Chromatographic Detection of LigninâCarbohydrate Complexes in

Article pubs.acs.org/Biomac

Chromatographic Detection of Lignin−Carbohydrate Complexes in Annual Plants by Derivatization in Ionic Liquid Anika Salanti, Luca Zoia,* Eeva-Liisa Tolppa, and Marco Orlandi Department of Environmental and Earth Sciences, University of Milano-Bicocca, Piazza della Scienza 1, Milan, I-20126 Italy ABSTRACT: The opportunity for detecting the presence and the amount of lignin−carbohydrate complexes (LCCs) in renewable feedstocks is a major issue for the complete utilization of biomass. Indeed, LCCs are known to shield cellulose from enzymatic hydrolysis, reducing the efficiency of the digestion processes needed for the production of biobased products. This study is focused on the chromatographic characterization of lignocellulose from agricultural residues (rice husk, wheat straw) and herbaceous energy crops (Arundo donax, Miscanthus sinesis) and their fractionation products (hemicellulose, cellulose, and lignin). Exploiting alternative chemical derivatizations on the aforementioned samples, it was possible to discern the connectivity among the various lignocellulosic components. The complete acetylation and benzoylation of the milled native substrates in ionic liquid media, and the systematic comparison between their GPC-UV chromatograms collected at different wavelengths has revealed itself as a straightforward technique in the detection of LCCs. This novel approach proved an extensive connectivity between the lignin and the hemicellulosic for all the analyzed specimens, whereas the cellulosic fraction was conceived as a substantially unbound moiety, accounting for the sample composition at higher molecular weights. Moreover, the collected lignin fractions were extensively characterized by means of 31P NMR and 2D-HSQC techniques.

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INTRODUCTION Traditionally, two types of materials have been regarded as renewable feedstock: woody-based and agricultural biomasses. In recent years, lignocellulosic biomasses from agricultural residues and herbaceous energy crops have been under intense investigation due to their annual renewability and large annual biomass stock.1 As opposed to fossil fuels, the herbaceous lignocellulosic biomass represents a promising alternative in the production of biofuels due to its naturally high content of fermentable reducing sugars.2,3 Lignocellulose is an extremely structured natural material made up of three main biopolymers: cellulose, hemicellulose, and lignin. Cell wall cellulose consists of linear chains of β(1− 4) linked D-glucopyranose units. It is difficult to break down into glucose because of its extensive H-bonded network and highly organized crystalline structure. Hemicellulose is a carbohydrate heteropolymer composed of several different sugars including five-carbon and six-carbon sugars and could be easily broken down into its building blocks. Lignin is a complex and irregular polymer network, composed of randomly crosslinked phenylpropanoid units,4,5 that acts as a glue holding cellulose and hemicellulose together. Lignocellulose biorefinery generally includes three fundamental steps: first, a pretreatment to fractionate the recalcitrant lignocellulose structure; second, the enzymatic hydrolysis of the isolated cellulose moiety to obtain fermentable sugars; and third, the fermentation to produce cellulosic ethanol or other biobased chemicals.6 Because of the resistant structure of crystalline cellulose and natural composite structure of lignocellulose, efficient pretreat© 2012 American Chemical Society

ment technologies are needed prior to the enzymatic hydrolysis. The recalcitrance of lignocellulosic materials to enzymatic hydrolysis is substantially attributed to the low accessibility of crystalline cellulose fibers, which restricts cellulase activity.7,8 The presence of lignin and hemicellulose on the surface of cellulose prevents cellulase from accessing the substrate. It is recognized that enzymes’ performance is reduced during lignocellulose hydrolysis by interaction with lignin and, especially, lignin−carbohydrate complexes (LCCs).9 There is consensus among scientists that lignin is cross-linked to different polysaccharides in the cell wall, but the basic nature of these interactions is still unclear. A lot of studies have been done from the first paper in the field.10 The most recent advances were based on the isolation11,12 and characterization13,14 of lignin−carbohydrate complexes from woody materials by Lawoko et al. A number of different approaches have been proposed so far for lignocellulosic pretreatment aimed at the removal of lignin (either in the form of LCCs or free polyphenols), including biological, chemical, physical, and thermal processes. However, all of them result in a substantial loss in fermentable sugars content of the residual polysaccharides.15 In the past few years, the development of ionic liquids and their application as green solvents for the pretreatment and fractionation of lignocellulosic biomass has led to intensive research, which allowed the selective extraction of a chemically Received: October 20, 2011 Revised: December 22, 2011 Published: January 5, 2012 445

dx.doi.org/10.1021/bm2014763 | Biomacromolecules 2012, 13, 445−454

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Figure 1. Fractionation and analytical scheme.

materials. The application of the method in the detection of LCCs was based on the comparison among the molecular weight distributions of the entire material and its fractions (namely holocellulose and lignin).Valuable information about elusive LCCs could be achieved by means of this approach, but a fractionation of the examined substrate could not be avoided. In this paper we propose a straightforward chromatographic method for the detection of LCCs based on the benzoylation and acetylation of the whole lignocellulose specimen. Extensively ball-milled samples of four herbaceous substrates (rice husk, wheat straw, Arundo donax, and Miscanthus sinesis) were dissolved in the ionic liquid 1-allyl-3-methylimidazolium chloride ([amim]Cl) and then reacted with either benzoyl chloride or acetyl chloride in the presence of pyridine under mild conditions. Both the highly substituted lignocellulosic esters exhibited an enhanced solubility in tetrahydrofuran (THF), but developed a different instrumental response when submitted for GPC-UV analysis. Benzoylated specimens enabled the UV-detection of all substrate components,24,25 namely, cellulose, hemicellulose, and lignin, regardless of possible chemical connection among them, whereas acetylated specimens accounted for the sole contribution of LCCs (and of course possibly free lignin) due to the lack of chromophores in the unbound acetylated polysaccharide portion. This effect has been exploited for the direct detection of LCCs in acetylated, unprocessed samples by GPC-UV, thanks to the high instrumental response of naturally occurring aromatic rings, that is, polysaccharides-linked polymeric and oligomeric lignin, and related monomers as well. Moreover, the GPC-UV analysis of each cellulosic and hemicellulosic fraction offered a valuable method to establish the nature of both free and LCC-bound polysaccharides. In conclusion, the method described in this paper modifies the previous one (Zoia et al.)24 to develop an improved methodology useful in lignocellulosic characterization. Furthermore, an exhaustive chromatographic and spectroscopic characterization of each extracted lignin is provided as a preliminary investigation.

unaltered lignin and simultaneously yielded an unaltered, highly biodegradable cellulose fraction.16−19 Ionic liquids (ILs) are defined as organic salts that melt below 100 °C and are entirely composed of ions, typically large organic cations and small inorganic anions. The most promising ILs in the dissolution of lignocelluloses are imidazolium-based chloride salts: the cations are able to solvate the aromatic character of lignin by means of π−π interaction, whereas the chloride counteranion is usually the most effective in disrupting the extensive inter- and intramolecular hydrogen bonds pattern of cellulose microfibrils.20−22 For these reasons, the aforementioned approach requires mild reaction conditions and is expected to decrease sugar degradation, inhibitor formation, processing costs, and capital investments thanks to the recycling opportunity associated with ionic liquids. Nevertheless, the presence of LCCs could not be avoided due to their intrinsic nature, that is, a covalent bond connecting a polysaccharide chain to a lignin moiety. Indeed, whereas a fairly large lignin fraction could definitely be solubilized and removed from the lignocellulosic substrate,16−18 polysaccharides are regenerated from the ionic liquid solution after the addition of an antisolvent such as water or ethanol.23 Even if the regenerated lignocellulosic material possesses a relatively homogeneous and amorphous morphology which contributes to the enhancement of its enzymatic digestibility, it would be widely useful to have available an experimental methodology able to describe the extent of connection between lignin and polysaccharides. Such information would be of considerable importance when choosing the most appropriate pretreatment in order to maximize the yield of fermentable sugars while minimizing the processing time and cost. Recently, Zoia et al.24 reported a methodology based on comparative GPC analyses of benzoylated intact wood and correspondent benzoylated holocellulose and lignin fractions. This method is based on ionic liquid solubilization and homogeneous reaction with benzoyl chloride for complete derivatization. The reaction improved the solubility of the entire material in THF and enabled its detection by GPC-UV, due to the abundant presence of aromatic groups in the derivatized materials. The method allowed for the first time the direct analysis of molecular weight distribution of woody

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MATERIALS AND METHODS

The analytical protocol applied in this work is reported in Figure 1. 446


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Reagents and Materials. All the reagents were purchased from Sigma-Aldrich and used as received without further purification. Rice husk was kindly provided by a local factory, Gariboldi S.p.A.. Arundo donax and Wheat straw samples were supplied by a local factory. Miscanthus sinensis samples were kindly provided during COST action FP0901. The four herbaceous substrates (10 g) were crushed in a blender for 5 min and passed through a 1 mm screen. The ground material thus obtained was Soxhlet extracted with about 250 mL of acetone for 24 h. The dry, extractives-free samples (3 g) were milled in a planetary ball mill for 20 h at 300 rpm, using a 100 mL zirconiumgrinding bowl (zirconium dioxide 95%) in the presence of six zirconium balls (10 mm in diameter each). Lignin Content. The amount of total lignin was calculated as the sum of the acid-insoluble (Klason lignin) and acid soluble lignin content, measured according to the method reported by Yeh et al.26 The values reported are the average of three analyses ±1.0% (P = 0.05, n = 3). Ash Content. Accurately weighed and dried samples (100 mg) were put in tared, well-desiccated porcelain crucibles and placed in a muffle furnace set at 550 °C for 3 h. The crucibles were then stored in a desiccator until room temperature was reached. The ash content was determined gravimetrically. Acidolysis Lignin. The lignin extraction was performed according to a modification of the milled wood method developed by Holmbom and Stenius.27 The powder (1 g) was refluxed under a nitrogen atmosphere for 2 h in a 0.1 M HCl dioxane−water solution (30 mL, 85:15) and then allowed to cool to room temperature. The insoluble material left after lignin solubilization was collected by centrifugation (3000 rpm, 15 min). The supernatant was added dropwise into a 0.01 M HCl aqueous solution (250 mL), which was then kept at +4 °C overnight to allow for a complete lignin precipitation. The precipitate was collected by centrifugation (3000 rpm, 15 min), washed with acidified distilled water (pH 2), and freeze-dried. Lignin Acetylation. A total of 60 mg of extracted lignin was acetylated in a pyridine−acetic anhydride solution (1:1 v/v, 4 mL) and kept overnight at 40 °C. After stripping with ethanol, toluene, and chloroform (25 mL × 3, each solvent), the sample was dried in vacuum. The acetylated lignin has been solubilized in THF for GPC analysis and in DMSO-d6 for 2D-HSQC-NMR analyses. Lignin: 31P NMR Derivatization. Accurately weighed lignin samples (30 mg) were dissolved in a pyridine−deuterated chloroform solution (1.6:1 v/v mL, 800 μL) containing 1 mg/mL of chromium(III) acetylacetonate (Cr(acac)3). Then 100 μL of an e-HNDI solution (121.5 mM, CDCl3/pyridine 4.5:0.5) was added, along with 100 μL of 2-chloro-4,4,5,5-tetramethyl-1,3,2-dioxaphospholane as the derivatizing agent28 to quantitate the amount of different labile hydroxyl groups (aliphatics, phenolics, and acidic). Furthermore, the same procedure was employed for the specific determination of aliphatic hydroxyl groups derived from β-O-4 moieties using 100 μL of 2-chloro-1,3,2dioxaphospholane as the phosphorus derivatizing agent.29 31P NMR spectra were recorded of 800 μL samples on a Bruker Avance 500 MHz instrument. The 31P NMR data reported in this paper are the average of three experiments. The maximum standard deviation was 2 × 10−2 mmol/g, while the maximum standard error was 1 × 10−2 mmol/g. Preparation of Holocellulose. Holocellulose was prepared according to a modification of the method outlined by Chang et al.30 Approximately 2 g of severely blended herbaceous material was placed in a 250 mL Erlenmeyer flask and allowed to soak in 60 mL of water. Afterward, 60 mL of a NaClO2 aqueous solution was added and 25 mL of glacial acetic acid was slowly incorporated under stirring. After being covered with a small inverted Erlenmeyer, the mixture was heated to 90 °C. After 2 h, the solid residue was filtered in a sintered glass crucible and oven-dried. Extraction of Hemicellulose and α-Cellulose..31 About 1 g of holocellulose was transferred in a three-neck wide-mouth 250 mL round-bottom flask. Using a dropping funnel, 50 mL of 5% aqueous potassium hydroxide solution and 0.014 g of NaBH4 were added under a nitrogen atmosphere and constantly stirred to extract hemicellulose A. After 2 h, the mixture was filtered off through a sintered funnel and

the filtrate was acidified with glacial acetic acid until pH 5−6 was reached. The solid residue was treated in the same way for the extraction of hemicellulose B, but 24% of aqueous potassium hydroxide was used instead. The acidified filtrate was then added to the previously recovered hemicellulose A solution. After the addition of ethanol (200 mL), the solution was kept at +4 °C overnight to allow for hemicellulose precipitation. The exceeding supernatant liquor was removed with a vacuum-assisted pipet. The precipitate was then recovered by centrifugation (3000 rpm, 15 min), washed with ethanol, and freeze-dried. The insoluble residue that was left, designated as α-cellulose, was thoroughly washed with deionized water and ethanol and dried with diethyl ether. Enzymatic Hydrolysis. The milled samples from RH, AD, MS and WS were treated with cellulase (from Tricoderma resei, ATCC 26921, Sigma-Aldrich). The enzymatic hydrolyses were carried out at 40 °C for 48 h using 50 mM citrate buffer (pH 4.5) at 5% consistency on an orbital water bath shaker. The insoluble material that remained after the enzymatic hydrolysis was collected by centrifugation (3000 rpm), washed twice with acidified deionized water (pH 2), and freeze-dried. The weight loss percentage was calculated, dividing the weight of the recovered materials by the weight of the starting materials. Milled Herbaceous Plants: Acetylation in Ionic Liquid. Ionic liquid, 1-allyl-3-methylimidazolium chloride ([amim]Cl, 950 mg), was added to the pulverized herbaceous material (50 mg) in a 8 mL dried sample bottle equipped with a magnetic stirrer, vortexed, and heated at 80 °C until the solution was clear (18 h, overnight). After the addition of pyridine (350 μL, 4.3 mmol) the solution was vortexed until homogeneous and allowed to cool to room temperature. Acetyl chloride (250 uL, 3.5 mmol) was added in one portion, and the mixture was vortexed until a homogeneous yellow paste was formed. Afterward, CHCl3 was added in two portions (250 μL each) and the mixture was vortexed as described by King et al.32 Additional CHCl3 (two portions, 2 mL each) was included, and the sample was heated at 50 °C for 5 min. The sample was transferred in a 100 mL roundbottom flask with additional CHCl3 to ensure complete recovery of the entire sample; CHCl3 was then removed under rotary evaporation. Then a deionized water−ethanol solution (1:3, 20 mL) was added to induce the product precipitation, and the mixture was vigorously shaken and vortexed for 2 min. The solid was filtered off through a sintered funnel (grade 3), washed with additional ethanol, and purified with methanol. The acetylated samples were solubilized in THF, filtered through a 0.45 μm GHP Acrodisc syringe filter, and subjected to GPC analysis. Milled Herbaceous Plants: Benzoylation in Ionic Liquid. Ionic liquid, 1-allyl-3-methylimidazolium chloride ([amim]Cl, 950 mg), was added to the pulverized herbaceous material (50 mg) in an 8 mL dried sample bottle equipped with a mechanical stirrer, vortexed, and heated at 80 °C until the solution was clear (18 h, overnight). Pyridine (350 μL, 4.3 mmol) was added; the solution was vortexed until homogeneous and allowed to cool to room temperature. Then benzoyl chloride (380 μL, 3.3 mmol) was added in one portion and vortexed until a homogeneous paste was formed. The sample was kept under magnetic stirring at room temperature for 2 h. To precipitate the benzoylated product, a deionized water−ethanol solution (1:3 v/v, 20 mL) was added and the mixture was vigorously shaken and vortexed for 5 min. The solid was filtered off through a sintered funnel (grade 3), washed with additional ethanol, and purified with methanol. The benzoylated samples were solubilized in THF and passed through a 0.45 μm GHP Acrodisc syringe filter for GPC analyses. GPC Analyses. The analyses were performed on a Waters 600 E liquid chromatography connected to a HP1040 ultraviolet UV detector set at 240 or 280 nm. The injection port was a Rheodyne loop valve equipped with a 20 μL loop. The GP-column system was composed by a sequence of an Agilent PL gel 5 μm, 500 Ǻ , and an Agilent PL gel 5 μm, 104 Ǻ . The solvent used was tetrahydrofuran (Fluka 99.8%). PL Polymer Standards of Polystyrene from Polymer Laboratories were used for calibration. The evaluation of the number-average molecular weight (Mn) and the weight-average molecular weight (Mw) of the extracted lignin samples was performed according to the methodology developed by Himmel.33 The peak molecular weight Mp is defined as 447


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the molecular weight of the species with maximum absorbance. Moreover, the ratio I = Mw/Mn, defined as Polydispersity Index was also calculated. The Mn, Mw, and Mp values reported are the average of three analyses (Mw: 1000 g/mol; Mn, Mp: 70 g/mol, P = 0.05, n = 3). Acetylated and benzoylated samples were dissolved in THF (1 mg/ mL) and analyzed at a flow rate of 1 mL/min. 2D-HSQC-NMR Analyses. 2D-HSQC spectra were run in DMSOd6 on acetylated samples to avoid the fractionation of the material before NMR analysis and to increase both solubility and chemical shift dispersion of the side chain units.34 The inverse detected 1H−13C correlation spectra (HSQC) were measured on a Varian Mercury 400 MHz instrument at 308 K. The spectral width was set at 5 kHz in F2 and 25 kHz in F1. Altogether 128 transients in 256 time increments were collected. The polarization transfer delay was set at the assumed coupling of 140 Hz and a relaxation delay of 2 s was used. The spectra were processed using Π/2 shifted squared sinebell functions in both dimensions before Fourier transformation.35 The assignment of the predominant signals was based on the chemical shift data of lignin model compounds and milled wood lignin (MWL), as reported in literature.36,37

Figure 3. Overlapped GPC-UV profiles of cellulose and lignin, acetylated and benzoylated at 1 mg/mL concentration.

Table 2. Compositional Evaluation of Rice Husk,33 Arundo donax, Wheat Straw, and Miscanthus sinensis Expressed as Klason Lignin, Ash, and Holocellulose Percentage Content

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RESULTS AND DISCUSSION Setup of the Chromatographic Method. To set up the chromatographic system for the detection of LCCs, UV spectra

lignin (%) ash (%) holocellulose (%)

Table 1. Results of UV-Vis Analyses of Acetylated and Benzoylated Cellulose and Lignin Samples Extracted from Rice Huska sample cellulose acetylated cellulose benzoylated lignin acetylated lignin benzoylated

ε 240 nm (mL/mg *cm)

ε 280 nm (mL/mg *cm)

0.1

4.00

0.96

0.1

30.37

9.43

0.1 0.1

25.65 34.12

19.13 16.07

Arundo donax

Miscanthus sinensis

wheat straw

21.8 16.0 62.2

29.9 4.7 65.4

25.6 3.1 71.3

18.6 9.4 72.0

both), while the spectrum of acetylated cellulose was not well resolved, with a weak and broad absorbance around 330 nm. At 240 nm, benzoylated cellulose and lignin presented similar ε values (30.37 vs 34.12, Table 1), whereas at 280 nm, the acetylated cellulose sample showed a very low ε value, with an instrumental response 20 times lesser than lignin (0.96 vs 19.13, Table 1). On the basis of these results, we decided to choose 240 nm as the recording wavelength for GPC analysis of benzoylated samples and 280 nm as the recording wavelength for GPC analysis of acetylated samples. The GPC analyses of acetylated and benzoylated cellulose and lignin (concentration 1 mg/mL) are reported in Figure 3. As expected, the chromatograms highlighted that after benzoylation both cellulose and lignin are readily detected at 240 nm with a similar instrumental response, while after acetylation at 280 nm, lignin showed a much higher response in comparison to the acetylated cellulose. This is related to the absence of strong chromophores in the polysaccharide fraction after acetylation and represents the key point for the recognition of LCCs in native substrates. Residual absorbance of cellulose after acetylation was ascribed to byproduct formed after reaction with either pyridine or the ionic liquid. Starting with the aforementioned results, this paper proposes a chromatographic method for the detection of LCCs based on the acetylation and benzoylation of the whole lignocellulose specimens in ionic liquid media. Both of the highly substituted lignocellulosic esters exhibited an enhanced solubility in tetrahydrofuran (THF) but developed a different instrumental response when submitted for GPC-UV analysis. Benzoylated specimens (240 nm) enabled the UV-detection of all substrate components,24,25 namely, cellulose, hemicellulose, and lignin, regardless of possible chemical connection among them, whereas acetylated specimens (280 nm) accounted for the sole contribution of LLCs, and possibly free lignin, due to the lack of chromophores in the unbound acetylated polysaccharide portion. GPC Analyses of the Annual Plants. This study is focused on the developed GPC-UV chromatographic detection

Figure 2. UV spectra 230−430 nm of cellulose and lignin, acetylated and benzoylated at 0.1 mg/mL concentration.

concentration (mg/mL)

rice husk

a

The extinction coefficient was calculated according to the LambertBeer law.

of acetylated and benzoylated samples of cellulose and lignin extracted from Rice Husk were acquired between 230 and 430 nm. The UV spectra are reported in Figure 2. The concentration of each specimen was 0.1 mg/mL in THF. The spectra of acetylated and benzoylated lignin, along with the spectrum of benzoylated cellulose showed different absorption bands between 230 and 380 nm. The two major bands in these samples were ascribed to the presence of aromatic units (either from lignin itself or the derivatization with benzoyl chloride or 448


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were then analyzed by GPC-UV at 240 and 280 nm, respectively, to maximize their analytical response. The resulting chromatograms are reported in Figure 4. As previously outlined, after benzoylation, polysaccharides and lignin have a similar instrumental response. Therefore, the chromatograms of benzoylated samples are supposed to report the molecular weight distribution of the whole material (cellulose, hemicelluloses, lignin, and LCCs).24 On the other hand, acetylated sample chromatograms almost exclusively account for the molecular weight distribution of those lignocellulosic fractions that naturally contain aromatic groups (LCCs and free lignin) due to the higher instrumental response observed for lignin compared to free polysaccharides. The chromatograms of acetylated and benzoylated materials were then overlapped: the comparison between them gave a clue about the presence of polysaccharides whether or not connected to lignin. For all the analyzed samples (RH, AD, WS, and MS), the chromatograms of benzoylated ones showed a pronounced shoulder in the higher molecular weight region than the acetylated ones, supporting the presence of free polysaccharides (Figure 4). The profile of acetylated samples in Figure 4 showed the presence of moieties with a larger molecular weight in regard to the corresponding extracted lignin specimen. These components, detected at 280 nm, could be associated with the presence of polysaccharides extensively connected with aromatic compounds. However, the GPC technique herein developed was not able to discern the nature of aromatic compounds covalently bounded to the polysaccharide fraction. They may be ascribed either to lignin (aromatic polymer) or to p-coumaric and ferulic acids (phenolic aromatic compounds), known to be connected by ester bonds to hemicellulose.38 Moreover, the presence of different shoulders in the region of lower molecular weights, noticed in all the benzoylated and acetylated chromatograms, could be related to the presence of free hemicellulose (detectable only after benzoylation at 240 nm) and free lignin (detectable in any instrumental configuration). Afterward, the main lignocellulosic fractions (namely, cellulose, hemicellulose, and lignin) of all four herbaceous samples were extracted and derivatized (benzoylated and acetylated) in IL. They were then subjected to GPC analysis, with the aim to rationalize composition and components distribution in each chromatogram describing the whole derivatized samples. The results are reported in Figure 5. Pure cellulose and hemicellulose samples were both acetylated and benzoylated. In the case of cellulose samples, Figure 5 reports only benzoylated chromatograms due to the low instrumental response obtained from acetylated ones, according to the observation discussed in the first section (Setup of the Chromatographic Method). Where hemicellulose was concerned, both acetylated and benzoylated specimens were detectable by GPC-UV, giving a similar molecular weight distribution. Such observation itself may be considered evidence of the presence of LCCs, and may be justified by the presence of any aromatic compound bounded to the hemicellulose structure and not removed by the preliminary oxidative step involving NaClO2. For the purposes of this study, we decided to compare the acetylated chromatograms of hemicellulose to the acetylated chromatograms acquired after the derivatization of all materials. It is worth noting that the presence of residual aromatic compounds bounded to the hemicellulose fraction did not affect the molecular weight

Figure 4. Overlapped GPC-UV profiles of native milled benzoylated (black line) and native milled acetylated (gray line) herbaceous materials under investigation (top to bottom: rice husk, RH; Arundo donax, AD; wheat straw, WS; and Miscanthus sinensis, MS). The chromatograms of the acidolysis lignin samples (acetylated, dotted line) are also reported as reference signals.

of lignocelluloses from agricultural residues (rice husk, wheat straw), herbaceous energy crops (Arundo donax, Miscanthus sinesis), and their fractionation products (hemicellulose, cellulose, and lignin). As a preliminary investigation, ash, lignin, and polysaccharide content in the starting materials was determined. Results are reported in Table 2. Moreover, extensively ball-milled samples of the four herbaceous plants were dissolved in the ionic liquid 1-allyl-3methylimidazolium chloride ([amim]Cl) and then reacted with benzoyl chloride or acetyl chloride in the presence of pyridine under mild conditions. Benzoylated and acetylated samples 449


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Figure 5. Identification of the main polysaccharide components in the UV-detected fractions of benzoylated and acetylated native samples (Clockwise: rice husk, RH; Arundo donax, AD; wheat straw, WS; and Miscanthus sinensis, MS). Top: overlapped GPC-UV profiles of native milled benzoylated (black line) and benzoylated extracted cellulose (gray line). Bottom: overlapped GPC-UV profiles of native milled acetylated (black line), acetylated extracted hemicellulose (gray line), and acetylated acidolysis lignin (dotted line).

almost the same Mp, except for a sharper peak for RH and WS, while in the case of AD and MS, the distributions were sharper and completely shifted at lower molecular weight with respect to the corresponding native material. In the case of RH and WS, the slight differences in the overlapped chromatograms may be ascribed to the abundant presence of naturally bonded p-coumaroyl and feruloyl esters on the hemicellulose fraction, comprehensively resulting in low molecular weight LCCs (specifically phenolics−polysaccharide bonds). As far as AD and MS are concerned, the molecular weight distribution of acetylated native samples is located much higher in regards to the corresponding hemicellulose fractions. It is like a fraction is missing: a fraction in the highest molecular weight region, which was interpreted as the LCCs (specifically lignin−carbohydrate complexes). The fractionation process cleaved away the lignin moiety by a progressive oxidation, thus,

distribution due to their limited contribution to the polymer molecular weight. Looking at the benzoylated chromatograms in Figure 5, where the comparison between all the native materials and the corresponding extracted cellulose is reported, it was possible to verify a significant overlapping of the two GPC profiles in the higher molecular weight area, thus, confirming that the polysaccharide describing the higher molecular weight fractions of all the samples involved ought to be identified with cellulose. The same results have already been observed in a study concerning the comparison among native specimens of benzoylated Eucalyptus grandis, Norway spruce, corn stover, and holocellulose samples.24 On the other hand, Figure 5 showed a substantial disagreement between the chromatograms of acetylated native materials and the corresponding extracted hemicelluloses. In fact, the extracted hemicellulose specimens were described by 450


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Figure 6. Overlapped GPC-UV profiles of the enzymatically treated herbaceous materials under investigation (Clockwise: rice husk, RH; Arundo donax, AD; wheat straw, WS; and Miscanthus sinensis, MS). Molecular weight distributions of benzoylated (black line) and acetylated (gray line) samples after recovery. Chromatographic profiles of each acidolysis lignin (acetylated, dotted line) are also reported as reference signals.

milled native specimens were hydrolyzed by cellulase T. reesei. The recovered materials were acetylated and benzoylated and then submitted to GPC characterization. The chromatograms are reported in Figure 6. The results from gravimetric analyses showed that RH, AD, MS, and WS lost 36, 49, 52, and 72% in weight, respectively, after the cellulolytic treatment. RH showed recalcitrance to the cellulase hydrolysis, AD and MS were extensively hydrolyzed, while WS was almost completely hydrolyzed (holocellulose contents 62, 65, 71, and 72%, respectively, as reported in Table 2). As a consequence of the extensive milling of the samples, the cellulose crystallinity was assumed to be completely destroyed, facilitating the enzymatic digestion. These gravimetric results were moreover confirmed by GPC. The benzoylated and acetylated chromatograms reported in Figure 6 showed that, for the recovered WS sample, almost all the sugars were hydrolyzed, leaving residual GPC profiles very close to the extracted lignin ones. In the case of AD and MS, the comparison between benzoylated and acetylated GPC profiles proved the presence of residual polysaccharides not completely hydrolyzed, but mainly as the shoulder in the higher molecular weight region of the benzoylated specimens. Finally, the benzoylated and acetylated

leaving a missing contribution in molecular weight that could be explained only by taking into account an extensive chemical connection between hemicellulose and the lignin polymer. The experimental observations are in agreement with the present knowledge about ferulates in grasses.38−40 The presence of naturally bonded feruloyl esters on the hemicellulose fraction is well-known indeed (phenolics−polysaccharides bonds, as detected in RH and WS).41 Moreover, these structures could undergo dimerization producing a whole range of diferulates and result in polysaccharide−polysaccharide cross-linking.42 Cross-coupling of ferulates with lignin monomers incorporates them into lignin and results in lignin− polysaccharide cross-linking.43 Moreover, there is evidence that ferulates could act as nucleation sites for lignification in grasses.44 The predominant of each of these cross-linking mechanisms could be detected and rationalized by our GPC system. This kind of information is really important because the lignin−polysaccharide complexes negatively impact the availability of polysaccharides for the production of bioethanol9 or animal feeds.45 Applications. To apply the developed GPC system and observe the role of LCCs during enzymatic hydrolysis, the 451


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chromatograms of the recovered RH sample endorsed the presence of a high amount of polysaccharides, especially connected to aromatic moieties (shown by the acetylated profile) but also in a free form (shown by the benzoylated profile). Lignin Characterization. The extracted lignin specimens were thoroughly characterized by means of GPC, 2D-HSQC, and 31P NMR analyses.46 Results are reported in Table 3, which displays similar results for each sample. The isolated lignins were a p-coumaryl, guaiacyl, and syringyl lignins, typical of herbaceous species, with each of the three main constituent units well represented (31P NMR). GPC analysis confirmed a similar molecular weight distribution for every annual plant under investigation. Arylglycerol β-arylether (β-O-4) units are the major interunit structure of lignins, followed by phenylcoumaran (β-5) and pinoresinol (β-β) units (2D-HSQC). As a consequence of the huge development of bioethanol production platforms, large amounts of lignin are produced as side stream. Uniform chemical features in lignin process streams are therefore profitable and desired properties as they could be utilized as feedstock for green chemicals, offering a significant opportunity for enhancing the operation of a lignocellulosic biorefinery, regardless of the origin of the feeding material. Moreover, the 2D-HSQC-NMR analyses also provide a qualitative assessment of the presence of pcoumarates and ferulates,39 particularly abundant in herbaceous plants and involved in lignification. In Figure 7 it is reported the aromatic region of 2D-HSQC spectrum from acetylated lignin extracted from rice husk sample as an example. NMR work on grasses47,48 established that the acylation by p-coumaric acid was exclusively at the γ-position. The DFRC (Derivatization Followed by Reductive Cleavage) method, which leaves such γesters intact, further elucidated the γ-acylation and indicated that p-coumarates were predominantly on syringyl units.47 This was recognized as the reason why both syringyl and pcoumarates units were so well represented in the HSQC spectra of all four herbaceous substrates examined. In grasses, most lignin molecules have ferulates fully incorporated into them. Ferulates radically cross-coupled during lignification analyze as lignin.Therefore, it has been suggested that ferulates are a

Table 3. GPC and NMR Data for Acidolysis Lignin (AL) Specimens Extracted from Rice Husk, Arundo donax, Wheat Straw, and Miscanthus sinensisa

GPC Mn (g/mol) Mw (g/mol) Mp (g/mol) I 2D-HSQC-NMR Side Chains Region β-O-4(A) β-5(B) β-β(C) 2D-HSQC-NMR Aromatic Region S−OH(D) S−OH, α-ketone(E) G−OH(F) ferulate(G) P−OH(H) p-coumarate(I) 31 P NMR aliphatic −OH, tot (mmol/g) aliphatic −OH, β-O-4(A) (mmol/g) cond PhOH(L) + S−OH (D) (mmol/g) G−OH(F) (mmol/g) P−OH(H) (mmol/g) COOH (mmol/g)

rice husk, AL

Arundo donax, AL

wheat straw, AL

Miscanthus sinensis, AL

10200 41000 5100 4.0

15000 81800 6400 5.5

10200 57500 4900 5.6

9600 36000 5500 3.7

+++ ++ n.d.

+++ ++ ++

+++ + +

+++ n.d. +

+++ n.d. +++ + n.d. +++

+++ n.d. +++ + ++ +++

+ + + + + +

+++ n.d. ++ n.d. ++ +++

2.89

4.35

3.42

3.47

1.27

1.74

1.31

1.69

0.21

0.32

0.29

0.27

0.61 0.66 0.22

0.61 0.53 0.15

0.67 0.43 0.29

0.58 0.77 0.17

++ + ++ ++ ++

a

2D-HSQC-NMR qualitative analyses outcomes are reported as relative abundance (the number of + marks indicates the relative amounts of the structure in the lignin sample; n.d. not detected) and differentiated in side chain and aromatic region absorptions. 31P-NMR quantitative data are expressed as mmol/g of extracted lignin. Each assigned NMR signal has its corresponding chemical structure reported in Figure 7.

Figure 7. On the left, intermonomeric (A, B, C), phenolic (D, F, H, L), and aromatic (D, E, F, G, H, I) units detected by 2D-HSQC-NMR and 31P NMR spectroscopy. On the right, aromatic region of 2D-HSQC spectrum from acetylated lignin extracted from rice husk in DMSO-d6. 452


Biomacromolecules

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natural component of lignins in grasses. 49 Their full incorporation into the polymer network could justify the lower amount of ferulates units detected by the HSQC experiment if compared to p-coumarates. Both ferulates and pcumarates, identified during lignin analysis and thoroughly discussed with the GPC chromatograms, could therefore be recognized as fundamental units in LCCs formation in herbaceous plants.

(6) Ragauskas, A. J.; Williams, C. K.; Davison, B. H.; Britovsek, G.; Cairney, J.; Eckert, C. A.; Frederick, W. J. Jr; Hallett, J. P.; Leak, D. J.; Liotta, C. L. Science 2006, 311 (5760), 484−489. (7) Arantes, V.; Saddler, J. N. Biotechnol. Biofuels 2010, 3, 4. (8) Berlin, A.; Balakshin, M.; Gilkesa, N.; Kadla, J.; Maximenko, V.; Kubo, S.; Saddler, J. J. Biotechnol. 2006, 125 (2), 198−209. (9) Balan, V.; da Costa Sousa, L.; Chundawat, S. P. S; Marshall, D.; Lansing, E.; Sharma, L. N.; Chambliss, C. K.; Dale, B. E. Biotechnol. Prog. 2009, 25 (2), 365−375. (10) Freudenberg, K.; Grion, G. Chem. Ber. 1959, 92, 1355−1363. (11) Lawoko, M.; Henriksson, G.; Gellerstedt, G. Holzforschung 2003, 57, 69−74. (12) Lawoko, M.; Henriksson, G.; Gellerstedt, G. Holzforschung 2006, 60, 156−161. (13) Lawoko, M.; Berggren, R.; Berthold, F.; Henriksson, G.; Gellerstedt, G. Holzforschung 2004, 58 (6), 603−610. (14) Lawoko, M.; Henriksson, G.; Gellerstedt, G. Holzforschung 2006, 60, 162−165. (15) Galbe, M.; Zacchi, G. Adv. Biochem. Eng. Biotechnol. 2007, 108, 41−65. (16) Lee, S. H.; Doherty, T. V.; Linhardt, R. J.; Dordick, J. S. Biotechnol. Bioeng. 2009, 102 (5), 1368−1376. (17) Tan, S. S. Y.; MacFarlane, D. R.; Upfal, J.; Edye, L. A.; Doherty, W. O. S.; Patti, A. F.; Pringle, J. M.; Scott, J. L. Green Chem. 2009, 11, 339−345. (18) Fu, D.; Mazza, G.; Tamaki, Y. J. Agric. Food Chem. 2010, 58 (5), 2915−2922. (19) Li, B.; Asikkala, J.; Filpponen, I.; Argyropoulos, D. S. Ind. Eng. Chem. Res. 2010, 49 (3), 2477−2484. (20) Swatloski, R. P.; Spear, S. K. H.; Rogers, R. D. J. Am. Chem. Soc. 2002, 124, 4974−4975. (21) Kilpelainen, I.; Xie, H.; King, A.; Granstrom, M.; Heikkene, S.; Argyropoulos, D. S. J. Agric. Food. Chem. 2007, 55, 9142−9148. (22) Holm, J.; Lassi, U. Ionic. In Ionic Liquids: Applications and Perspectives; Kokorin, A., Ed.; Intech Press: Rijeka, Croatia, 2011; p 545. (23) Swatloski, R. P.; Holbrey, J. D.; Spear, S. K.; Rogers, R. D. Molten Salts XIII. In Proceedings of the 13th International Symposium on Molten Salts; De Long, H. C., Bradshaw, R. W., Matsunaga, M., Stafford, G. R., Truelove, P. C., Eds.; The Electrochemical Society Proceeding Series: Pennington, NJ, 2002; p 155. (24) Zoia, L.; King, A. W. T.; Argyropoulos, D. S. J. Agric. Food Chem. 2011, 59 (3), 829−838. (25) Salanti, A.; Zoia, L.; Tolppa, E.-L.; Giachi, G.; Orlandi, M. Microchem. J. 2010, 95, 345−352. (26) Yeh, T. F.; Yamada, T.; Capanema, E.; Chang, H. M.; Chiang, V.; Kadla, J. F. J. Agric. Food Chem. 2005, 53, 3328−3332. (27) Holmbom, B.; Stenius, P. In Forest Products Chemistry. Stenius, P., Ed.; Tappi Press: Atlanta, GA, 2000; Book 3, p 105. (28) Granata, A.; Argyropoulos, D. S. J. Agric. Food Chem. 1995, 43, 1538−1544. (29) Argyropoulos, D. S. J. Wood Chem. Technol. 1994, 14 (1), 45− 63. (30) Yokoyama, T.; Kadla, J. F.; Chang, H. M. J. Agric. Food Chem. 2002, 50, 1040−1044. (31) Pinto, P. Ph.D. Thesis, University of Aveiro, Aveiro, Portugal, 2005. (32) King, A.; Järvi, P.; Kilpeläinen, I.; Heikkinen, S.; Argyropoulos, D. S. Biomacromolecules 2009, 10, 458−463. (33) Himmel, M. E.; Tatsumoto, K.; Oh, K. K.; Grohmann, K.; Johnson, D. K.; Chum, H. L. In Lignin - Properties and Materials; Glasser, W. G., Sarkanen, S., Eds.; American Chemical Society: Washington, DC, 1989; Vol. 397, p 82. (34) Adler, E. Wood Sci. Technol. 1977, 11, 169−218. (35) Canevali, C.; Orlandi, M.; Zoia, L.; Scotti, R.; Tolppa, E.-L.; Sipila, J.; Agnoli, F.; Morazzoni, F. Biomacromolecules 2005, 6 (3), 1592−1601.

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CONCLUSION Extensively milled lignocellulosic materials from rice husk, Arundo donax, wheat straw, and Miscanthus sinensis have been characterized by means of GPC-UV after derivatization in an ionic liquid. The use of acetyl chloride and benzoyl chloride as different derivatizing agents resulted in different instrumental responses (respectively at 280 and 240 nm), providing a clue to discern the connectivity among the various lignocellulosic components. This novel approach proved an extensive connectivity between lignin (or any other aromatic compounds) and the hemicellulosic fraction in all the analyzed specimens, in any case the amount and the type of those chemical bonds were different. Moreover, extracted lignin specimens were completely characterized revealing a similar structure for all the materials under examination. This similarity surely represents an important feature for future systematic production of biobased chemicals since the enzymatic digestion of biomass for the production of biofuels leaves a large amount of lignin as a byproduct. Abbreviations

RH, rice husk; AD, Arundo donax; WS, wheat straw; MS, Miscanthus sinensis; e-HNDI, endo-N-hydroxy-5-norbornene2,3-dicarboximide; AL, acidolysis lignin; IL, ionic liquid; [amim]Cl, 1-allyl-3-methylimidazolium chloride; GPC, gel permeation chromatography; Mn, number average molecular weight; Mw, weight average molecular weight; Mp, peak molecular weight; I, polydispersity index; HSQC, heteronuclear single-quantum coherence; cond. PhOH, condensed phenols; S−OH, syringyl phenols; G−OH, guaiacyl phenols; P−OH, pcoumaryl phenols; LCCs, lignin−carbohydrate complexes

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AUTHOR INFORMATION

Corresponding Author

*Tel.: +39 02 64482812. Fax: +39 02 64482835. E-mail: luca. [email protected].

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ACKNOWLEDGMENTS The authors thank the Cariplo 2008 for financial support and the COST action FP0901 for the Miscanthus sinensis samples.

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REFERENCES

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