 262. 31. Juang, R.-S.; Wu, F.-C.; Tseng, R.-L., Characterization and use of activated carbons prepared from bagasses for liquid-phase adsorption. Colloids and Surfaces A: Physicochemical and Engineering Aspects 2002, 201 (1–3), 191–199. 32. Lado, J. J.; Zornitta, R. L.; Calvi, F. A.; Tejedor-Tejedor, M. I.; Anderson, M. A.; Rotuolo, L. A. M., Study of sugar cane bagasse fly ash as electrode material for capacitive deionization. Journal of Analyticala and Applied Pirolysis 2016, 120, 389-398. 33. Batra, V. S.; Urbonaite, S.; Svensson, G., Characterization of unburned carbon in bagasse fly ash. Fuel 2008, 87, 2972-2976. 34. Srivastava, V. C.; Swamy, M. M.; Mall, I. D.; Prasad, B.; Mishra, I. M., Adsorptive removal of phenol by bagasse fly ash and activated carbon: Equilibrium, kinetics and thermodynamics. Colloids and Surfaces a-Physicochemical and Engineering Aspects 2006, 272 (1-2), 89-104. 35. Srivastava, V. C.; Prasad, B.; Mishra, I. M.; Mall, I. D.; Swamy, M. M., Prediction of breakthrough curves for sorptive removal of phenol by bagasse fly ash packed bed. Industrial & Engineering Chemistry Research 2008, 47 (5), 1603-1613. 24 ACS Paragon Plus Environment

Page 25 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


36. Shah, B.; Tailor, R.; Shah, A., Adaptation of Bagasse Fly Ash, a Sugar Industry Solid Waste into Zeolitic Material for the Uptake of Phenol. Environmental Progress & Sustainable Energy 2011, 30 (3), 358-367. 37. Sarkar, M.; Acharya, P. K., Use of fly ash for the removal of phenol and its analogues from contaminated water. Waste Management 2006, 26 (6), 559-570. 38. Gupta, V. K.; Sharma, S.; Yadav, I. S.; Mohan, D., Utilization of bagasse fly ash generated in the sugar industry for the removal and recovery of phenol and p-nitrophenol from wastewater. Journal of Chemical Technology and Biotechnology 1998, 71 (2), 180-186. 39. Silva, V. F.; Arruda, P. V.; Felipe, M. G.; Goncalves, A. R.; Rocha, G. J., Fermentation of cellulosic hydrolysates obtained by enzymatic saccharification of sugarcane bagasse pretreated by hydrothermal processing. Industrial Microbiololy Biotechnology 2011, 38 (7), 809-17. 40. Foo, K. Y.; Hameed, B. H., Insights into the modeling of adsorption isotherm systems. Chemical Engineering Journal 2010, 156 (1), 2–10. 41. Ghose, T. K., Measurement of cellulase activities. Pure & Aplied Chemistry 1987, 59 (2), 257-268. 42. Xu, B.; Chang, S. K. C., Total phenolic, phenolic acid, anthocyanin, flavan-3-ol, and flavonol profiles and antioxidant properties of pinto and black beans (Phaseolus vulgaris L.) as affected by thermal processing. American Chemical Society 2009, 57 (11), 4754–4764. 43. Miller, G. L., Use of Dinitrosalicylic Acid Reagent for Determination of Reducing Sugar. Analytical Chemistry 1959, 31 (3), 426–428. 44. IUPAC, Reporting physisorption data for gas/solid systems with special reference to the determination of surface area and porosity (Recommendations 1984). Pure & Applied Chemistry 1985, 57 (4), 603-619. 45. Hsieh, C.-T.; Teng, H., Influence of mesopore volume and adsorbate size on adsorption capacities of activated carbons in aqueous solutions. Carbon 2000, 38 (6), 863869. 46. László, K., Adsorption from aqueous phenol and aniline solutions on activated carbons with different surface chemistry. Colloids and surface A: physicochemical and engineering aspects 2005, 265 (1-3), 32-39. 47. Biniak, S.; Szymański, G.; Siedlewski, J.; Świtkowski, A., The characterization of activated carbons with oxygen and nitrogen surface groups. Carbon 1997, 35 (12), 17991810. 48. Ishizaki, C.; I., M., Surface oxide structures on a commercial activated carbon. Carbon 1981, 409-412. 49. Bansal, R. C.; Goyal, M., Activated Carbon Adsorption. CRC Press: Boca Raton, 2005; p 497 p.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 26 of 40

50. Ania, C. O.; Parra, J. B.; Pis, J. J., Effect of texture and surface chemistry on adsorptive capacities of activated carbons for phenolic compounds removal. Fuel Processing Technology 2002, 77-78, 337-343. 51. László, K.; Podkoscielny, P.; Dabrowski, A., Heterogeneity of Polymer-Based Active Carbons in Adsorption of Aqueous Solutions of Phenol and 2,3,4-Trichlorophenol. Langmuir 2003, 19, 5287-5294. 52. László, K.; Bóta, A.; Nagyu, L. G., Characterization of activated carbons from waste materials by adsorption from aqueous solutions. Carbon 1997, 35 (5), 593-598. 53. Terzyc, A. P., Further insights into the role of carbon surface functionalities in the mechanism of phenol adsorption. Journal of Colloid and Interface Science 2003, 268 (2), 301–329. 54. Villacañas, F.; Pereira, M. F. R.; Órfão, J. J. M.; Figueiredo, J. L., Adsorption of simple aromatic compounds on activated carbons. Journal of Colloid and Interface Science 2006, 293 (1), 128–136. 55. Lanning, F. C.; Ponnaiya, B. W. X.; Crumpton, C. F., The Chemical Nature of Silica in Plants. Plant Physiol 1958, 33 (5), 339-343. 56. Valix, M.; Cheung, W. H.; Zhang, K., Role of chemical pre-treatment in the development of super-high surface areas and heteroatom fixation in activated carbons prepared from bagasse. Microporous and Mesoporous Materials 2008, 116 (1-3), 513-523. 57. Adinaveen, T.; Kennedy, L. J.; Vijaya, J. J.; Sekaran, G., Studies on structural, morphological, electrical and electrochemical properties of activated carbon prepared from sugarcane bagasse. Journal of Industrial and Engineering Chemistry 2013, 19 (5), 14701476. 58. Xiao, G. Q.; Long, L. P.; Wang, J. L., Synthesis of the water-compatible pacetaminophen resin and its adsorption performances for vanillin in aqueous solution. Chinese Chemical Letters 2012, 23 (1), 123–126. 59. An, J.-H.; Dultz, S., Adsorption of tannic acid on chitosan-montmorillonite as a function of pH and surface charge properties. Applied Clay Science 2007, 36 (4), 256–264. 60. Furuya, E. G.; Chang, H. T.; Miura, Y.; Noll, K. E., A fundamental analysis of the isotherm for the adsorption of phenolic compounds on activated carbon. Separation and purification technology 1997, 11 (2), 69-78. 61. Moreno-Castilla, C.; Rivera-Utrilla, J., Carbon Materials as Adsorbents for the Removal of Pollutants from the Aqueous Phase. MRS Bulletin 2001, 26 (11), 890-894. 62. Singh, S.; Kushwaha, J. P., Tannic acid adsorption/desorption study onto/from commercial activated carbon. Desalination and Water Treatment 2013, 52, 3301-3311. 63. Liu, F.; Guo, Z.; Zheng, S.; Xu, Z., Adsorption of tannic acid and phenol on mesoporous carbon activated by CO2. Chemical Engineering Journal 2012, 183, 244–252.


Page 27 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


64. Du, B. W.; Sharma, L. N.; Becker, C.; Chen, S. F.; Mowery, R. A.; van Walsum, G. P.; Chambliss, C. K., Effect of Varying Feedstock-Pretreatment Chemistry Combinations on the Formation and Accumulation of Potentially Inhibitory Degradation Products in Biomass Hydrolysates. Biotechnology and Bioengineering 2010, 107 (3), 430-440. 65. Martin, C.; Galbe, M.; Nilvebrant, N. O.; Jonsson, L. J., Comparison of the fermentability of enzymatic hydrolyzates of sugarcane bagasse pretreated by steam explosion using different impregnating agents. Applied Biochemistry and Biotechnology 2002, 98, 699716. 66. Michelin, M.; Ximenes, E.; Polozeli, M. d. L. T. d. M.; Ladisch, M. R., Effect of phenolic compounds from pretreated sugarcane bagasse on cellulolytic and hemicellulolytic activities. Bioresource Technology 2016, 199, 275–278. 67. González-Bautista, E.; Santana-Morales, J. C.; Ríos-Fránquez, F. J.; Poggi-Varaldo, H. M.; Ramos-Valdivia, A. C.; Cristiani-Urbina, E., Phenolic compounds inhibit cellulase and xylanase activities of Cellulomonas flavigena PR-22 during saccharification of sugarcane bagasse. Fuel 2017, 196, 32–35. 68. Dodd, D.; Cann, I., Enzymatic deconstruction of xylan for biofuel production. Global Change Biology Bioenergy 2009, 1 (1), 2-17. 69. Sheldon, R. A., Green and sustainable manufacture of chemicals from biomass: state of the art. Green Chemistry 2014, 16 (3), 950-963. 70. Mohan, S. V.; Nikhil, G. N.; Chiranjeevi, P.; Reddy, C. N.; Rohit, M. V.; Kumar, A. N.; Sarkar, O., Waste biorefinery models towards sustainable circular bioeconomy: Critical review and future perspectives. Bioresource Technology 2016, 215, 2-12.



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Figure legends Figure 1. Differential pore volumes of the sugarcane bagasse fly ash and the activated carbon.

Figure 2. FTIR spectra (a), X-ray diffractograms (b), and zeta potentials (c) of the sugarcane bagasse fly ash and the commercial activated carbon.

Figure 3. Adsorption of phenolic compounds onto sugarcane bagasse fly ash at different pHs at 30 °C (a) and at different temperatures at pH 5 (b), at 30 rpm, using 2% (w/v) of adsorbent.

Figure 4. Isotherms for adsorption of vanillin (a) and tannic acid (b) onto sugarcane bagasse fly ash, and for adsorption of vanillin (c) and tannic acid (d) onto activated carbon. Adsorption conditions: 30 °C, pH 5, 30 rpm, for 24 h, using 1% (w/v) of adsorbent.

Figure 5. Time profiles for adsorption of vanillin (a) and tannic acid (b) onto sugarcane bagasse fly ash at 30 °C, pH 5, 30 rpm, using 5% (w/v) of adsorbent.

Figure 6. Enzymatic hydrolysis of sugarcane bagasse for 24 h at 50 °C and pH 5, before and after adsorption of phenolic compounds from LHW pretreated sugarcane bagasse liquor. CAC – commercial activated carbon. Adsorption conditions: 30 °C, pH 3, 30 rpm, for 24 h, using 5% (w/v) of adsorbent.

Figure 7. Relative concentration of reducing sugars (RS) released in the enzymatic hydrolysis reaction of a cellulose source (filter paper) for 1 h at 50 °C and pH 5, before and 28 ACS Paragon Plus Environment

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after adsorption of phenolic compounds onto fly ash. Adsorption conditions: 30 °C, pH 3, 30 rpm, for 24 h, using 5% (w/v) of adsorbent. Reference solution: 50 mM sodium citrate buffer, pH 4.8.

Figure 8. Adsorption of sugars from synthetic pure solutions (glucose and xylose) and from the LHW pretreated sugarcane bagasse liquor. Adsorption conditions: 30 °C, pH 3, 30 rpm, for 24 h, using 5% (w/v) of adsorbent. CAC – commercial activated carbon.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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Table 1. Pore structural properties of the sugarcane bagasse fly ash. Activated Property

Fly ash

Total BET surface area (m2/g) (SBET)

431

Micropore area (m2/g) (Smic)

325

76a

678

85a

External area (m2/g) (Sext)

105

24b

116

15b

Total pore volume (cm3/g) (VT)

0.20

Micropore volume (cm3/g) (Vmic)

0.13

65c

0.27

77d

External volume (cm3/g) (Vext)

0.07

35c

0.08

23d

Average pore diameter (nm)

1.89

a

(%)

carbon

(%)

794

0.35

1.80

Smic/SBET; bSext/SBET; cVmic/VT; dVext/VT.


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1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Table 2. Langmuir, Freundlich, and Redlich-Peterson constants for the adsorption of vanillin and tannic acid onto sugarcane bagasse fly ash and a commercial activated carbon. Sample

Model

R2

K (L/g)

Q (g/g)

n

a (L/g)

b

Langmuir

0.90

2.683±0.713

0.11±0.00

-

-

-

Vanillin

Freundlich

0.95

0.000

-

3.56±0.30

-

-

onto fly ash

Redlich0.96

0.989±0.552

-

-

12.1±7.6

0.80±0.04

Langmuir

0.99

0.292±0.039

0.16±0.01

-

-

-

Tannic acid onto

Freundlich

0.98

0.003±0.001

-

1.72±0.10

-

-

fly ash

Redlich0.98

0.058±0.018

-

-

0.5±0.04

0.82±0.21

Langmuir

0.82

19.322±7.354

0.32±0.01

-

-

-

Vanillin onto

Freundlich

0.89

0.001±0.000

-

4.98±0.66

-

-

activated carbon

Redlich0.88

32.306±43.647

-

-

121.8±183.5

0.83±0.05

Langmuir

0.95

5.689±1.080

0.06±0.00

-

-

-

Tannic acid onto

Freundlich

0.90

0.000

-

4.39±0.56

-

-

activated carbon

Redlich0.96

0.502±0.137

-

-

9.5±3.0

0.91±0.04

Peterson

Peterson

Peterson

Peterson


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Differential Pore Volume (cm³/g.nm)


0.12

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Activated Carbon Fly Ash

0.10 0.08 0.06 0.04 0.02 0.00 0

2

4

6

8 10 12 14 16 18 20 22 Pore width (nm)

Figure 1


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2905 3432

1389 1573 1095

471

Transmittance (%)

A)

600

Activated carbon Fly ash

1200 1800 2400 3000 3600 -1

Wavelength (cm )

B)

Intensity (a.u.)


0

10

20

30

40

50

60

70

80

2-Theta (degrees)

C) 10


0 Zeta Potential (mV)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


-10 -20 -30 -40 -50 0

1

2

3

4

5 6 pH

7

8

9 10 11

Figure 2 33 ACS Paragon Plus Environment


Percent phenolics removed (%)

A)

100 90 80 70 60 50 40 30 20 10 0

Vanillin Tannic acid

2

3

4 pH

5

6

B) Percent phenolics removed (%)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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100 90 80 70 60 50 40 30 20 10 0

Vanillin Tannic acid

30

35 40 45 Temperature (°C)

50

Figure 3


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Tannic Acid

0.14

0.14

(a)

0.12

0.12

0.10

0.10

0.08

0.08

Q (g/g)

Q (g/g)

0.06 Experimental data Freundlich Langmuir Redlich-Peterson

0.04 0.02 0.00

0

1

2

3

4

5

6

7

0.04

8

Experimental data Freundlich Langmuir Redlich-Peterson

0

1

2

3

4

5

6

7

8

Ce (g/L)

0.07

(c)

0.35

0.06

0.00

Ce (g/L)

0.40

(b)

0.02

(d)

0.06

0.30 0.05

Q (g/g)

0.25 Q (g/g)

Sugarcane bagasse fly ash

Vanillin

Activated carbon

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


0.20 0.15 Experimental data Freundlich Langmuir Redlich-Peterson

0.10 0.05 0.00 0

1

2

3

4

5

6

7

0.04 0.03

Experimental data Freundlich Langmuir Redlich-Peterson

0.02 0.01 8

0

Ce (g/L)

1

2

3

4

5

6

7

Ce (g/L)

Figure 4


8


A)

70 Vanillin 60

Q (mg/g)

50 40 30 20 Activated carbon Fly ash

10 0

0

30

60

90 120 150 180 210 Time (min)

B) 70 Tannic acid

60 50 Q (mg/g)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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40 30 20 Activated carbon Fly ash

10 0 0

30

60

90 120 150 180 210 Time (min)

Figure 5


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12 11 10

Concentration (g/L)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


9 8

L TLFA

Untreated Liquor Liquor treated by fly ash

Phenolics Glucose

TLCAC Liquor treated by CAC

Phenolics removal

Enzymatic hydrolysis

7 6 5 4 3 2 1 0

L

TLFA

TLCAC

L

TLFA

TLCAC

Figure 6



Initial (g/L)

Relative RS concentration (%)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

After fly ash (g/L)

Vanillin

Tannic acid

1/2 van + 1/2 tannic

3.06±0.11

2.80±0.08

2.85±0.02

0.19±0.01

0.36±0.01

0.02±0.01

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Untreated Treated by fly ash

100 80 60 40 20 0 Vanillin

Tannic acid

Van+Tannic acid

Figure 7


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20 18 16

Concentration (g/L)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Initial concentration After fly ash After CAC

14 12 10 8 6 4 2 0

Glucose

Xylose

RS- liquor

Figure 8



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For Table of Contents Use Only

Graphical Abstract

Synopsis The use of sugarcane bagasse fly ash as a no-cost adsorbent of phenolic compounds increased the efficiency of enzymatic hydrolysis of lignocellulosic biomass, contributing to implementation of future biorefineries.


Sugarcane bagasse fly ash as a no-cost adsorbent for removal of

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