Article pubs.acs.org/EF
Structural Analysis of Pyrolytic Lignins Isolated from Switchgrass Fast-Pyrolysis Oil Michael Fortin,† Megan Mohadjer Beromi,† Amy Lai,† Paul C. Tarves,† Charles A. Mullen,‡ Akwasi A. Boateng,‡ and Nathan M. West*,†,§ †
Department of Chemistry and Biochemistry, University of the Sciences, 600 South 43rd Street, Philadelphia, Pennsylvania 19104, United States ‡ USDA-ARS, Eastern Regional Research Center, 600 East Mermaid Lane, Wyndmoor, Pennsylvania 19038, United States S Supporting Information *
ABSTRACT: Structural characterization of lignin extracted from the bio-oil produced by fast pyrolysis of switchgrass (Panicum virgatum) is reported. This is important for understanding the utility of lignin as a chemical feedstock in a pyrolysis-based biorefinery scheme. Pyrolysis induces a variety of structural changes to lignin in addition to reduction in molecular weight. The guaiacol structural units remain largely intact, and some hemicellulose stays covalently linked to the lignin. However, twodimensional 1H−13C HSQC NMR analysis shows an absence of γ-methylene hydrogens from β-O-4 linkages, implying that rearrangements in the propyl linking chains have occurred. Ferulate and hydroxyl phenol esters are still present in the pyrolyzed lignin, but at lower concentrations than in unpyrolyzed switchgrass lignin.
■
secondary reactions of phenolic lignin monomers.10−12 In the solid phase, biomass undergoes cracking reactions to form lower-molecular-weight oligomers that are able to form aerosols in the inert gas atmosphere.13 Small-angle neutron scattering of bio-oil and gas-phase chromatography combined with 13C NMR spectroscopy indicate that pyrolytic lignin is composed of trimers, tetramers, and larger oligomeric units. This suggests that most of the pyrolytic lignin results directly from ejected lignin oligomers rather than lignin monomers undergoing recombination reactions, as this would be expected to produce chains with greater polydispersity.14,15 Studies of the pyrolysis of phenolic model compounds have revealed that loss of propyl side chains, ejection of the aromatic-ring methyoxy group, and homolytic cleavage of methyl groups are all common reactions possible during pyrolysis.16 Despite the possibility of propyl chain cleavage from model studies, there is evidence that during pyrolysis of lignin some side chains remain intact while others are released, resulting in condensed phenolic structures containing Caryl−Caryl bonds.17 Here we report the isolation, fractionation, and characterization of pyrolytic lignin from switchgrass. The individual separated fractions have been structurally analyzed using multidimensional NMR spectroscopy, Fourier transform infrared spectroscopy (FTIR), gas chromatography (GC), gelpermeation chromatography (GPC), and derivatization followed by reductive cleavage (DFRC)18 analysis. Comparison with lignin isolated from nonpyrolyzed switchgrass yields insight into the effect of fast pyrolysis on lignin structure. This analysis will provide more understanding of the mechanisms of formation and properties of pyrolytic lignins. Having an accurate structural model of pyrolytic lignins is critical to
INTRODUCTION Non-food-crop biomass is a vast potential source of carbonneutral fuels and chemicals.1,2 A fully functional biomass-based refinery will require methods to produce commodity chemical feedstocks, especially aromatics, from plants. A promising source material for arenes is lignin, which is abundant in plant cell walls. Lignin is an amorphous polymer consisting of substituted phenolic rings linked by propyl chains. Lignin is the second most abundant biopolymer behind cellulose.2 There are few examples of lignin degradation in nature aside from woodrotting lignicolous fungi.3 In fact, even termites appear to utilize merely the carbohydrate portion present in wood as a nutrition source; the lignin is excreted mostly intact with only minor changes made to the structural units.4 There is a great need for the development of lignin depolymerization catalysts, but in order to develop lignin conversion techniques, detailed structural information about the fine structure of different lignin sources is needed. Fast pyrolysis is a promising technique for the conversion of biomass to fuels. Fast pyrolysis involves high-temperature thermal decomposition of biomass in an inert atmosphere to rapidly convert solid biomass into viscous bio-oil, which can used to produce hydrocarbon fuels and chemicals.5,6 The net reactions of lignin during pyrolysis result only in partially degraded lignin, and this material remains in the bio-oil. This so-called pyrolytic lignin can be separated by fractionation with water; the majority of the bio-oil is water-soluble, whereas the lignin fractions are not and thus can be isolated as a solid.7−9 The pyrolytic lignin can be further separated into different fractions using organic solvent fractionation and/or chromatography. The effect of fast pyrolysis on the structure of lignin is not fully understood. The formation of pyrolytic lignin has been proposed to occur by some combination of thermal ejection of lignin oligomers into the gas phase from biomass particles and © 2015 American Chemical Society
Received: July 28, 2015 Revised: October 23, 2015 Published: October 27, 2015 8017
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels
Quantitative 31P NMR Analysis of Phosphitylated Lignins. According to the literature procedure,9,25 30 mg of dried sample was dissolved in a solvent mixture (0.50 mL) of anhydrous pyridine and deuterated chloroform (1.6:1.0 v/v) containing a relaxation agent (chromium(III) acetylacetonate) and a phosphitylation reagent (2chloro-4,4,5,5-tetramethyl-1,3,2-dioxaphospholane, ∼0.05−0.10 mL). The vial was capped and stirred for 60 min at room temperature. The reaction mixture was transferred into an NMR tube. According to an established procedure;26,27 the quantitative 31P NMR spectrum was acquired in the range of 160−120 ppm using a relaxation delay of 5 s with 1000 transients to ensure a high signal/noise ratio. Experiments were performed in duplicate. Chemical shifts and integrals were calibrated to an internal standard (phosphitylated N-hydroxynaphthalimide) that has a sharp peak at 152.2 ppm. Gel-Permeation Chromatography. Molecular weight data were determined by GPC. Samples were dissolved in THF (1:4 w/v, mg/ mL), cotton-filtered, and injected without further treatment. The lignin samples were analyzed with a Polymer Laboratories GPC-50 chromatograph (Varian, Inc.) and eluted at a flow rate of 1.2 mL/min with THF on a column set made of two OligoPore GPC columns (particle size 6 μm, 300 mm × 7.5 mm) maintained at 35 °C. A differential refractometer was used as detector. The molecular weight distribution was determined using an Integrated GPC system with Cirrus GPC software on the basis of a calibration curve pre-established with six poly(ethylene glycol)/poly(ethylene oxide) standards in the molecular weight range of 106 to 12 140.
facilitate the development of downstream conversion technologies utilizing pyrolytic lignin as a feedstock for value-added chemicals or advanced materials in a pyrolysis-based biorefinery.
■
EXPERIMENTAL SECTION
General Considerations. Bio-oil was produced from the Carthage variety of switchgrass by the published method using the USDA-ARS fluidized-bed pyrolysis process development unit (PDU);19,20 this is described in further detail in the Supporting Information. Structural analyses using the DFRC method18 and KMnO4 oxidation21 were done using published methods and are described further in the Supporting Information. Unless otherwise stated, all chemicals were purchased from Sigma-Aldrich and used as received. KMn(malonate)2(H2O)2 was synthesized according to a literature procedure.22 FTIR data were collected with a Thermo Scientific Nicolet i510 spectrometer from 1 mg of powdered lignin pressed with 100 mg of potassium bromide in absorbance mode. The 4000−400 cm−1 spectra were taken using high resolution with 64 scans. The data were obtained using OMNIC software. GC−MS analyses were done with either a Varian 3900 GC/Saturn 2100T MS or Thermo GC Ultra Trace/DSQ II MS instrument. Fractionation of Pyrolytic Lignin. By means of a modified version of the published method,8,9 bio-oil was fractionated by dissolution in water and isolation of the insoluble solids by filtration; approximately 8 g of dried pyrolytic lignin was isolated from 50 mL of bio-oil. The solid (2 g) was then extracted with methanol and filtered; removal of the solvent by rotary evaporation provided a dark solid material. This residue was washed with dichloromethane, dried, and then further washed by Soxhlet extraction with hexanes. The solid pyrolytic lignin (1.06 g, 53 wt %) was then separated into fractions 1− 4 by solvent fractionation (see Figure 1). The solid was washed with tetrahydrofuran (THF), and the insoluble portion was dried in vacuo to yield fraction 1 as a red-brown powder (328 mg, 16.4 wt %). The THF filtrate was concentrated in vacuo, and the residue was washed with methanol. The methanol filtrate was dried in vacuo to yield fraction 2 as a tan solid (70 mg, 3.5 wt %). The methanol-insoluble solid was triturated with THF, dried, and then washed with methanol once more. The methanol filtrate was dried in vacuo to yield fraction 3 as a brown solid (104 mg, 5.2 wt %). The methanol-insoluble solid was dried in vacuo to yield fraction 4 as a dark-brown powder (462 mg, 23.1 wt %). The isolated fractions were dried overnight under vacuum at 40 °C with P2O5 and stored in a desiccator. Fraction 4 could be further separated into fractions 4a−d by washing with solvents of increasing polarity. The solid (200 mg) was placed on a frit and washed sequentially with different solvents using vacuum filtration; the soluble portion was collected and dried in vacuo to yield each fraction. The soluble fractions were isolated with chloroform (fraction 4a, 1.2 mg, 0.6 wt %), ethyl acetate (fraction 4b, 50 mg, 25 wt %), and acetone (fraction 4c, 75 mg, 37.5 wt %); the insoluble solid remaining on the frit was fraction 4d (74 mg, 37 wt %). NMR Analyses. Saturated samples were dissolved in perdeuterated dimethyl sulfoxide (DMSO-d6), and two-dimensional (2D) 1H−13C heteronuclear single-quantum coherence (HSQC) correlation NMR spectra were recorded on a Bruker Advance 400 MHz spectrometer with a double-resonance broad-band probe at 60 °C. The HSQC analysis was performed with the Bruker phase-sensitive gradient-edited HSQC pulse sequence “hsqcetgpsi.2” using 1024 data points for an acquisition time of 0.11 s, a recycle delay of 1.5 s, a 1JC−H coupling constant of 145 Hz, and 256 scans. Spectra were recorded and analyzed by literature methods.23,24 Heteronuclear multiple-bond correlation (HMBC) spectra were collected using the pulse program hmbcgplpndqf. The spectra were obtained with 128 transients per increment for 256 increments into 4096 data points covering spectral widths of 5208.3 and 22 347 Hz on the 1H and 13C axes, respectively. All of the distortionless enhanced polarization transfer (DEPT) spectra were collected using the pulse program DEPT 135. The spectra were obtained using 40 000 scans.
■
RESULTS AND DISCUSSION Isolation of Pyrolytic Lignins. Pyrolytic lignin was isolated from switchgrass bio-oil using solvent fractionalization, similar to published methods.28 The bulk pyrolytic lignin was isolated by adding water to the bio-oil and collecting the insoluble solids, followed by removal of much of the ash and inorganics by dissolving the lignin in methanol and filtering away the insolubles. In this study, we performed additional solvent fractionation to further purify this pyrolytic lignin into additional fractions containing unique structural properties. This additional separation aided in the characterization and potential utility of the materials, as the characteristics of each fraction were more uniform. The sequential solvent fractionation described in the Experimental Section and displayed in Figure 1 was used to isolate fractions 1−4 and subfractions 4a− d; fractions 1 (16 wt %) and 4 (23 wt %) were the largest fractions. Elemental Analysis. Comparison of the elemental analyses of nonpyrolyzed and pyrolyzed switchgrass shows that pyrolysis results in an increase in carbon content and a decrease in oxygen content; the switchgrass used as a feedstock to produce the bio-oil from which these lignins were isolated has a composition of 49.41% C, 6.06% H, 44.33% O, and 0.51% N, providing a higher heating value (HHV) of 19.81 MJ/kg. Separation of the lignin from the bulk material provides a further increase in carbon content and decrease in oxygen content. Previous studies on pyrolyzed switchgrass examined the elemental composition of the water-insoluble “lignin” fraction and found values of 65.28% C, 6.53% H, 0.82% N, and 27.37% O, providing an HHV of 26.56 MJ/kg,7 but this material was not further fractionated and purified as was the material reported here and thus likely contained significant amounts of ash and inorganics. The fractionated and purified pyrolyzed switchgrass lignin fractions (Table 1) show a decrease in C and H contents with an increase in oxygen content versus the nonfractionated pyrolyzed lignin. These results are consistent with the removal of oxygen-poor hydrocarbons and impurities to yield a product with an 8018
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels Table 2. Molecular Weight Data and Degrees of Condensation for Pyrolytic Lignin Samples fraction
Mn
Mw
PDI
I1510/I1595b
1 2 3 4 SGLa
864 638 761 705 2070
1945 1417 1820 2059 5100
2.25 2.22 2.39 2.92 2.5
0.826 0.992 0.985 1.009 0.993
a
Ball-milled switchgrass lignin.29,30 bRatio of the peak intensities at 1510 and 1595 cm−1.
Figure 1. Solvent fractionation of the water-insoluble solids (pyrolytic lignin) from switchgrass bio-oil produced by fast pyrolysis.
elemental profile similar to that of vanillin (63.15% C, 5.30% H, 31.55% O), consistent with the cleavage of some of the propyl groups from lignin. Gel-Permeation Chromatography of Pyrolytic Lignins. The GPC results indicate that significant cleavage and molecular weight reduction occur during pyrolysis. All four fractions contain similar number-average (Mn) and weightaverage (Mw) molecular weights (Table 2), all of which are reduced by more than half versus nonpyrolyzed ball-milled switchgrass lignin.29,30 The polydispersity index (PDI) does not change dramatically following pyrolysis, suggesting uniformity in the degradation process. The polydispersities measured here are also similar to that of ball-milled poplar lignin (2.64)31 but significantly lower than that of organosolv switchgrass lignin (4.3).29,30 FTIR Analysis of Pyrolytic Lignins. The infrared spectra show similar behavior for all four fractions (Figure 2). The major peaks for most of the fractions are OH and carbonyl C− O stretches from carbohydrate moieties. The signals for C−O stretches at 1050 cm−1 in fractions 1 and 4 are attributed to the xylan backbone.32,33 The remaining fractions (2 and 3) do not appear to contain any carbohydrate. The conjugated carbonyl stretches (1680 cm−1) are from ferulate ester linkages, and these stretches diminish along with the bands associated with carbohydrate when the pyrolytic lignin is treated with refluxing 0.1 N HCl in dioxane. This suggests that the ferulate linkages
Figure 2. FTIR spectra of fractions 1−4 of the pyrolytic lignins isolated from bio-oil. The spectrum of acid-treated lignin isolated from switchgrass is shown for comparison. Region A: nonconjugated carbonyl CO stretch (1713 and 1715 cm−1) and conjugated CO stretch (1680 cm−1). Region B: intensities at 1510 and 1595 cm−1, whose ratio is used to determine the degree of cross-linking. Region C: xylan C−O stretch (1050 cm−1) and aliphatic alcohol C−O stretch (1033 cm−1).
are primarily associated with the hemicellulose−lignin interface. Fractions 2 and 3 have no signals at 1033 cm−1, indicating the absence of primary alcohol groups in the propyl chain, which are lost during pyrolysis.34,35 The presence of carbonyl stretches from 1713 to 1715 cm−1 in all four fractions results from the cleavage of β-O-4 linkages.36 The degree of cross-linking can be estimated by FTIR using the ratio of intensities at 1510 and 1595 cm−1. Others have reported that the absorbance at 1510 cm−1 is more intense for lignins with highly cross-linked structures.37−40 Lignin isolated
Table 1. Elemental Composition and Higher Heating Values of Lignin Fractions (Dry Ash-Free Basis) fraction
yield (%)
C (%)
H (%)
O (%)a
N (%)
S (%)
HHV (MJ/kg)
1 2 3 4 Py-SGLb Py-SGc SGd
16.4 3.5 5.2 23.1
62.36 62.84 63.23 63.84 65.28 53.56 49.41
5.33 5.23 5.48 5.83 6.53 6.50 6.06
31.65 31.13 30.51 29.2 27.37 39.56 44.33
0.64 0.78 0.76 0.70 0.82 0.38 0.51
0 0 0 0.40 0 0 0
23.18 23.79 24.74 23.18 26.56 20.36 19.81
a
By difference. bWater-insoluble material from switchgrass pyrolysis oil with no further fractionation.7 cSwitchgrass pyrolysis oil.7 dDried switchgrass before pyrolysis. 8019
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels
position. Similar results were obtained in Meier’s chemical degradation studies of pyrolytic lignin from woody biomass;10 the observed degradation products were due largely to the cleavage of carbohydrate structural units. The ether dimer 6 is a cross-linker between lignin and carbohydrate and is observed in grass.10 The condensed structures (e.g., structures 7−9) are not associated with native lignin. These structures are formed during pyrolysis. Structure 8 may result from a dehydration and recombination step whereby the original ether oxygen is ejected and a C−C bond is formed. Next we turned to a catalytic oxidative cleavage procedure to get further information on the structure of the isolated pyrolyic lignins. Because of the ability of manganese peroxidase (MnP) to cleave lignin utilizing carboxylate-ligated Mn(III) ions, we postulated that KMn(malonate)2(H2O)2 may catalyze oxidative cleavage of the pyrolytic lignins.42 Indeed, monolignols were observed when fraction 4 was treated with KMn(malonate)2(H2O)2 in THF. GC−MS analysis of the products clearly indicated fragments derived from guaiacol and syringyl units (10−14 and 16−18; Figure 4). The MS fragmentation
from poplar was reported to have typical absorbance values of 0.201 and 0.266, corresponding to a ratio of 0.756. Higher ratios indicate the presence of more cross-linked and condensed structures.41 This analysis (Table 2) shows that fraction 4 has the highest amount of condensed structures. Fraction 2 has a slightly more condensed structure than native switchgrass. Chemical Degradation Studies. In an effort to further determine the structural composition of the pyrolytic lignin fractions, we employed several chemical degradation and analysis methods. The materials proved to be resistant to oxidative cleavage by treatment with HCl and bleach. These methods modify the functional groups, making the majority of the material water-soluble, presumably by oxidizing alcohols to carboxylic acids, but do not produce any significant quantities of volatile organic compounds. The application of the derivatization followed by reductive cleavage (DFRC) technique to each fraction also failed to provide any ligninderived fragments.18 The method is based upon activation of benzylic ether positions with acetyl bromide followed by reductive cleavage with zinc dust. After this treatment the only volatiles found were anhydrosugars; these are monomers of cellulose formed by its depolymerization. Carbohydrate units were detected in fractions 1 and 4. After this treatment, the only volatiles found were anhydrosugar residues resulting from the carbohydrates contained in fractions 1 and 4. Oxidation with KMnO4 can be used to detect ether linkages by release of fragments bound by ether bonds regardless of a benzylic C−H site.21 This method was applied to fraction 4 and proved useful, as signals consistent with the series of monomers 1−4 and dimers 5−9 (Figure 3) were detected by GC−MS. One significant observation from the KMnO4 oxidation of fraction 4 is that there are no degradation products from β-O-4 ether type linkages; rather, the products are from ester linkages and linkages bearing carbonyl functional groups at the benzylic
Figure 4. Structures observed after catalytic oxidation of pyrolytic lignin fraction 4.
peaks of the major species are reported in Table 3. Structures 10 and 12 may have originated as ferulate ester linkages to hemicellulose. The anhydrosugar 15 is a byproduct of hemicellulose depolymerization. The assignment of fragments 10, 11, 13, and 14 originating from guaiacol units is based on the presence of fragment peaks at m/z 177 and 163. The m/z 177 value corresponds to ferulic aldehyde and has been observed in the fragmentation of hycandinic acid ester in positive-ion mode.43 Syringyl-derived units are assigned to 12 and 16 on the basis of the presence of both m/z 181 and m/z 193 or 196 fragments. Low-resolution mass spectrometry cannot distinguish between syringaldehyde
Figure 3. Structures observed after potassium permanganate oxidation of fraction 4. 8020
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels Table 3. GC−MS Results for Catalytic Oxidation of Pyrolytic Lignin Fraction 4 conditionsa
chemical species
retention time (min)
A A A A A A A A A B B B
10 11 12 13 14 15 16 17 18 19 20 21
14.978 16.978 17.731 17.859 18.552 18.969 20.566 28.245 32.567 17.928 20.790 25.437
m/z valuesb,c 234, 248, 236, 220, 262, 220, 237, 220, 306, 236, 237, 209,
219, 191, 177, BP163, 135 233, 219, 192, BP177, 163 221, 193, 181, BP165 205, BP177, 163, 149, 135 247, 220, 206, BP191, 177, 163 BP205, 177, 145, 57 196, 181, BP153 205, BP71 291, 250, 235, 219, BP189, 177, 161, 147, 133, 119 221, 208, 193, 180, BP165, 151, 137, 123, 109, 91 196, BP153, 137 195, 181, 167, 153, 138, 125, 111, BP95
Conditions A: KMn(malonate)2(H2O)2, THF, 80 °C, 14 h. Conditions B: KMn(malonate)2(H2O)2, CH3CN, 80 °C, 14 h. bGC−MS samples were derivatized with trimethylsilyl chloride (TMSCl). cBP = biggest peak. a
Figure 5. Two-dimensional 1H−13C HSQC spectra of pyrolyzed switchgrass lignin (fractions 1−4 before THF fractionation) (blue contours) and native lignin extracted from switchgrass (red contours).
and ethyl syringate, as both have nominal values of m/z 181. Fortunately, 16 contains a peak at m/z 196, which only occurs for the methyl syringyl ketone. In the case of 12, a peak at m/z 193 is observed, indicating syringyl allyl cation. Compounds 17 and 18 are proposed to be fragments from guaiacol−guaiacol and syringyl−syringyl resinol dimers. These structures are believed to be involved in cross-linking chains, but the amounts of these fragments are quite small, preventing isolation of these species for further characterization. Interestingly, when the same oxidation reaction is carried out in acetonitrile rather than THF, syringyl-based products 19−21 are the primary products
observed. One possible cause of the difference is the competing oxidation of THF to form γ-butyrolactone, among other species, which may react with lignin-based aromatics. Oxidation of THF is observed under these reaction conditions, whereas no solvent oxidations are observed to occur in the acetonitrile reactions. NMR Analysis of Pyrolytic Lignins. Pyrolytic lignin fractions were further characterized by 1H−13C HSQC (Figure 5), 1H−13C HMBC (Figure 6), and 13C DEPT (Figure 7) analyses. NMR analysis of lignocellulose materials is complicated because lignin and carbohydrate signals overlap 8021
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels
Figure 6. Two-dimensional 1H−13C HMBC spectra of pyrolyzed switchgrass lignin (fractions 1−4 before THF fractionation) (blue contours) and native lignin extracted from switchgrass (red contours).
which should appear at 4.8 ppm/71.5 ppm and 4.3 ppm/84.8 ppm, respectively, in DMSO-d6.46 These data, coupled with the lack of any product formation from DFRC, suggest that this linkage has been oxidized to a keto group at the α position as opposed to being a β-O-4 linkage. The lignin isolated from bio-oil before further purification was used in an initial NMR HSQC study (Figure 5), and ferulate esters and hemicellulose xylan could be easily assigned. The reducing-atom and anomeric cross-peak signals are of high intensity. Some Caryl−Caryl linkages can be seen in the form of stilbene and diarylmethylenes. Comparison of the HSQC spectrum of fraction 2 with that of the mixture of fractions 1−4 shows that fraction 2 contains stilbene linkages (6.73 and 115.0 ppm) and almost no β-O-4 linkages. Fraction 2 may be the result of pyrolytic oxidation and/or dehydration of a linear β-O4 fragment of lignin. Fraction 3 exhibits weak NMR crosspeaks, making the assignments difficult; the HSQC spectrum was featureless except for the methoxy (3.73 and 56.5 ppm) and guaiacyl C5 (6.73 and 115.2 ppm) linkage cross-peaks. Fraction 4 is a complex mixture of at least three types of oligomers that can be further fractionated with different solvents. Solvation in the following solvents in the order listed provided the following yields of subfractions: chloroform, 0.6 wt % (fraction 4a); ethyl acetate, 25 wt % (fraction 4b); acetone, 37.5 wt % (fraction 4c). The remaining 37 wt % is fraction 4d. The chloroform-soluble portion (fraction 4a) contains mostly guaiacyl units followed by syringyl units, and the 13C signal for nonconjugated carboxylic acids attached to guaiacyl units appears at 178 ppm. Ferulate ester linkages are also indicated by the C6 peak at 123 ppm. Syringyl groups bearing carboxyl groups on the α carbon lead to a peak at 130 ppm. Presumably they are connected to the chain by ether linkages at C4, as a syringyl monomer cannot cross-link at the C5 position (a methoxy group is attached). Resonances due to syringyl benzoic acid moieties are observed only in the pyrolyzed lignin fractions, as indicated by HMBC cross-peaks at 6.87/148.4 ppm (4c) and 6.76/144.6 ppm (4b) (Figure 6). Lignin isolated from switchgrass contains aldehydes, guaiacyl,
in the ether region (∼3−5 ppm). Assignments were made on the basis of literature sources.35,44,45 Upon examination of the HSQC spectrum (Figure 5) it can be seen that many aryl methoxy groups are still present in the lignin materials post hydrolysis. The presence of guaiacyl units is indicated by comparison of the proton and carbon chemical shifts with those of extracted nonpyrolyzed switchgrass lignin. Evidence of the existence of xylose and arabinose units implies that the hemicellulose xylan backbone is largely intact and possibly is connected to lignin components through ferulate ester linkages. Fraction 1 bears a galactose unit derived from hemicellulose. The sugar ring is connected to the lignin polymer by two types of linkages: a phenyl glycoside linkage (102 ppm, from C1 of the carbohydrate to the C4 ether of a guaiacyl unit) and an ester linkage (65 ppm, from the C6 position of a galactose ring to the propane chain with a keto carbon at the γ position). Fraction 1 is rich in syringyl units, as evidenced by the signals for the carbons bearing the methoxy groups (C1, 125 ppm; C3; C5, 151 ppm). Guaiacyl units involved in ether linkages (C5, 119 ppm) as well as guaiacyl units containing phenolic positions (C4, 145 ppm) are observed in the HMBC spectrum (Figure 6). The thermal degradation results in the cleavage of the phenyl propane bonds. Ferulate esters are also present at 170 ppm. Importantly, in the post pyrolysis lignin fractions, the aryl groups do not appear to be connected by β-O-4 linkages. If any β-O-4 linkages exist in the pyrolyzed material, their amounts are too small to detect. The majority of the connections are due to propane chains with keto groups at the benzyl α position along the alkyl chain (191 ppm), indicating that oxidation of lignin benzyl alcohol positions occurs during pyrolysis. Fraction 2 is a more purely lignin-derived material with no detectable amount of carbohydrates. This makes assignments of the NMR spectra easier. In addition to the presence of guaiacyl and conjugated ester aromatic units the presence of CγHγ moieties (3.87 ppm/64.7 and 3.50 ppm/64.7 ppm) from β-O-4 type linkages can be detected. However, there are no corresponding methine signals for α and β of a β-O-4 linkage 8022
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels
Figure 7. Comparison of 13C DEPT 135 spectra for ethyl acetate-soluble fraction 4b (spectrum 2, top) and extracted and acid-treated switchgrass lignin (spectrum 1, bottom).
and syringyl units as determined by NMR analysis.47,48 The HMBC spectrum of ethyl acetate-soluble fraction 4b shows the same type of galactose unit as in fraction 1, as determined by both fractions having an anomeric carbon at 102 ppm. In addition, fraction 1 contains an additional anomeric methine at 4.43 ppm (1H) and 103.38 ppm (13C) that is not shared by fraction 4b. Again, two types of sugar lignin linkages could be identified: phenyl glycoside and ester linkages with the keto group on the lignin propane carbon. Very few syringyl units could be detected; the oligomer is mostly guaiacyl units, some involved in ether linkages (guaiacyl Cipso at 137 ppm) and some in ester linkages (carbonyl carbon at 170 ppm). The ferulate ester linkages are connected as dimers, as has been reported for grasses. Only fraction 4b contains nonconjugated ferulates, as indicated by a 13C signal at 173 ppm (ester carbonyl carbon), where the C−C double bond has been reduced.49 The linkages again are mostly C−C bonds with some ethers; no α- or β-O-4 methines could be detected in the DEPT 135 spectra (Figure 7). Fraction 4c is unique in that only trace amounts of galactose could be detected in the DEPT 135 spectrum. It is the most diverse portion of fraction 4 and contains syringyl phenolic (C3 and C5, 152 ppm), guaiacyl ether, and phenolic linkages as well as ferulate ester linkages. This is the most pure lignin material of all the pyrolytic lignin fractions. For comparison, lignin was isolated from nonpyrolyzed switchgrass, and this material contains a significant amount of attached carbohydrate that can be removed by refluxing in 9:1 0.1 N HCl/dioxane for 1 h. The 1H−13C HMBC spectrum shows that β-O-4 linkages, ferulate esters, and guaiacyl ether linkages all remain, indicating a significant difference from the pyrolyzed material, which is devoid of β-O-4 linkages. The solubility of this material is different from that of the isolated lignin, as it is soluble in methanol, water, and DMSO. Before acid treatment, the lignin isolated from switchgrass is soluble in pyridine, THF, and acetone. As opposed to the pyrolyzed lignin, no fractionation can be obtained by subjecting the nonpyrolytic lignin to different solvents. This implies more ether linkages and more homogeneity among the chains compared with the pyrolytic lignin. The NMR assignments for the carbohydrate portions of the fractions are tabulated in the Supporting Information; these signals are consistent with xylan as the predominant
carbohydrate. Residues associated with glucomanan are not observed; they may have not survived the pyrolysis. The assignment of ether linkages for lignin is difficult because of the overlapping of xylan ring atom cross-peaks. Cross-peaks for guaiacyl and syringyl aromatic rings are easy to detect (Table 4). Ferulate esters are associated with fractions 1 and 4. These Table 4. Assignment of 13C−1H Correlation Signals in the HSQC Spectrum Observed for Lignin Functional Groups chemical shift δC/δH (ppm) 20.7/2.1 53.0/3.4 53.5/3.07 55.6/3.7 62.5/3.7 69.7/3.5 89.5/5.0 104.1/6.5 107.1/7.1 112.7/6.9 115.5/5.8 115/5.6 119.1/6.6 123.2/7.4 126.5/7.3 129.4/7.3 129.4/6.8 129.4/5.2 132.1/7.7 154.1/7.9
assignment biaryl methylene Cβ-Hβ phenylcoumarin Cβ-Hβ resinol OMe (Cγ-Hγ) phenylcoumarin (Cγ-Hγ) resinol (Cα-Hα) phenylcoumarin (C2,6/H2,6) syringyl Cα/β−Hα/β stilbene CO syringyl (C2/H2) CO guaiacyl (Cβ-Hβ) p-coumaryl ester (C5/H5) in guaiacyl CO C6/H6 in guaiacyl phenolic ether-linked ferulate ester (Cα/β−Hα/β) stilbene (C2/H2) p-OH phenyl (cinnamyl alcohol) (C6/H6) p-OH phenyl (cinnamyl alcohol) (C2,6/H2,6) coniferyl alcohol (allylic alcohol) (C2,6/H2,6) p-hydroxybenzoate (phenolic OH free) (Cα-Hα) ferulate ester
fractions also contain stilbene linkages and guaiacyl and syringyl linkages bearing a carbonyl (CO) at the benzylic position. This can account for the low reactivity with the acetyl bromide reagent for DFRC analysis. Phenylcoumarin and resinol linkages are present in both pyrolyzed and native lignin, indicating the stability of these linkages toward pyrolysis. 31 P NMR Analysis of Pyrolytic Lignins. NMR analysis of alcohol types was performed by reacting the lignin samples with 2-chloro-4,4,5,5-tetramethyl-1,3,2-dioxaphospholane using pre8023
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels
Figure 8. 31P NMR spectra of phosphitylated lignin fractions. On the basis of a comparison to those of known model compounds, the spectra can be divided into seven regions: (1) aliphatic alcohols (including carbohydrate); (2) diphenylmethane; (3) 5−5′ diphenyl; (4) syringol; (5) guaiacol; (6) p-OH phenol; (7) carboxylic acid.51
Table 5. Quantification of Lignin Alcohol Groups by Phosphitylationa lignin 1 2 3 4 4b 4c 1−4 avgc SGLd
aliphatic 6.53 1.34 1.43 3.81 0.85 0.62 4.30 2.99
± ± ± ± ± ± ± ±
0.59 0.58 0.20 2.28 0.01 0.01 1.36 0.10
DPMb 0.51 0.27 0.49 0.52 0.27 0.32 0.50 0.04
± ± ± ± ± ± ± ±
0.07 0.13 0.06 0.03 0.03 0.01 0.05 0.02
5−5′ 1.23 0.50 0.82 1.11 0.74 0.45 1.08 0.32
± ± ± ± ± ± ± ±
syringyl
0.01 0.21 0.11 0.65 0.02 0.01 0.34 0.04
0.45 0.25 0.39 0.38 0.25 0.19 0.40 0.09
± ± ± ± ± ± ± ±
0.04 0.05 0.05 0.24 0.01 0.01 0.14 0.02
guaiacyl 3.44 1.18 1.89 2.54 1.68 0.84 2.68 0.64
± ± ± ± ± ± ± ±
0.33 0.51 0.23 1.55 0.04 0.01 0.92 0.08
p-OH phenol 1.78 0.65 1.00 1.36 1.05 0.60 1.41 0.28
± ± ± ± ± ± ± ±
0.15 0.26 0.12 1.07 0.13 0.01 0.60 0.03
carboxyl 0.59 0.13 0.27 0.12 0.14 0.02 0.30 0.15
± ± ± ± ± ± ± ±
0.68 0.10 0.02 0.57 0.02 0.01 0.51 0.04
a
All values are in units of mmol/g of lignin. bDPM = diphenylmethane. cWeighted average based on the yields of fractions 1−4. dExtracted switchgrass lignin treated with 1 N HCl.
viously reported methods.9,50 Three types of alcohols are detected by the analysis: aliphatic, aromatic, and aromatic condensed (Figure 8). The guaiacyl OH peak at 140−138 ppm is present in all three lignin oligomer types. The syringyl OH peak is shifted downfield to 144 ppm compared with that of nonpyrolyzed wheat lignin (142 ppm). The carboxylic acid content is low for fractions 2−4. Only fraction 1 has more carbohydrate than nonpyrolyzed switchgrass. Compared with milled wood lignin, the pyrolytic lignin has a poorly defined βO-4 guaiacol region. All of the pyrolyzed fractions have significantly more aryl OH groups than the extracted switchgrass lignin, except for 4b and 4c (Table 5). The fractions all show a large increase in condensed OH groups as well, indicating that the formation of these structures is favored by pyrolysis. The number of aliphatic groups is significantly higher in fraction 1, most likely because of the presence of a significant amount of carbohydrate. Fractions 2 and 3, which do not appear to contain carbohydrate, both show a decrease in the amount of aliphatic groups, consistent with the destruction of β-O-4 linkages. The overall types of alcohols present remain fairly constant through the fractions. Fractions 1 and 4, which constitute the majority of the material, have similar S:G:H alcohol ratios as the
extracted lignins, although there are some differences in the minor fractions. Development of a Structural Model. The data reported here can be combined to create a structural model of a typical pyrolytic lignin oligomer (Figure 9). A typical oligomer has a mass of 1000 Da and contains a varying degree of xylan-derived carbohydrate covalently attached through ferulate linkages. Xylan is the most abundant component of hemicellulose and is composed of xylose, arabinose, and glucuronic acid. The failure of DFRC and KMnO4 to produce any cleaved ether products implies the absence of any hydroxyl groups at the α and β positions of the propyl chains. The absence of any apparent βO-4-type linkages indicates that these are particularly prone to cleavage in fast pyrolysis. The fast-pyrolysis process leaves mainly aryl carbonyl fragments and condensed aliphatic and phenolic structures. The aromatic syringyl and guaiacol units are left intact along with the xylan and ferulate linkages. Some of the observed linkages are pine resinol, phenylcoumarin, and stilbene, which can be produced by the degradation of β-1 linkages. Pyrolysis leads to an increase in the concentration of phenolic positions, but the S:G:H ratio is not substantially changed by pyrolysis. 8024
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels
Figure 9. Proposed structural model of pyrolytic switchgrass lignin.
■
Present Address
CONCLUSION
§
N.M.W.: Eastman Chemical Company, 200 S. Wilcox Dr. Kingsport, TN 37660.
The analysis of pyrolytic lignins shows that the pyrolysis produces oligomers with differing amounts of hemicellulose present, which affects the solubility of the fractions and their utility for value-added product synthesis. The pyrolysis leads to a highly condensed phenolic structure that is not well modeled by traditional lignin analogues and will require different conversion technologies than extracted or biorefinery lignins. Solvent fractionation of the water-insoluble pyrolytic lignin from bio-oil leads to the isolation of four main fractions of increased purity. This aids in structural analysis and helps to enable high-value product synthesis by providing a more uniform and reproducible starting material. Having betterresolved structural features, such as the amount of carbohydrate and the types of lignin linkages present, is very important for future uses like catalytic cleavage of the lignin into its monomers or incorporation of lignin into advanced materials.52
■
Funding
We acknowledge funding support from U.S. Department of Agriculture Biomass Research and Development Initiative (BRDI) grant (USDA-NIFA 2012-10008-20271). Notes
Disclaimer: Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. The authors declare no competing financial interest.
■ ■
ACKNOWLEDGMENTS We acknowledge the technical assistance of Dr. Robert Y. Nsimba.
ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.energyfuels.5b01726. Experimental details for bio-oil production and fractionation, native lignin extraction, DFRC analysis, and lignin oxidation analysis; FTIR and NMR spectra and GC−MS chromatograms; tabulation of IR assignments and NMR chemical shifts of carbohydrate fragments (PDF)
■
■
ABBREVIATIONS DFRC = derivatization followed by reductive cleavage HSQC = heteronuclear single-quantum coherence HPLC = high-pressure liquid chromatography THF = terahydrofuran GPC = gel-permeation chromatography TEMPO = 2,2,6,6-tetramethylpiperidin-1-oxyl REFERENCES
(1) Mullen, C. A.; Boateng, A. A. Energy Fuels 2008, 22, 2104. (2) Zakzeski, J.; Bruijnincx, P. C. A.; Jongerius, A. L.; Weckhuysen, B. M. Chem. Rev. 2010, 110, 3552. (3) Ruiz-Duenas, F. J.; Martinez, A. T. Microb. Biotechnol. 2009, 2, 164. (4) Ke, J.; Laskar, D. D.; Singh, D.; Chen, S. Biotechnol. Biofuels 2011, 4, 17. (5) Czernik, S.; Bridgwater, A. V. Energy Fuels 2004, 18, 590. (6) Jahirul, M. I.; Rasul, M. G.; Chowdhury, A. A.; Ashwath, N. Energies 2012, 5, 4952.
AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. 8025
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
Article
Energy & Fuels (7) Mullen, C. A.; Boateng, A. A. J. Anal. Appl. Pyrolysis 2011, 90, 197. (8) Nsimba, R. Y.; West, N.; Boateng, A. A. J. Agric. Food Chem. 2012, 60, 12525. (9) Nsimba, R. Y.; Mullen, C. A.; West, N. M.; Boateng, A. A. ACS Sustainable Chem. Eng. 2013, 1, 260. (10) Bayerbach, R.; Meier, D. J. Anal. Appl. Pyrolysis 2009, 85, 98. (11) Bai, X.; Kim, K. H.; Brown, R. C.; Dalluge, E.; Hutchinson, C.; Lee, Y. J.; Dalluge, D. Fuel 2014, 128, 170. (12) Zhou, S.; Garcia-Perez, M.; Pecha, B.; Kersten, S. R. A.; McDonald, A. G.; Westerhof, R. J. M. Energy Fuels 2013, 27, 5867. (13) Piskorz, J.; Maherski, P.; Radlein, D. In Biomass, A Growth Opportunity in Green Energy and Value-Added Products: Proceedings of the 4th Biomass Conference of the Americas; Overend, R. P., Chornet, E., Eds.; Elsevier Science: Amsterdam, 1999; p 1153. (14) Fratini, E.; Bonini, M.; Oasmaa, A.; Solantausta, Y.; Teixeira, J.; Baglioni, P. Langmuir 2006, 22, 306. (15) Scholze, B.; Hanser, C.; Meier, D. J. Anal. Appl. Pyrolysis 2001, 58−59, 387. (16) Hao, C.; Wu, S.; Liu, C. BioResources 2014, 9, 4441. (17) Meier, D.; Jesussek, G.; Radtke, S. In Pyrolysis and Gasification of Biomass and Waste; Bridgewater, A. V., Ed.; CPL Press: Newbury, U.K., 2003; p 221. (18) Lu, F.; Ralph, J. J. Agric. Food Chem. 1997, 45, 2590. (19) Boateng, A. A.; Daugaard, D. E.; Goldberg, N. M.; Hicks, K. B. Ind. Eng. Chem. Res. 2007, 46, 1891. (20) Boateng, A. A.; Mullen, C. A.; Goldberg, N.; Hicks, K. B.; Jung, H.-J. G.; Lamb, J. F. S. Ind. Eng. Chem. Res. 2008, 47, 4115. (21) Gellerstedt, G. In Methods in Lignin Chemistry; Lin, S. Y., Dence, C. W., Eds.; Springer: Berlin, 1992; p 322. (22) Cartledge, G. H.; Nichols, P. M. J. Am. Chem. Soc. 1940, 62, 3057. (23) del Río, J. C.; Rencoret, J.; Prinsen, P.; Martínez, Á . T.; Ralph, J.; Gutiérrez, A. J. Agric. Food Chem. 2012, 60, 5922. (24) Ralph, J.; Landucci, L. L. In Lignin and Lignans: Advances in Chemistry; Heitner, C., Dimmel, D., Schmidt, J., Eds.; CRC Press: Boca Raton, FL, 2010; p 137. (25) Pu, Y.; Cao, S.; Ragauskas, A. J. Energy Environ. Sci. 2011, 4, 3154. (26) Scholze, B.; Meier, D. J. Anal. Appl. Pyrolysis 2001, 60, 41. (27) Granata, A.; Argyropoulos, D. S. J. Agric. Food Chem. 1995, 43, 1538. (28) Sukhbaatar, B.; Steele, P. H.; Kim, M. G. BioResources 2009, 4, 789. (29) Hu, G.; Cateto, C.; Pu, Y.; Samuel, R.; Ragauskas, A. J. Energy Fuels 2012, 26, 740. (30) Tolbert, A.; Akinosho, H.; Khunsupat, R.; Naskar, A. K.; Ragauskas, A. J. Biofuels, Bioprod. Biorefin. 2014, 8, 836. (31) Kim, J. Y.; Shin, E. J.; Eom, I. Y.; Won, K.; Kim, Y. H.; Choi, D.; Choi, I. G.; Choi, J. W. Bioresour. Technol. 2011, 102, 9020. (32) Karatzos, S.; Edye, L.; Doherty, W. O. Biotechnol. Biofuels 2012, 5, 62. (33) Faix, O.; Bremer, J.; Schmidt, O.; Tatjana, S. J. J. Anal. Appl. Pyrolysis 1991, 21, 147. (34) Wang, S.; Lin, H.; Ru, B.; Sun, W.; Wang, Y.; Luo, Z. J. Anal. Appl. Pyrolysis 2014, 108, 78. (35) Trajano, H.; Engle, N.; Foston, M.; Ragauskas, A.; Tschaplinski, T.; Wyman, C. Biotechnol. Biofuels 2013, 6, 110. (36) Zhou, S.; Garcia-Perez, M.; Pecha, B.; McDonald, A. G.; Kersten, S. R. A.; Westerhof, R. J. M. Energy Fuels 2013, 27, 1428. (37) Sarkanen, K. V.; Chang, H. M.; Ericsson, B. Tappi 1967, 50, 572. (38) Hergert, H. L. J. Org. Chem. 1960, 25, 405. (39) Stewart, D.; Yahiaoui, N.; McDougall, G. J.; Myton, K.; Marque, C.; Boudet, A. M.; Haigh, J. Planta 1997, 201, 311. (40) Hargert, H. L. In Lignins: Occurrence, Formation, Structure and Reactions; Sarkanen, K. V., Ludwig, C. H., Eds.; Wiley: New York, 1970; p 267.
(41) Zhong, R.; Morrison, W. H., III; Himmelsbach, D. S.; Poole, F. L., II; Ye, Z. H. Plant Physiol. 2000, 124, 563. (42) Wariishi, H.; Valli, K.; Gold, M. H. J. Biol. Chem. 1992, 267, 23688. (43) Wu, Z. J.; Ma, X. L.; Fang, D. M.; Qi, H. Y.; Ren, W. J.; Zhang, G. L. Eur. Mass Spectrom. 2009, 15, 415. (44) Wen, J. L.; Xue, B. L.; Xu, F.; Sun, R. C. BioEnergy Res. 2012, 5, 886. (45) Samuel, R.; Foston, M.; Jiang, N.; Allison, L.; Ragauskas, A. J. Polym. Degrad. Stab. 2011, 96, 2002. (46) Samuel, R.; Pu, Y.; Raman, B.; Ragauskas, A. J. Appl. Biochem. Biotechnol. 2010, 162, 62. (47) Yelle, D. J.; Wei, D.; Ralph, J.; Hammel, K. E. Environ. Microbiol. 2011, 13, 1091. (48) Kim, H.; Ralph, J.; Lu, F.; Ralph, S. A.; Boudet, A.-M.; MacKay, J. J.; Sederoff, R. R.; Ito, T.; Kawai, S.; Ohashi, H.; Higuchi, T. Org. Biomol. Chem. 2003, 1, 268. (49) Li, M.-F.; Fan, Y.-M.; Sun, R.-C.; Xu, F. BioResources 2010, 5, 1762. (50) Granata, A.; Argyropoulos, D. S. J. Agric. Food Chem. 1995, 43, 1538. (51) Ahvazi, B.; Wojciechowicz, O.; Ton-That, T.-M.; Hawari, J. J. Agric. Food Chem. 2011, 59, 10505. (52) Deuss, P. J.; Barta, K. Coord. Chem. Rev. 2015, DOI: 10.1016/ j.ccr.2015.02.004.
8026
DOI: 10.1021/acs.energyfuels.5b01726 Energy Fuels 2015, 29, 8017−8026
