Evidence for Complex Molecular Architectures for Solvent-Extracted

Apr 17, 2012 - Sabornie ChatterjeeTomonori SaitoOrlando RiosAlexander Johs. 2014 .... Recent advances in low-cost carbon fiber manufacture from lignin...
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Evidence for Complex Molecular Architectures for Solvent-Extracted Lignins Shane E. Harton,*,† Sai Venkatesh Pingali,*,‡ Grady A. Nunnery,† Darren A. Baker,§ S. Hunter Walker,∥ David C. Muddiman,∥ Tadanori Koga,⊥ Timothy G. Rials,§ Volker S. Urban,‡ and Paul Langan‡ †

Materials Science and Technology Division and ‡Biology and Soft Matter Division, Oak Ridge National Laboratory, Oak Ridge, Tennessee, 37831, United States § Center for Renewable Carbon, The University of Tennessee, Knoxville, Tennessee, 37996, United States ∥ Department of Chemistry, North Carolina State University, Raleigh, North Carolina 27695, United States ⊥ Chemical and Molecular Engineering Program, Department of Materials Science and Engineering, Stony Brook University, Stony Brook, New York, 11794, United States S Supporting Information *

ABSTRACT: Lignin, an abundant, naturally occurring biopolymer, is often considered “waste” and used as a simple fuel source in the paper-making process. However, lignin has emerged as a promising renewable resource for engineering materials, such as carbon fibers. Unfortunately, the molecular architecture of lignin (in vivo and extracted) is still elusive, with numerous conflicting reports in the literature, and knowledge of this structure is extremely important, not only for materials technologies, but also for production of biofuels such as cellulosic ethanol due to biomass recalcitrance. As such, the molecular structures of solvent-extracted (sulfur-free) lignins, which have been modified using various acyl chlorides, have been probed using small-angle X-ray (SAXS) and neutron (SANS) scattering in tetrahydrofuran (THF) solution along with hydrodynamic characterization using dilute solution viscometry and gel permeation chromatography (GPC) in THF. Mass spectrometry shows an absolute molecular weight ≈18−30 kDa (≈80−140 monomers), while GPC shows a relative molecular weight ∼3 kDa. A linear styrene oligomer (2.5 kDa) was also analyzed in THF using SANS. Results clearly show that lignin molecular architectures are somewhat rigid and complex, ranging from nanogels to hyperbranched macromolecules, not linear oligomers or physical assemblies of oligomers, which is consistent with previously proposed delignification (extraction) mechanisms. Future characterization using the methods discussed here can be used to guide extraction processes as well as genetic engineering technologies to convert lignin into value added materials with the potential for high positive impact on global sustainability.

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Lignin, a completely amorphous biopolymer, can be processed in an analogous manner as many synthetic linear and branched polymers, but it lacks the strength of conventional synthetic polymers such as polyethylene, polystyrene, and poly(methyl methacrylate). This characteristic limits its use as a neat thermoplastic. The high phenolic content, on the other hand, makes lignin a useful component in thermosets, such as phenol-formaldehyde resins, polyurethanes, and epoxies.5 It has also been examined for the manufacture of low-cost carbon fibers due to the high carbon yield upon pyrolysis, but the low tensile strengths encountered with current lignin feedstocks have prevented its use in engineering applications such as automotive and aerospace technologies.6 Despite these challenges there is great interest in developing

ith the ever increasing demand for petroleum, and the inevitable disruptions in supply, material feedstocks from clean renewable resources are becoming increasingly necessary for global sustainability.1 Therefore, there is continued interest in extracting energy (e.g., cellulosic ethanol, butanol, etc.) and engineering materials (e.g., carbon fibers) from renewable resources.2 Conversion of biomass to biofuels is an attractive means of providing future energy needs; however, conversion of biomass into technologically and commercially relevant materials, thereby bypassing the need for a petroleum-based feedstock, is another promising path to sustainability. One such material is lignin, a naturally occurring phenolic resin.3,4 Although it is one of the most abundant naturally occurring macromolecules, lignin is often burned as a fuel source in the paper-making process.5 As such, its value as a material is often determined relative to its value as a simple fuel source and measured against the need to recover process chemicals. © 2012 American Chemical Society

Received: January 23, 2012 Accepted: April 11, 2012 Published: April 17, 2012 568

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though the chemical and physical properties of lignin have been exhaustively investigated for over half a century,3−5,14 there is still little known about the molecular architectures of lignins. It has been postulated,10,14 and later supported,11,13 that lignin first forms as a low molecular weight oligomer and then a network is formed from these oligomers within the plant cell wall. The somewhat rigid structure of lignin-containing plant species is dependent on this strong network.8 However, there is still conflicting speculation as to the molecular structures and potential hierarchies of extracted lignins,15 whether linear, branched, hyperbranched, cross-linked, or physical assemblies. Such a fundamental understanding, with further simulations and feedback16 to guide choice of biomass feedstock, extraction processes, and genetic engineering technologies,9 is crucial in moving this potentially valuable resource beyond current materials capabilities. Furthermore, knowledge of the molecular structures of protolignins as well as extracted lignins is extremely important for the production of cellulosic ethanol due to the complex binding interactions between hydrolyzing enzymes and lignins (recalcitrance), thereby directly coupling biofuels and materials technologies.2 Using small angle neutron (SANS) and X-ray (SAXS) scattering, as well as hydrodynamic characterization and mass spectrometry, we have probed the molecular architectures of solvent-extracted (sulfur-free) lignins in dilute tetrahydrofuran (THF) solution with a comparison to a model linear oligomer (oligostyrene). Results show that extracted lignins are rigid, compact, and polydisperse, with complex characteristics ranging from nanogels to hyperbranched macromolecules. Lignin (sulfur free), which was provided by Lignol Innovations, was extracted from a mixture of hardwoods using the so-called Alcell process, which typically involves water, ethanol, and in situ formed acetic acid at temperatures ∼195 °C and pressures ∼3 MPa.17 The extraction process used for this lignin is much milder than the kraft and sulfite pulping processes, resulting in lignins with lower molecular weights (less secondary condensation reactions) and greater solubility in organic solvents such as THF and dioxane.5 The lignin was purified, filtered (0.45 μm pores), and partially fractionated to remove organic and inorganic contaminants as well as very low molecular weight fractions. This treatment resulted in a clear increase in the glass transition temperature from ∼110 to ∼135 °C, although UV−vis spectroscopy in THF as well as Fourier transform infrared spectroscopy (FTIR) showed immeasurable changes (see Supporting Information) in the lignin chemistry before and after treatment. Therefore, the changes in the glass transition temperatures most likely resulted from removal of very low molecular weight fractions as well as residual plasticizing solvents (e.g., furfural and ethanol). Equilibrium water absorption was determined to be ∼2−3 wt % at 100% relative humidity, and physisorption of water shows a desorption endotherm with a peak ∼50 °C, confirming that this is a relatively hydrophobic material. Supporting Information contains more information regarding the purification, modification, and spectroscopic analysis. The regenerated lignin was dried and stored under N2 until it was modified with (1) acetyl chloride (AcL), (2) bromoacetyl chloride (BrAcL), (3) chloroacetyl chloride (ClAcL), (4) 4bromobenzoyl chloride (BrBenL), or (5) 4-chlorobenzoyl chloride (ClBenL) according to previously established procedures,18 which resulted in ester (acetate or benzoate) formation with the aromatic and aliphatic hydroxyls. Proton nuclear magnetic resonance spectroscopy (NMR) was

methods to convert lignin into value added materials because of the potentially high positive impact on global sustainability.1,2 For example, to reduce automotive carbon footprints, carbon fiber produced from lignin can potentially be used to reduce vehicle weight, while cellulosic ethanol can be produced from the same feedstock and used to fuel the combustion engines with greatly improved fuel efficiency due to the weight reductions.6 Most lignins are synthesized starting within a polysaccharide gel in the cell corners and continue within the secondary cell wall of plants as a covalently bound complex with hemicellulose that encrusts crystalline bundles of cellulose fibers.7,8 Nondestructive methods (e.g., NMR) for studying lignifications in plant cell walls suggest that it is highly heterogeneous in both its structure and its relative composition of three monomeric components, with significant differences in chemical compositions and structures from lignins formed in the cell corners (higher H concentrations) to lignins formed in the secondary cell walls.7,8 Figure 1 shows the three basic monomeric

Figure 1. Three primary monolignols, (H) p-coumaryl alcohol, (G) coniferyl alcohol, and (S) sinapyl alcohol. G and S tend to be the dominant species, with ratios depending on several factors, including the plant source (e.g., S/G ∼ 2:1 for the lignins used here).

constituents (monolignols), namely, p-coumaryl (H), coniferyl (G), and sinapyl (S) alcohols, with relative compositions depending on the plant source, that polymerize primarily via a complex radical process.9 Hardwood lignins typically have S/G ratios ∼2:1, while softwood lignins tend to have S/G ratios ∼1:2 to 1:3, and lignins from grasses have S/G ∼ 1:1 to 1:2. Lignins from softwoods and grasses also tend to have higher concentrations of H monolignols, although the chemical structures of lignins are strongly dependent on plant or tree species and age, climate, geographical region, location within the plant cell, and genetic modification.7−9 For example, a transgenic poplar has been reported to have S/G > 35:1.9 Numerous processes have been utilized to extract and isolate lignins, including kraft, sulfite, soda, solvent-extraction (sulfurfree), and mechanical pulping.5 The commonly accepted process of delignification (extraction) involves degelation or gel degradation.10 Delignification theories postulate that the lignin network, or gel, within the cell wall is extracted by reversing a gelation process, primarily proceeding with bonds being broken (ether linkages), and secondarily with some condensation reactions taking place (e.g., forming diaryl ethers),11,12 plus incorporation of sulfonates (-SO3−) or thiols (-SH) into the lignins for sulfite or kraft pulping processes, respectively.5 This is an extremely complex process resulting in high polydispersities and an increase in extracted lignin molecular weights as extraction time progresses.13 Each lignin extraction process, as well as the biomass source (e.g., hardwood, softwood, switchgrass, etc.), generates isolated lignins with varying chemical and physical properties. Even 569

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performed on AcL, and results suggest a quantitative reaction (ca. 1.1−1.2 acetyl groups per C9).19 It was also determined using FTIR that the acyl chloride reaction strongly favors formation of esters with the aliphatic hydroxyls in the early stages, but that the reaction inevitably goes to completion with all hydroxyls, aliphatic and aromatic. This is most likely due to accessibility of the aliphatic hydroxyls, as the aromatic hydroxyls can be hindered by adjacent methoxy groups. FTIR and UV−vis spectroscopy were performed on all samples (see Supporting Information). FTIR spectra of BrAcl, ClAcL, BrBenL, and ClBenL, when compared to the AcL spectrum, confirm quantitative reactions for all materials. NMR and FTIR revealed an aromatic/aliphatic hydroxyl ratio ∼3:1, which is consistent with previously reported results for this lignin, and the S/G ratio has been reported previously to be ∼2:1 (see Figure 1).19 Analysis of AcL using gel permeation chromatography (GPC) with THF eluent and calibration using polystyrene standards showed a relative Mw ∼ 3 kDa and Mw/Mn ∼ 3, and reverse phase liquid chromatography coupled online with electrospray ionization (ESI) mass spectrometry20 showed an absolute Mw ≈ 18−30 kDa (see Supporting Information), which corresponds to ≈80−140 monomers. The GPC-determined molecular weights are consistent with previously reported values for this particular solvent-extracted lignin.21−23 An oligostyrene (OS) reference (Mw = 2460 Da, Mw/Mn = 1.01) was purchased from Scientific Polymer Products and dried under vacuum for 4 days at room temperature. To probe the hydrodynamic properties and predict the overlap concentration, dilute solution viscometry of AcL was performed using a Cannon-Fenske viscometer. A concentration range of 3−8 vol % AcL in THF, a known good solvent for AcL,21−24 was used, resulting in an intrinsic viscosity [η] = 5.2 mL/g and a hydrodynamic volume (Vh) 2.8 times that of an Einstein (hard) sphere. Results provided a range of overlap concentrations from 150 mg/mL, which is the lower limit assuming linear flexible chains (0.77/[η]), to 490 mg/mL, which is an upper limit assuming dense spheres (2.5/[η]).25,26 The halogen-containing lignins (BrAcL, ClAcL, BrBenL, and ClBenL) were analyzed using small-angle X-ray scattering (SAXS) at beamline X10A at the National Synchrotron Light Source (NSLS) at Brookhaven National Laboratory (BNL) with a photon wavelength of 1.095 Å. The halogen-containing moieties are essential for this analysis, as they add electron density contrast against the THF solvent.27 Samples with nominal concentrations of 10, 50, and 100 mg/ mL were prepared by dissolving in anhydrous THF for 24 h, filtering with 0.45 μm pore polypropylene syringe filters, and loading the solutions into 2 mm diameter (0.01 mm wall thickness) quartz capillary tubes from Charles Supper Co. The z-average gyration radii (Rg) were determined from Zimm plots, after subtraction of a pure THF reference, as shown in Figure 2 for 50 and 100 mg/mL,27 according to the equation ⎛ q2R g2 ⎞ 1 1 ⎜ ⎟ = 1+ P(q) P(O) ⎜⎝ 3 ⎟⎠

Figure 2. Zimm plots of the SAXS data for halogen-containing modified lignins at 50 and 100 mg/mL nominal concentrations. The data have been offset vertically by arbitrary amounts.

Table 1. Results of Halogen-Containing Modified Lignins from SAXS Using Eq 1 sample

10 mg/mL Rg (Å)

50 mg/mL Rg (Å)

100 mg/mL Rg (Å)

BrAcL BrBenL ClAcL ClBenL

25 28 27 28

25 28 24 25

23 25 22 23

implies that the molecular structures of the solvent-extracted lignins are somewhat rigid, unlike linear polymer chains in solution, and that the modifications imposed negligible changes to the basic lignin molecular structures.27,28 Therefore, the acyl chloride reactants behaved like ligands. Unfortunately, the q-range and sensitivity were not sufficient to extract more information from the SAXS data, even with the enhanced electron density contrast. Therefore, small-angle neutron scattering (SANS) was performed on AcL at the BioSANS beamline at the High-Flux Isotope Reactor (HFIR) at Oak Ridge National Laboratory (ORNL) with neutron wavelengths of 12 Å at a detector distance of 15.338 m and 6 Å at detector distances of 6.838 and 1.138 m. Neutron scattering relies on scattering length density contrast rather than electron density contrast, and therefore only AcL was analyzed with SANS, as halogens can actually reduce scattering contrast against an isotopically labeled solvent. Samples were dissolved in deuterated THF (dTHF, 99.5% C4D8O), which provides high coherent scattering contrast while minimizing incoherent scattering, at concentrations of 90 (OS and AcL), 150 (AcL), and 250 (AcL) mg/mL, filtered with 0.2 μm pore polypropylene syringe filters, placed in 2 mm path length cylindrical quartz cells, sealed, and stored for 4 days prior to SANS analysis in order to allow complete dissolution of the somewhat high concentration solutions.24 Scattering of the samples, pure dTHF reference, and an empty cell, was determined over a q-range of approximately 0.002 to 0.6 Å−1, with good signal-to-noise over the q-range ∼0.006 to 0.6 Å−1. Figure 3 shows absolute scattering profiles (Figure 3a), after subtracting both buffer and incoherent scattering, as well as Kratky plots (Figure 3b) of all four samples. The solid lines in Figure 3a are fits to the Debye function27 for OS and a spherical form factor convoluted using the maximum entropy method29 for polydispersity for AcL. The Debye function and maximum entropy method are described further in Supporting Information. The OS results show behavior typical of a linear oligomer, with a clear plateau at low-q (Figure 3a) and non-Gaussian

(1)

Table 1 shows the Rg values measured using eq 1. The decrease in Rg with increasing concentration is due to excluded volume effects, a well-known phenomenon with nonassociating macromolecules in good solvents.28 However, the relative invariability of Rg ( 0).27 From the intrinsic viscosity results, the hydrodynamic radius (Rh) could be approximated by42 ⎛ 3M[η] ⎞1/3 Rh ≈ R η = ⎜ ⎟ ⎝ 10πNA ⎠

(2)

where M is the molecular weight and NA is the Avogadro constant. From this, Rh/Rg ≈ 1.0−1.2 using the ESI-determined absolute molecular weight range (18−30 kDa) and the zaverage Rg at 90 mg/mL (24 Å). Note that polydispersity was not taken into account with respect to Rg, but the Zimm analysis approximates the z-average Rg (high-end of the distribution), and accounting for polydispersity will only shift 571

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the approximation of Rh/Rg upward (i.e., Rh/Rg > 1.0−1.2), implying a rather compact molecular structure. The compact structure is further evidenced with the discrepancy between the GPC-determined Mw and the ESIdetermined Mw. The absolute Mw determined using ESI is approximately an order of magnitude greater than the relative GPC-determined Mw. This type of phenomenon has been shown previously upon intramolecularly cross-linking individual linear polystyrene chains, thereby converting them into nanoparticles (nanogels), which reduces the relative GPCdetermined Mw significantly due to changes in hydrodynamic volumes, even though light scattering shows negligible changes in absolute Mw.43 Here, the GPC-determined Mw of AcL was measured relative to linear polystyrene standards, an analogous situation as that reported in ref 43. Finally, Kratky plots and maximum entropy fits to spherical form factors also show compact molecular structures indicative of polydisperse nanogels (see Figure 3).44 However, Rh/Rg = 1.29 for hard spheres, Rh/Rg ∼ 0.6−0.7 for linear polymers,45 and Rh/Rg ∼ 0.8−1 for hyperbranched polymers.46−48 Therefore, this would indicate that the complex molecular architectures of the lignins investigated here range from compact nanogels to hyperbranched macromolecules. These results are consistent with a degelation process for extraction of lignins.13 With network ether bonds being stochastically broken, as well as a small amount of condensation reactions taking place in an already heterogeneous chemical “landscape,” one would anticipate a complex nanogel-type of structure with the presence of free branches. The OS analyzed in this work shows Rh/Rg ∼ 1.2, as determined from eq 2 using previously reported [η] = 4.2 mL/ g (interpolation)49 and the Debye Rg determined here (10 Å). The segmental distribution identifiable from the Kratky plot is indicative of a linear oligomer,27,30 with Rh/Rg > 1 supporting the linear oligomeric structure of OS50 (as chain length increases, Rh/Rg → 0.6−0.7).49 When scattering and viscometric results for AcL and OS are compared, it is clear that lignins are not simple linear oligomers or physical assemblies of linear oligomers, further demonstrating the complex architectures of extracted lignins and the three-dimensional network structures of protolignins. Unfortunately, unlike structural and dynamic analyses of model linear synthetic polymers with low polydispersities,27 the results determined here must be considered ensemble averages, as individual lignin macromolecules will vary significantly, both chemically and physically, within a particular batch of lignins extracted from a single source using a single extraction method.8 The molecular weights and architectures will also vary with time over the course of the extraction process, and the lignin chemistry will depend on the location within the plant cell.13 Such complexities explain the challenges in replacing traditional synthetic thermoplastics with lignins for many applications,5,6 but the methods outlined here can be used to guide genetic engineering technologies as well as extraction and postextraction processing methods in order to meet commercial and industrial needs by providing lignin architectures more suitable for advanced materials technologies.9 We have shown that lignin, a ubiquitous biopolymer, has a somewhat rigid molecular structure that ranges from nanogels to hyperbranched macromolecules. This is consistent with a lignin existing in the plant cell as an extremely complex, heterogeneous network.7 Future work should focus on the comparison of structures of lignins from various sources,

including hardwoods and softwoods, with particular attention to switchgrasses, which show great promise as an abundant biomass feedstock.2 We anticipate analogous, complex structures will be determined with other feedstocks and extraction methods, as can be inferred from similar trends in GPC and dilute solution viscosity data over a large permutation of sources and delignification methods (e.g., [η] ∼ M0.1−0.3).3,21−23 The scattering techniques employed here (SANS and SAXS) could also be used, along with various modification and labeling techniques, to probe structures of lignins in vivo (protolignins) coupled to kinetic evaluations of lignin structures as a function of extraction time using various extraction techniques and conditions in order to optimize future materials and biofuels production from renewable resources.51

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ASSOCIATED CONTENT

S Supporting Information *

Information regarding materials preparation and characterization. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]; [email protected]. Notes

The authors declare no competing financial interests.

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ACKNOWLEDGMENTS The authors gratefully acknowledge discussions with Prof. Wolfgang Glasser (Virginia Polytechnic Institute and State University) and Dr. Frederick Baker (Big Island Carbon, Hawaii, U.S.A.). S.E.H. and G.A.N. acknowledge funding from the Department of Energy, Office of Energy Efficiency and Renewable Energy. S.V.P., V.S.U., and P.L. were partly supported by a grant (ERKP752) from the Department of Energy, Office of Biological and Environmental Resources. BioSANS is operated as a user facility at Oak Ridge National Laboratory for the Department of Energy, Office of Biological and Environmental Resources. SAXS was performed at beamline X10A at the National Synchrotron Light Source, which is supported by the Department of Energy, Office of Basic Energy Sciences. The authors also wish to thank Steve Bennett (NSLS).

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NOTE ADDED AFTER ASAP PUBLICATION This Letter posted ASAP on April 17, 2012. The Supporting Information file has been revised. The correct version posted on April 19, 2012.

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