Two-Dimensional 1H-Nuclear Magnetic ... - ACS Publications

Research Article pubs.acs.org/journal/ascecg

Two-Dimensional 1H‑Nuclear Magnetic Resonance Relaxometry for Understanding Biomass Recalcitrance Tina Jeoh,*,† Nardrapee Karuna,†,‡ Noah D. Weiss,§ and Lisbeth G. Thygesen§ †

Biological and Agricultural Engineering, University of California, One Shields Avenue, Davis, California 95616, United States Biotechnology, Faculty of Engineering and Industrial Technology, Silpakorn University, 6 Rajamankha Nai Road, Amphoe Muang Nakhon Pathom, Thailand 73000 § Geosciences and Natural Resource Management, University of Copenhagen, Rolighedsvej 23, 1958 Frederiksberg C, Denmark ‡

S Supporting Information *

ABSTRACT: Low-field nuclear magnetic resonance (LFNMR) relaxometry examines the influence of the local environments within porous materials on the responses of the water-associated 1H to magnetic fields, yielding information on the chemical and physical surroundings of the water. 1D NMR relaxometry has been used to examine the relationship between water constraint within lignocellulosic biomass microstructure and its enzymatic digestibility; however, the effect of physical confinement and magnetic dephasing by the local chemistry could not be decoupled. This limitation is overcome by twodimensional T1T2 1H NMR relaxometry, where simultaneously probing the spin−lattice and spin−spin relaxation times of water 1 H resolves physical and chemical contributions to relaxation times of unique water environments within the sample. 2D T1T2 relaxation revealed four water environments in Norway spruce assigned to lumen and cell wall water based on water mobility in the pools. Sulfur dioxide (SO2) pretreatment of the spruce eliminated the cell wall water environments, while increasing the mobility of water in the lumens. Subsequent dewatering of pretreated spruce to high dry matter content in the samples significantly decreased water mobility in the lumens without changing the local chemical composition. For the first time, the use of 2D T1T2 relaxation revealed that osmotic pressure exerted by solute (glucose or bovine serum albumin (BSA)) uptake into the microstructure of lignocellulosic biomass expands the volume of the confined spaces such as the cell lumens. The uptake of BSA was associated with increased water retention and enzymatic digestibility of SO2 pretreated spruce. Overall, 2D T1T2 relaxation results suggest a relationship where increasing water mobility in the biomass microstructure reduces its recalcitrance. KEYWORDS: Lignocellulosic biomass, Recalcitrance, 2D T1T2 1H NMR relaxometry, T1 relaxation, T2 relaxation

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INTRODUCTION

Studies using X-ray diffraction and scattering, molecular dynamics simulations and size-exclusion chromatography have shown that pretreatment impacts molecular arrangements within cellulose microfibrils,4 nanoscale interfibril interactions,5,6 and micron-scale porosity and particle sizes7 of biomass. Moreover, pretreatment alters the overall3 and localized chemical composition of biomass.8 The resulting accessibility of the structural polysaccharides to lignocellulytic enzymes9−11 is assessed by measuring physical accessibility of molecular probes within the biomass structure12,13 or cellulose hydrolysis rates by cellulase enzymes.14 Greater accessibility of molecular probes in the biomass suggest successful disruption of the biomass microstructure to minimize mass transfer resistances and higher hydrolysis rates suggest improved opportunities for cellulase enzymes to complex to and hydrolyze cellulose.15 However, to design an effective pretreat-

Biomass recalcitrance, that is, the resistance of plant cell walls to deconstruction by chemical and biological processes, serves the plant well during its lifetime but presents a challenge when the biomass is used as a feedstock for the production of chemicals and liquid fuels, such as cellulosic bioethanol.1 Overcoming biomass recalcitrance is critical to the economic viability of industrial bioconversion processes as it will minimize capital and operation costs by reducing energy, water, and chemical inputs.2 The current state of technology for availing fermentable sugars from plant biomass is initiated by a thermochemical pretreatment step to improve downstream enzymatic hydrolysis of structural carbohydrates to glucose and other monosaccharides.3 The thermochemical pretreatment step, conducted at temperatures ranging from 160−200 °C with or without alkali, acid, or oxidative catalysts, physically disrupts the structure of the biomass, while altering the cell wall chemistry by a combination of solubilization, hydrolysis and oxidation of the polysaccharides and lignin. © 2017 American Chemical Society

Received: May 19, 2017 Revised: July 17, 2017 Published: August 25, 2017 8785

DOI: 10.1021/acssuschemeng.7b01588 ACS Sustainable Chem. Eng. 2017, 5, 8785−8795

Research Article

ACS Sustainable Chemistry & Engineering

biomass leads to improved enzymatic digestibility of the biomass.

ment with predictable impact on the enzymatic digestibility of biomass, the relationship between physical and chemical changes to the biomass and enzyme accessibility in pretreated biomass must be elucidated. A complementary approach to understand biomass recalcitrance has been to explore the state of water within the biomass by 1H NMR relaxometry.16−21 The underlying hypothesis of this approach is that localized biomass-water interactions can provide insights into both the physical and chemical environments and the accessibility of lignocellulytic enzymes within the biomass structure. While 1D NMR relaxation have provided insights into the biomass porosity16 and surface chemistry,18 the work carried out so far has also made it clear that interpretation of water populations is tentative and requires additional information.21 The challenge is that pore sizes, particle sizes, surface chemistry of the solids and solutes in the liquid phase all affect the relaxation times of water-associated 1 H. One strategy is to untangle these effects by eliminating confounding factors. For example, the contribution of porosity and surface chemistry was minimized in the study of the effect of soluble monosaccharides present during hydrolysis by using low (5%) solids,19 while the contribution of solutes was minimized by extensive washing of the solid biomass before hydrolysis.22 Another strategy is by multidimensional NMR relaxation. Two-dimensional (2D) T1T2 1H NMR relaxation records a matrix of correlated spin−spin (T2) and spin−lattice (T1) relaxation times for each sample that facilitates resolution of physical confinement and chemical composition influence on cellular and subcellular water populations in plant biomass.23 For example, 2D NMR relaxation is used to assess postharvest produce quality and shelf-stability by studying changes in cellular water distribution due to microstructural and compositional changes in the fruit or vegetable.24−26 Two published studies with lignocellulosic biomass27,28 identified three unique water pools in the T1T2 1H NMR relaxation spectra of softwood at varying moisture contents. The pool with the longest T1 and T2 relaxation times was assigned to water in the cell lumens and two pools with the same T2, but different T1 relaxation times were assigned to water in the cell walls. However, while Cox et al.27 speculated that one pool is likely water associated with cell wall polysaccharides while the other pool resides in clusters/voids within the cell wall, Bonnet et al.28 hypothesized that both populations comprise water bound to biopolymers, but in two different types of locations. 1D T2 relaxation studies of wood only identify one cell wall water pool because of the inability to further resolve this pool.29,30 T1T2 1H NMR relaxation has the power to resolve the contribution of porosity and local chemical influences on water relaxation times within a porous structure, a distinct advantage for lignocellulosic biomass that is a porous structure with spatial heterogeneity in chemical composition. Where 1D relaxation profiles can reveal porous structures of relatively chemically homogeneous material, such as rocks31 and hydrogels,32 such applications have eluded lignocellulosic biomass.18 Thus, the aim of the present work was to explore the use of 2D T 1T 2 1H NMR relaxation to understand biomass recalcitrance by studying the impact of mechanical and thermochemical treatment of lignocellulosic biomass on local water environments and enzyme accessibility in the biomass. We demonstrate the use of the method to resolve changes to the physical and chemical properties of intracellular and subcellular water environments within lignocellulosic biomass and show that increasing physical mobility of water in the

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EXPERIMENTAL SECTION

Spruce Sample Preparation. Norway Spruce (Picea abies (L.) Karst.) was harvested at ATA Timber Widtskövle AB, Everöd, Sweden, and chipped to pass a 20 mm screen. Thereafter, the material was pretreated in a steam gun at Lund University, Sweden, as previously reported.22 Pretreatment of the spruce with SO2 was carried out at three different temperatures (195, 200, and 210 °C) for 5 min. Compositional data was obtained following NREL standard procedures33 and has also been reported previously.22 The pretreated materials were washed thoroughly to remove solutes before all analysis. Unpretreated spruce chips were water extracted under reflux in a Soxhlet system for 24 h prior to analysis. Samples for NMR were prepared from the washed pretreated material by removing excess water via centrifugation (1000 rcf for 10 min), decanting the liquid fraction, and adding approximately 0.4 g (wet basis) of the insoluble residue to HPLC vials with a screw cap to retain moisture. This resulted in dry matter (DM) contents of 16%, 15%, and 12% for the 195, 200, and 210 °C pretreated samples, respectively. High solids samples were prepared by first removing water from the pretreated samples by centrifugation over a 0.45 μm filter (3000 rcf for 15 min) and then adding water back to achieve 30% DM. Samples were then weighed into HPLC vials with a total wet sample weight of 0.4 g. Samples were prepared in triplicate and stored at 4 °C, and equilibrated to 40 °C before analysis by NMR.

Figure 1. Low (16%, 15%, and 12% for 195, 200, and 210 °C, respectively) and high (30% w.b.) solids pretreated spruce samples. Equilibration of Samples in Solutes. Glass filter frits with a pore size 10−16 μm, unpretreated spruce, and pretreated spruce were soaked in nanopure water, 50% α-D-(+)-glucose (CAS no. 50-99-07, Sigma-Aldrich, USA), or 10% (w/v) bovine serum albumin (BSA) (CAS no. A7906, Sigma-Aldrich, USA) solution in 15 mL centrifuge tubes. The samples equilibrated in water or 50% glucose were incubated at 65 °C, with shaking at 100 rpm. The samples equilibrated in BSA solution were incubated at room temperature (∼22 °C) at 250 8786


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Figure 2. Interpreting the 2D T1T2 correlation spectra. (A) The dependence of relaxation times on the molecular tumbling rate of water molecules. The Larmor frequency, fo, is a function of the gyromagnetic ratio and magnetic field strength, fo = γBo. At frequencies below the fo, T1 relaxation times increase, while T2 relaxation times continue to decrease (after an initial small increase in T2 relaxation time reflecting the T1 relaxation process). The Larmor frequency in these studies is 20.0 MHz. Adapted from Bloembergen et al. 1948.34 (B) An example 2D correlation spectrum illustrating hypothetical water pools in a heterogeneous, porous structure, such as lignocellulosic biomass. T1 = T2 along the solid line bisecting the plot area. Pool w is free water with longest T1 and T2 relaxation times. Pool 2 has shorter T1 and T2 relaxation times than pool w, but similar T1/T2 ratios, which can be visually seen as a displacement parallel to the T1 = T2 line. Pool 3 shares a similar T1 relaxation time as pool 2 but a considerably shorter T2 relaxation time. rpm to avoid BSA denaturing. All samples were incubated for >1 week to reach equilibrium. After equilibrium, the solids and supernatant were separated by centrifugation at 2000g for 10 min. ∼1 g of the solids and 0.9 mL of supernatant were transferred to HPLC vial with cap and equilibrated to 40 °C before NMR measurements. NMR Measurements and Data Analysis. Two-dimensional 1H T1T2 NMR correlation signals were collected using a 0.47 T (20.0 MHz 1H resonance frequency) permanent magnet NMR (Bruker mq20 minispec NMR, Massachusetts, USA) at 40 °C. The T1T2 correlation spectra were acquired by a saturation recovery-CPMG sequence (Samsi A/S, Trollåsen, Norway). The saturation recovery measurement started after 3 to 6 ms (depended on the sample) to avoid receiver artifacts. Each recovery step of 40-steps of saturation recovery (T1 measurement) was followed by a CPMG sequence (T2 measurement) with echo time (TE) 0.2 ms for 12 000−32 000 echoes to obtain full decay. The T1T2 correlation data were analyzed using a 2D Laplace Inversion program (Victoria University of Wellington, Wellington, New Zealand). Determining the Water Retention Value (WRV) of Biomass Samples. Water retention value (WRV) was measured according to a modified version of the standard Scandinavian pulp, paper, and board testing committee method (SCAN-C 62:00, 2000). WRV was measured on either washed pretreated materials or on pretreated material soaked in 10% (w/v) BSA solutions as described above. WRV measurement was carried out by centrifugation at 3000 rcf for 15 min above a 45 μm nylon filter supported by a 200 mesh metal grating in 50 mL centrifuge tubes, with a target dry material weight of 0.48 g the wet mass of the material was measured after centrifugation and then again after 48 h at 40 °C under vacuum. The added mass of the BSA solution (10% of liquid mass) was taken into account in the determination of the WRV of the materials, and is reported as such for the BSA soaked samples. Enzymatic Saccharification of Spruce Samples. Enzymatic hydrolysis was carried out on washed pretreated spruce samples, BSAsoaked pretreated spruce samples, and Avicel, both with and without soaking with BSA. Hydrolysis reactions were carried out in 50 mL tubes free falling in a tumbler reactor at 15 rpm for 72 h at 50 °C. The commercial enzyme preparation, Cellic CTec 2 (Novozymes A/S, Baagsvaerd Denmark), was used for the hydrolysis at a constant loading of 15 mg enzyme protein/g cellulose. The pH was adjusted to 5.1 using a 50 mM citrate buffer solution at the stated pH. An initial total insoluble solids concentration of 10% (w/w) was the target for the reaction start, however this was modified when adding BSA, which is described as follows. BSA soaked biomass samples were prepared for hydrolysis by combining washed pretreated biomass (1 g dry basis for a 10 g total reaction mass) with 1 g BSA dissolved in makeup water to produce a slurry with 100 g/kg insoluble biomass (assuming BSA is a

soluble solid). Non BSA samples were also prepared in this manner except without BSA addition. Samples were then incubated at 22 °C for 72 h on a shaker table to allow the BSA and biomass to equilibrate. Citrate buffer and the appropriate amount of enzyme were then added and the samples were then placed in the tumbler reactor for 72 h. The reaction was quenched by addition of 2 mL of 0.5 M HCl to each vial. Glucose and cellobiose concentrations were measured using a Ultimate 3000 (Dionex) HPLC coupled to a Refractive Index Detector (RID). Separation was done with a Phenomenex Resex ROA column run at 80 °C with an isocratic method using 5 mM H2SO4 as eluent.

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RESULTS AND DISCUSSION T1T2 Correlation Spectroscopy Decouple the Influence of Physical Constraint and Chemical Composition in Local Water Environments in Lignocellulosic Biomass. T1T2 1H NMR relaxation experiments measure T1 and T2 relaxation simultaneously in the samples. The low-field NMR (LFNMR) method used in these studies limits detection to water-associated 1H within the biomass. T1 relaxation within a given water environment is the most efficient with the shortest T1 relaxation time when the rotational frequency of the water molecules, here described as molecular tumbling rates match the Larmor or resonance frequency (fo) set by the spectrometer (Figure 2A).34 In the example spectra in Figure 2B, free/bulk water is shown as pool “w”, with the longest T1 relaxation time of the water molecules with highest molecular tumbling rates (i.e., most mobile) in the sample. Water in pool 2 has lower molecular tumbling rates (is less mobile) than in pool ‘w’, as evidenced by the shorter T1 relaxation time. In porous structures such as rocks,31 porous glass frits (Supporting Information S1) and lignocellulosic biomass,27 water molecules subject to compartmentalization, that is, greater physical confinement in smaller spaces/pores, exhibit shorter T1 relaxation times. The figure on which Figure 2A is based refers to a model system where the sample is a homogeneous liquid and all interactions have the same correlation time. Here we use changes to T1 and T2 relaxation times with preserved T1/T2 ratio as a measure of the extent to which the local environment is constraining the physical movement of the water molecules. We refer to this as the extent of “physical constraint” of the environment in which the pool of water resides. 8787


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Figure 3. T1T2 1H NMR relaxation spectra of unpretreated, water extracted spruce (A, E) and spruce pretreated with SO2 at (B, F) 195, (C, G) 200, and (D, H) 210 °C. T1 and T2 relaxation times are shown in Table 2. (A−D) show contour plots while (E - H) show 3D plots of the same data. The gray line represents T1 = T2.

biomass. Pool 1 (T1 = 129 ± 30 ms, T2 = 48 ± 7 ms) has the longest relaxation times and the largest magnitude. Thus, the largest fraction of water in the spruce sample (79 ± 2%) is also the most mobile water residing in the least physically confining environment. The T1/T2 ratio of pool 1 (3 ± 1) was about 2fold higher than that of pure water (T1 = 4.2 s, T2 = 3.2 s, T1/T2 = 1.3, Figure S1 in Supporting Information). The water in pool 2 (T1 = 47 ± 14 ms, T2 = 6 ± 1 ms) was less mobile than in pool 1 and experience greater influence of local static magnetic fields within the environment (T1/T2 = 9 ± 3). Water in pool 3 (T1 = 27 ± 14 ms, T2 = 1 ± 0 ms) was even less mobile than in pool 2 and with greater influence of localized static magnetic fields (T1/T2 = 23 ± 2). Thus, from pool 1 to pool 2 to pool 3, the water environments become increasingly physically confining and with increasing influence of the local chemistry on T2 relaxation. Pool 4, with the shortest longitudinal relaxation time (T1 = 3 ± 0 ms) is thus the most physically confining water environment; but this pool shares the same T2 relaxation time as pool 3 (T2 = 1 ± 0 ms). The T1/T2 ratio of pool 4 of 4 ± 1 is similar to that of pool 1 with T1/T2 = 3 ± 1, which indicates that in both pool 4 and pool 1, the relative contribution of T1 relaxation processes to T2 relaxation time is the same. In other words, although pool 4 is a much more physically confining environment than pool 1, water in both pools experience similar extents of spin dephasing. This can come about if the pool 4 environment is simply a smaller version of pool 1 such as a smaller pore with the same chemical composition. From comparable 2D T1T2 relaxation spectra of the heartwood and sapwood of spruce, Cox et al.27 assigned pool 1 to the cell lumens, and pools 3 and 4 to water associated with the cell wall. A comparable pool 2, however, was not observed in Cox et al’s spruce sample, perhaps because the measurements were performed on samples below fiber saturation while samples in this study were above fiber saturation. Fredriksson and Thygesen38 identified two cell lumen water peaks in the T2 spectra of water saturated spruce xylem, with one component having a T2 in the 5−15 ms range of pool 2. Bonnet et al.28 also observed three water pools in Douglas fir early and late wood below the fiber saturation point, one assigned to remnants of liquid water and two to bound water within cell walls. Thus,

In the liquid state, the T2 relaxation process of water 1H reflects the T1 relaxation process with maximum efficiency at the Larmor frequency (Figure 2A). Additionally, T2 relaxation also occurs due to the dephasing influence of local magnetic fields in the immediate vicinity of the water 1H.35 These local magnetic fields are generated by chemical constituents within the environment such as sugars, proteins and surfaces that only influence adjacent 1H. Shorter T2 relaxation times of water in biomass, therefore, can be due to both decreased mobility of the water molecules in a physically confining environment (referred to as “physical effects”) and the greater abundance of local static magnetic fields in close proximity to the water molecules within the local environment (referred to as “chemical effects”, i.e., a reduction in T2 that causes an increase in the T1/T2 ratio) Previous work applying 1H NMR relaxation methods with cellulosic materials measured only T2 relaxation16−18,36 where it was challenging to decouple physical and chemical effects on T2 relaxation times. When T1 and T2 processes are measured simultaneously, however, the T1/T2 ratio provides insight into the relative influence of the microstructure (affecting physical mobility) and composition (affecting magnetic dephasing) of the water 1H.26,37 In a pool with a T1/T2 ratio of 1, the T2 relaxation time is solely a reflection of the T1 relaxation process. Increased influence of the local chemistry on T2 relaxation can be seen by larger T1/T2 ratios.26,31 For example, in Figure 2B, pool 3 and pool 2 have similar T1 relaxation times, indicating that water in the two pools are similarly physically constrained. However, the shorter T2 relaxation time of pool 3 relative to pool 2 indicates a stronger influence of the local chemical composition in pool 3. Visually, relative locations of pools along a diagonal parallel to the T1 = T2 line shows decreases in T1 and T2 times with conserved T1/T2 ratios indicating only differences in the physical mobility of water molecules in these pools31 (Figure 2B). 2D T1T2 1H NMR correlation spectra, therefore, resolve water environments within lignocellulosic biomass and provides insight into the relative influence of microstructure and chemical composition within each pool. T1T2 1H NMR relaxation spectra of spruce. The 2D-T1T2 1 H NMR spectra of water-extracted spruce (Figure 3A and E) show 4 pools, suggesting 4 distinct water environments in the 8788


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Table 1. Composition, Water Retention Value, and Cellulase Digestibility of SO2 Pretreated Spruce in This Study22 composition (wt %) pretreatment temperature (°C) 195 200 210

glucan

acid insoluble lignin

hemicellulose (xylan, mannan, galactan, and arabinan)

water retention value (g H2O/g biomass)

cellulose hydrolysis yield after 72 h (wt %)

45 45.1 49.5 47.9

30 50.6 57.3 56.5

17.4 1.3 1.0 0.7

1.1 1.4 1.9

56 ± 0.5 76 ± 1.7 77 ± 0.7

Table 2. Relaxation Times and Magnitudes of Water Pools in the 2D T1T2 1H-NMR Spectra of Raw and Pretreated Spruce at Low Solids Content (12−16% w.b.)a 195 °C

unpretreated pool 1

pool 2

pool 3b

pool 4b

T1 (ms) T2 (ms) magnitude T1 (ms) T2 (ms) magnitude T1 (ms) T2 (ms) magnitude T1 (ms) T2 (ms) magnitude

(%)

(%)

(%)

(%)

129 ± 30 48 ± 7 79 ± 2 47 ± 14 6±1 5±1 27 ± 14 1±0 12 ± 2 3±0 1±0 3±2

T1/T2 3±1 T1/T2 9±3

966 ± 109 100 ± 16 69 ± 3 211 ± 24 5±0 5±0

200 °C T1/T2 10 ± 2 T1/T2 45 ± 6

589 ± 136 54 ± 21 88 ± 4 211 ± 24 6±3 11 ± 5

210 °C T1/T2 11 ± 5 T1/T2 38 ± 16

451 ± 51 51 ± 7 99 ± 1 135 ± 0 4±1 1±1

T1/T2 9±2 T1/T2 37 ± 6

T1/T2 23 ± 12 T1/T2 4±1

a

Standard deviation of triplicates shown. bPools 3 and 4 were only observed in two of three replicates of the unpretreated spruce samples (Figure S2).

The T1T2 1H NMR relaxation spectra of the pretreated spruce samples are shown in Figure 3. Pools 3 and 4 observed in unpretreated spruce are absent in all the pretreated samples, suggesting that these cell wall associated water environments were eliminated by the pretreatment. Consistent with the speculation that pools 3 and 4 are water associated with hemicelluloses, loss of these pools coincides with the drastic reduction of hemicellulose content in the spruce (Table 1). The T1 and T2 relaxation times of pools 1 and 2 increased significantly compared to the spectra of unpretreated spruce (Figure 3 and Table 2). The increase in T1 relaxation times indicates that pretreatment increased the mobility of the water in the two pools. The pretreatment likely disrupted the cells, resulting in a less physically confining environment for water in the lumens. Moreover, the significant increase in T1/T2 ratios post pretreatment (p < 0.05) suggest that the pretreatment impacted the local chemical composition of the two pools resulting in increasing spin dephasing influence likely due to increased concentrations of solutes within these environments despite the washing. The 2D T1T2 relaxation spectra thus indicate that SO2 pretreatment of spruce results in a loss of cell and cell wall integrity to eliminate cell wall water and increases the homogeneity of the interstitial water. Mechanical Dewatering of Pretreated Spruce Leaves Less Mobile Water in Smaller Pores. High solids bioconversion of biomass to biofuels and chemicals has vast process economic advantages from the reduction in wastewater generation and energy requirements and smaller equipment size requirements.44,45 In general, high solids processing refers to processing in situations with little to no free water present.46 The pretreated spruce samples in this study were conditioned to 30% (w/v) solids by centrifugation to remove excess moisture from the samples. While the low solids samples (12− 16% w/v) had a slurry consistency and a visible liquid fraction,

here, we tentatively assign pools 1 and 2 to water in the lumens, and pools 3 and 4 to water associated with the cell-wall. In native wood, most cell wall water is thought to bind to the hydroxyl groups in the hemicelluloses,39 especially as recent results indicate that only 2/3 of the surface hydroxyls on cellulose fibrils partake in hydrogen bonding as H-donors to water molecules.40 The macromolecular structure of the dominant S2 layer of the spruce xylem tracheid cell wall is thought to consist of microfibrils with a diameter of about 3−4 nm surrounded by glucomannan and arranged in a web-like pattern creating lenticel shaped compartments consisting of xylan and lignin.41 Based on this structural model, and in agreement with Bonnet at al.,28 one could speculate that the two cell wall water pools identified both by Cox et al.27 and in the present study (pools 3 and 4) are water associated with the two different hemicelluloses, localized as water associated with the microfibrils or with the lenticel-shaped spaces between them. SO 2 Pretreatment of Spruce Reduces Physical Constraint of Water and Alters the Chemical Composition of the Local Water Environments in Biomass. The efficacy of sulfur dioxide catalyzed thermal pretreatment at increasing the enzymatic digestibility of softwoods is well documented.42,43 The SO2 pretreated spruce samples used in this study exhibited increasing cellulose conversion (56−77%) with increasing pretreatment temperatures of 195−210 °C (Table 1).22 As expected, higher pretreatment temperatures increased hemicellulose removal. Most interestingly, higher pretreatment temperatures also resulted in biomass with higher water retention values (WRV). The higher WRVs correlated with higher enzymatic digestibilities of the pretreated spruce samples, suggesting that water retention can be indicative of the physical accessibility of enzymes within the biomass structure.22 8789


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Figure 4. 2D 1H NMR spectra of water in spruce pretreated at (A) 195, (B) 200, (C) 210 °C. Spectra for dewatered samples at 30% solids are in color and overlaid onto spectra at low solids in gray. Dashed lines are drawn parallel to the solid diagonal line at T1 = T2 to guide the eye. The T1/T2 ratios of (D) pools 1 and (E) 2. Similar letters indicate no significant differences (p < 0.05). Error bars represent standard deviations of triplicate samples.

Figure 5. Unpretreated (A, E) and SO2 pretreated spruce (B−D, F−H) incubated for >10 days in 50% (w/v) glucose at 65 °C (A−D) and 10% (w/ v) BSA solution at room temperature (∼22 °C) (E−H). The spectra of the corresponding control (incubated in water) are under-laid in grayscale. Relaxation times for all pools are tabulated in Table S2.

the 30% (w/v) solids samples were no longer fluid but had a “crumbly” consistency (Figure 1). Compared to pretreated spruce samples with lower dry matter content (DM %), a clear shift in the pools of the 2D T1T2 relaxation spectra of the 30% (w/v) solids samples to shorter T1 and T2 times were observed (Figure 4). The diagonal shift parallel to the T1 = T2 line of both pools 1 and 2, also seen as no changes in T1/T2 ratios, indicate that while the environment became more physically constraining, the local chemistry of the water pools did not change. This interpretation is consistent with the preparation of the sample

where moisture was effectively drawn out of the biomass by centrifugal force that may also have collapsed the macropores such as lumens in the biomass without altering the chemistry within these. An interesting observation is that, while the T1 and T2 relaxation times of pools 1 and 2 spanned a relatively wide range in the pretreated spruce samples with lower DM %, they converged to the same relaxation times in the higher solids samples. Thus, the 2D T1T2 relaxation spectra analysis suggests that removing excess moisture from pretreated biomass leaves less mobile water in smaller spaces in the material. One possibility to confirm the hypothesis of pore collapse is to 8790


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ACS Sustainable Chemistry & Engineering measure pore size distribution in the wet material47 before and after centrifugation; however, such validation was out of the scope of the current study. One well-documented challenge of high-solids bioprocessing of lignocellulosic materials is the increasingly complex rheology that result in poor mixing and mass transfer limited saccharification reactions.48,49 Accordingly, the 2D T1T 2 relaxation spectra indicates that the water residing within the biomass becomes less mobile under low moisture conditions, which was previously implicated with decreased diffusivities of soluble enzymes and sugars that further limit hydrolysis rates.17 Solute Uptake into Spruce Exerts Osmotic Pressure within the Pores. Unpretreated and pretreated spruce were equilibrated in solutions with either 50% (w/v) glucose or 10% (w/v) BSA and examined by 2D T1T2 1H NMR to assess the accessibility of the water pools in the biomass to the solutes. The presence of these solutes had the general effect of decreasing T1 and T2 relaxation times of the bulk solutions, with increasing concentrations within the range studied here resulting in shorter relaxation times and increasing T1/T2 ratios (Figure S3). Moreover, both glucose and BSA increase the constraint of water molecules in porous glass frits that can be seen as a decrease in T1 and T2 relaxation times (Table S1). The expectation, therefore, was that the uptake of either glucose or BSA into the water pools in the spruce microstructure would be observable as decreases in the relaxation times with similar diagonal shifts in the 2D relaxation spectra. Water within unpretreated spruce incubated in 50% (w/v) glucose or 10% (w/v) BSA, however, exhibited distinctly different trends than what was observed in the solution phase or in the pores of glass frits (Figure 5). While incubation of the biomass in solution caused noticeable shifts in the relaxation times of the 2D 1H NMR spectra, the shifts were generally to longer relaxation times rather than shorter. For unpretreated spruce, incubation in either glucose or BSA solutions resulted in an “upward” shift in the water pools of the 2D spectra due predominantly to increased T1 relaxation times (Figure 5A and E, Tables S2 and S3). Accompanying increases in the T1/T2 ratio of the corresponding water pools suggest that, while the environment became physically less constraining (longer T1 relaxation times), the local chemistry had a greater dephasing influence (shorter T2 relaxation times) due to the solutes in the liquid fraction. These observations suggest that solutes taken-up into the biomass exerted osmotic pressure on the walls of the confined spaces and thereby expanded the lumens. Moreover, volumetric expansion of the pores was not observed with glass frits because osmotic stress within these pores were not sufficient to overcome the stiffness of the pore walls. Instead, the observed response in the rigid, porous glass substrate was an increase in physical constraint of the water within the pores. Accessibility of Glucose in Spruce. Glucose was used to probe the accessibility of a small molecule (∼0.5 nm) within the microstructure of spruce. The influence of glucose is apparent in the increased relaxation times of pools 1−3 of unpretreated spruce, which indicates the accessibility of glucose within these environments (Figure 5A). Of two replicates of unpretreated spruce incubated in the glucose solution, pool 4 was clearly detectable in one replicate but only a trace in the second replicate that was measured 15 days after (Figure S4). The interpretation that pools 1−3 of unpretreated spruce are physically accessible to glucose such that glucose diffuses into these environments from outside the biomass is consistent with the assignment of pools 1 and 2 to water in the cell lumens.

Moreover, the influence of glucose on the relaxation times of pool 3 confirms Cox et al.’s conclusion that the cell wall associated pool 3 is more likely water in small clusters rather than water bound directly to cell wall polysaccharides.27 The 2D 1H NMR relaxation spectra of all the spruce samples incubated in the glucose solution exhibited an additional pool 0 with longer relaxation times than the pools originally associated with the biomass and with a T1/T2 ratio of 1.4 (Figure 5A−D and Table S2). A separate measure of the glucose supernatant in which the spruce was incubated gave a single pool with longer relaxation times than pool 0, of T1 = 756 ms and T2 = 453 ms, but with similar T1/T2 ratio (1.7) (Table S2). The 1H relaxation response of pool 0 is consistent with that of glucose solution at a higher concentration (e.g., Figure S5). Pool 0 is thus most likely residual glucose solution on the outer surfaces of the biomass that concentrated during equilibration in the sample vial. The pretreated spruce samples incubated in the glucose solutions did not exhibit pools of longer T1 relaxation times that was observed with unpretreated spruce in glucose (Figure 5B−D). In contrast, there were almost negligible changes in the T1 relaxation times but a slight increase in T2 relaxation times of pools 1 and 2 of pretreated spruce after incubation in glucose. For example, for spruce pretreated at 200 °C, the T1 times of pool 1 were 589 ± 136 and 581 ± 62 ms before and after equilibration in glucose, respectively. For the same pool, the T2 time increased slightly from 54 ± 21 to 93 ± 1 ms after equilibrating in glucose (Tables 2 and S2). If the pretreated spruce had unyielding pore walls, then the uptake of glucose into the biomass microstructure would have resulted in decreased T1 and T2 relaxation times (as was the case for glucose uptake in glass frits that have unyielding pore walls, Table S1). The relaxation times in pretreated spruce did not decrease when equilibrated in the glucose solution, indicating that the uptake of glucose into the pores of pretreated spruce must have exerted sufficient osmotic pressure to yield the pore walls and expand the pore volumes. The diffusion of glucose into pretreated spruce is driven by a concentration differential between the supernatant and the lumens. Thus, relative to pretreated spruce equilibrated in water, the T1 relaxation times in these pools appear to be unchanged. Moreover, the slight increase in T2 relaxation times arise because in contrast to a more crowded environment in glass frit pores equilibrated with glucose, the expanded lumen volumes decrease the dephasing influence of the solutes on the T2 relaxation times of the 1H in these pools. Accessibility of BSA in Spruce. BSA is a globular protein of 66.5 kDa with a diameter of ∼5 nm, approximately the size of a cellulase enzyme, such as Trichoderma reesei Cel7A (∼5.1 nm).50 The motivation for equilibrating spruce samples in BSA solutions was to probe the physical accessibility of the water pools in the biomass to a protein similar in size to cellulases. The expectation was to observe differences in the physical accessibility of the water pools to glucose and to BSA where BSA was expected to be excluded from smaller pore spaces. In the spectra of unpretreated spruce incubated in BSA, the T1 relaxation times of pools 1 and 2 increased significantly after equilibration in the BSA solution, suggesting the uptake of BSA into these pools resulted in the expansion of the lumen volumes due to increased osmotic pressure. Pools 3 and 4 were undetected in all three replicate samples (one replicate shown in Figure 5E) likely because BSA was excluded from these pools and water was drawn out by osmotic pressure in pools 1 and 2. 8791


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Figure 6. (A) Water retention value (WRV) of SO2 pretreated spruce samples equilibrated in water and in 10% (w/v) BSA solutions. (B) Glucose yield from SO2 pretreated spruce samples and Avicel after 72 h hydrolysis with Cellic CTec2. Samples were either equilibrated in buffer or 10% (w/ v) BSA prior to enzyme addition. Error bars are standard deviation of triplicates. Similar letters indicate no significant difference (p < 0.05).

The 2D T1T2 relaxation spectra thus indicate that the cell wall associated water is physically inaccessible to BSA but that water can move in and out freely. Pool 3 was accessible to glucose but inaccessible to BSA. BSA had a significant increasing influence on the relaxation times of pools 1 and 2 in pretreated spruce (Figure 5F−H). The spectra show diagonal shifts of the pools in the upper right direction, parallel to T1 = T2, indicating that BSA decreased the physical constraint of the water in these pools without significantly changing the influence of the local chemistry. These trends suggest that the uptake of BSA into pools 1 and 2 exerted osmotic pressure to result in an uptake of water, thereby expanding the lumens. Water uptake into pools 1 and 2 must be from the surrounding solution, which was evidenced by a measurable increase in water retention value (WRV) of the pretreated spruce samples that were incubated in BSA (Figure 6A). The impact of BSA in the biomass was significantly more pronounced than that of glucose despite the fact that the BSA molar concentration was 3 orders of magnitude less than that of glucose and the incubation temperature was significantly lower to avoid protein denaturation (RT and 65 °C for BSA and glucose incubation, respectively). BSA has been shown to adsorb to lignocellulosic biomass.51 Thus, BSA transport into water-containing spaces in the biomass is likely driven by both diffusion and adsorption while glucose transport is only driven by diffusion. Moreover, the volume occupied by a molecule of BSA is approximately 3 orders of magnitude greater than that of a molecule of glucose. Together, the outcome is observed where the uptake of BSA into the pores in the biomass exert considerably higher osmotic pressures than the uptake of glucose such that there is a measurable increase in T1 relaxation times of these pools. Similar results were obtained with both rice straw and wheat straw where equilibration in 10% (w/v) BSA solutions increased T1 and T2 relaxation times of the dominant pools while eliminating short relaxation time pools (Figure S6). These results are again consistent with the interpretation that BSA only accessed larger pores within the biomass, in which the presence of the protein exerted sufficient osmotic pressure to expand the pore volumes such that measurable increases in relaxation times were observed. BSA Uptake into Cellulosic Biomass Improves Cellulose Hydrolysis Yields. Equilibration of pretreated spruce in the 10% (w/v) BSA, not only increased WRV of the biomass but also increased the glucose yields from subsequent

enzymatic saccharification (Figure 6B). The improvement was least pronounced for the most severely pretreated spruce sample at 210 °C (∼2%). For the less severely pretreated spruce samples at 200 and 195 °C, 14−16% increases in glucose yields after 72 h of hydrolysis were found. Improvements in enzymatic saccharification yields of dilute acid pretreated corn stover51 and hot water pretreated mixed hardwoods52 preincubated with BSA have been previously reported. The authors suggested that the hydrolysis yield improvements were due to the adsorption of BSA to lignin that reduce nonspecific cellulase−lignin interactions in the biomass. The previous studies, however, saw insignificant improvement in Avicel hydrolysis from preincubation in BSA that was suggested as further support of their hypothesis that reduction in nonspecific binding of cellulases to lignin was the root of the improvements observed with pretreated biomass.51,52 In contrast, in this study, equilibration of Avicel in 10% (w/v) BSA resulted in a 15% increase in 72 h glucose yields (Figure 6B). The BSA concentration used in this study was 10-fold that of the previous study, and therefore, the positive effects seen in this study may not have been observable at the order of magnitude lower BSA concentrations. Avicel, a nearly pure commercial cellulose, is a commonly used cellulose 'standard' in hydrolysis experiments. The extensive processing history of microcrystalline cellulose products, such as Avicel, completely eliminates the wood cell structure and results in particles that are formed by hornified aggregates of cellulose fibrils.15 Thus, the pores within Avicel refer to voids between the aggregates that form the particle. The porosity of Avicel has been implicated in aiding the enzymatic digestibility of the cellulosic substrate.53 The 2D T1T2 NMR relaxation spectra of Avicel equilibrated in a 10% (w/v) BSA solution exhibited a major pool with negligible shift in the T1 relaxation time and a slight increase in T2 relaxation time compared to Avicel equilibrated in water (Figure S7). A similar outcome was observed for pretreated spruce equilibrated in a 50% (w/v) glucose solution (Figure 5B−D). As speculated for pretreated spruce equilibrated in glucose, BSA uptake into Avicel particles by Fickian diffusion likely expand the volumes of the pores of the water environments, such that the T1 relaxation times are longer relative to a scenario where the pore walls cannot yield. In contrast to BSA uptake by pretreated spruce samples, however, the WRV value of Avicel decreased slightly, suggesting that the expansion of the pores within the Avicel particle structure was sufficient to 8792


Research Article

ACS Sustainable Chemistry & Engineering accommodate equilibrium concentrations of the BSA molecules without additional water uptake. Taken together, these results suggest that uptake of BSA into the microstructure of cellulosic substrates can expand the volume of pore spaces and correlate favorably with increased enzymatic cellulose digestibilities. A possible explanation is that the pore expansion improves physical accessibility of cellulase enzymes into the biomass microstructure; however, this hypothesis needs to be probed further with targeted experiments beyond the scope of the current study.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Phone: 530-752-1020.

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ORCID

CONCLUSIONS 2D T1T2 1H NMR of lignocellulosic biomass resolve water environments within lignocellulosic biomass that cannot be resolved by either 1D T1 or T2 1H NMR alone, and provides insight into the relative influence of microstructure and chemical composition within each pool. Shorter T1 and T2 relaxation times indicated reduced mobility of the water molecules (e.g., because of a more physically confining environment such as a smaller pore), while greater values for T1/T2 (>1) ratios indicated greater influence of the local chemical composition on decreasing T2 relaxation times. Water saturated Norway spruce contained four distinct water environments corresponding to two water pools in the lumen and two water pools associated with the cell wall. Increasing severity of SO2 pretreatment resulted in environments with more mobile water molecules in more complex local chemistry. These results align well with the general expectation that thermochemical pretreatment solubilizes the cell wall and “opens the structure” of lignocellulosic biomass. Similarly, reduction in relaxation times suggested that mechanical dewatering resulted in smaller pores of increasing physical constraint on water molecules without affecting the local chemical composition. However, further detailed studies on the porosity of these samples are needed to verify these conclusions. For the first time, we demonstrated a measurable impact of osmotic stress in the biomass upon solute uptake on the microstructure. Solute uptake into a rigid, porous structure with unyielding pore walls resulted in less mobile water molecules due to the increase in osmotic pressure. In a structure with pore walls that yield under stress, the osmotic pressure exerted because of the uptake of solutes resulted in the expansion of the pore volume and uptake of water. This effect was observed for spruce, pretreated spruce, rice straw, and wheat straw and correlated with an increase in the enzymatic digestibility of cellulose in pretreated spruce. One reason for this increase may simply be because of increased physical accessibility of cellulases into the microstructure of the biomass because of expanded pore volumes. However, further investigations are necessary to fully understand the observed results.

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relaxation spectra of spruce equilibrated in 50% (w/v) glucose at 65 °C, relaxation times of water pools in spruce equilibrated in glucose and BSA solutions, 2D 1H NMR relaxation spectra of rice straw and wheat straw, and 2D T1T2-NMR relaxation spectra of Avicel (PDF)

Tina Jeoh: 0000-0002-0727-4237 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors would like to acknowledge Dr. Michael McCarthy and Dr. Matthew Augustine for advice and consultation on NMR spin relaxation theory and methods. We also thank Dr. McCarthy for providing the 2D Laplace Inversion program. The authors thank Mats Galbe from Lund University for providing access to and assisting with pretreatment of material used in this study. This project was funded by the National Science Foundation Engineering Directorate (CBET Grant No. 1055518) and the IGN International Academy (University of Copenhagen). N.K. was supported by the Royal Thai Government Scholarship. This project was also partially funded by the BioValue SPIR project. The authors thank Claus Felby and his group at the University of Copenhagen for providing facilities, support and consultation on the project.

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REFERENCES

(1) Himmel, M. E.; Ding, S. Y.; Johnson, D. K.; Adney, W. S.; Nimlos, M. R.; Brady, J. W.; Foust, T. D. Biomass recalcitrance: Engineering plants and enzymes for biofuels production. Science 2007, 315, 804−807. (2) da Silva, A. R. G.; Torres Ortega, C. E.; Rong, B. Technoeconomic analysis of different pretreatment processes for lignocellulosic-based bioethanol production. Bioresour. Technol. 2016, 218, 561− 570. (3) Wyman, C. E.; Dale, B. E.; Balan, V.; Elander, R. T.; Mosier, N. S.; Lee, Y. Y.; Gupta, R.; Thomas, S. R.; Hames, B. R.; Warner, R.; Kumar, R. Comparative Performance of Leading Pretreatment Technologies for Biological Conversion of Corn Stover, Poplar Wood, and Switchgrass to Sugars. In Aqueous Pretreatment of Plant Biomass for Biological and Chemical Conversion to Fuels and Chemicals; Wyman, C. E., Ed.; John Wiley & Sons, Ltd.: Chichester, UK, 2013. (4) Bellesia, G.; Chundawat, S. P. S.; Langan, P.; Dale, B. E.; Gnanakaran, S. Probing the Early Events Associated with Liquid Ammonia Pretreatment of Native Crystalline Cellulose. J. Phys. Chem. B 2011, 115, 9782−9788. (5) Langan, P.; Petridis, L.; O’Neill, H. M.; Pingali, S. V.; Foston, M.; Nishiyama, Y.; Schulz, R.; Lindner, B.; Hanson, B. L.; Harton, S.; Heller, W. T.; Urban, V.; Evans, B. R.; Gnanakaran, S.; Ragauskas, A. J.; Smith, J. C.; Davison, B. H. Common processes drive the thermochemical pretreatment of lignocellulosic biomass. Green Chem. 2014, 16, 63−68. (6) Silveira, R. L.; Stoyanov, S. R.; Kovalenko, A.; Skaf, M. S. Cellulose Aggregation under Hydrothermal Pretreatment Conditions. Biomacromolecules 2016, 17, 2582−2590. (7) Ishizawa, C. I.; Davis, M. F.; Schell, D. F.; Johnson, D. K. Measurement of Porosity on Dilute Sulfuric Acid Pretreated Corn Stover. J. Agric. Food Chem. 2007, 55, 2575−2581. (8) Donohoe, B. S.; Decker, S. R.; Tucker, M. P.; Himmel, M. E.; Vinzant, T. B. Visualizing Lignin Coalescence and Migration Through

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acssuschemeng.7b01588. The effect of pore size on 2D T1T2 1H NMR relaxation spectra demonstrated with water-saturated porous glass frits, 2D 1H NMR relaxation spectra of water-extracted spruce, solutes decrease 2D 1H NMR relaxation times of the bulk solution and in porous glass media, 2D T1T2 8793


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ACS Sustainable Chemistry & Engineering Maize Cell Walls Following Thermochemical Pretreatment. Biotechnol. Bioeng. 2008, 101, 913−925. (9) Jeoh, T.; Ishizawa, C. I.; Davis, M. F.; Himmel, M. E.; Adney, W. S.; Johnson, D. K. Cellulase digestibility of pretreated biomass is limited by cellulose accessibility. Biotechnol. Bioeng. 2007, 98, 112− 122. (10) Wiman, M.; Dienes, D.; Hansen, M. A. T.; van der Meulen, T.; Zacchi, G.; Liden, G. Cellulose accessibility determines the rate of enzymatic hydrolysis of steam-pretreated spruce. Bioresour. Technol. 2012, 126, 208−215. (11) Zhao, X. B.; Zhang, L. H.; Liu, D. H. Biomass recalcitrance. Part I: the chemical compositions and physical structures affecting the enzymatic hydrolysis of lignocellulose. Biofuels, Bioprod. Biorefin. 2012, 6, 465−482. (12) Esteghlalian, A. R.; Bilodeau, M.; Mansfield, S. D.; Saddler, J. N. Do Enzymatic Hydrolyzability and Simons’ Stain Reflect the Changes in the Accessibility of Lignocellulosic Substrates to Cellulase Enzymes? Biotechnol. Prog. 2001, 17, 1049−1054. (13) Hong, J.; Ye, X.; Zhang, Y. H. P. Quantitative Determination of Cellulose Accessibility to Cellulase Based on Adsorption of a Nonhydrolytic Fusion Protein Containing CBM and GFP with Its Applications. Langmuir 2007, 23, 12535−12540. (14) Karuna, N.; Jeoh, T. The Productive Cellulase Binding Capacity of Cellulosic Substrates. Biotechnol. Bioeng. 2017, 114, 533−542. (15) Jeoh, T.; Cardona, M. J.; Karuna, N.; Mudinoor, A. R.; Nill, J. Mechanistic kinetic models of enzymatic cellulose hydrolysisa review. Biotechnol. Bioeng. 2017, 114, 1369−1385. (16) Foston, M.; Ragauskas, A. J. Changes in the Structure of the Cellulose Fiber Wall during Dilute Acid Pretreatment in Populus Studied by H-1 and H-2 NMR. Energy Fuels 2010, 24, 5677−5685. (17) Roberts, K. M.; Lavenson, D. M.; Tozzi, E. J.; McCarthy, M. J.; Jeoh, T. The Effects of Water Interactions in Cellulose Suspensions on Mass Transfer and Saccharification Efficiency at High Loadings. Cellulose 2011, 18, 759−773. (18) Karuna, N.; Zhang, L.; Walton, J. H.; Couturier, M.; Oztop, M. H.; Master, E.; McCarthy, M. J.; Jeoh, T. The Impact of Alkali Pretreatment and Post-Pretreatment Conditioning on the Surface Properties of Rice Straw Affecting Cellulase Accessibility to Cellulose. Bioresour. Technol. 2014, 167, 232−240. (19) Hsieh, C. W. C.; Cannella, D.; Jorgensen, H.; Felby, C.; Thygesen, L. G. Cellulase Inhibition by High Concentrations of Monosaccharides. J. Agric. Food Chem. 2014, 62, 3800−3805. (20) Selig, M. J.; Hsieh, C. W. C.; Thygesen, L. G.; Himmel, M. E.; Felby, C.; Decker, S. R. Considering water availability and the effect of solute concentration on high solids saccharification of lignocellulosic biomass. Biotechnol. Prog. 2012, 28, 1478−1490. (21) Ibbett, R.; Wortmann, F.; Varga, K.; Schuster, K. C. A morphological interpretation of water chemical exchange and mobility in cellulose materials derived from proton NMR T2 relaxation. Cellulose 2014, 21, 139−152. (22) Weiss, N.; Thygesen, L. G.; Felby, C.; Roslander, C.; Gourlay, K. Biomass-Water Interactions Correlate to Recalcitrance and are Intensified by Pretreatment: An Investigation of Water Constraint and Retention in Pretreated Spruce Using Low Field NMR and Water Retention Value Techniques. Biotechnol. Prog. 2017, 33, 146−153. (23) Hills, B. P. Relaxometry: Two-Dimensional Methods. In Encyclopedia of Magnetic Resonance; John Wiley & Sons, Ltd., 2009. (24) Furfaro, M. E.; Marigheto, N.; Moates, G. K.; Cross, K.; Parker, M. L.; Waldron, K. W.; Hills, B. P. Multidimensional NMR crosscorrelation relaxation study of carrot phloem and xylem. Part I: Peak assignment. Appl. Magn. Reson. 2009, 35, 521−535. (25) Marigheto, N.; Venturi, L.; Hills, B. Two-dimensional NMR relaxation studies of apple quality. Postharvest Biol. Technol. 2008, 48, 331−340. (26) Zhang, L.; McCarthy, M. J. Effect of controlled atmosphere storage on pomegranate quality investigated by two dimensional NMR correlation spectroscopy. LWT - Food Science and Technology 2013, 54, 302−306.

(27) Cox, J.; McDonald, P. J.; Gardiner, B. A. A study of water exchange in wood by means of 2D NMR relaxation correlation and exchange. Holzforschung 2010, 64, 259−266. (28) Bonnet, M.; Courtier-Murias, D.; Faure, P.; Rodts, S.; Care, S. NMR determination of sorption isotherms in earlywood and latewood of Douglas fir. Identification of bound water components related to their local environment. Holzforschung 2017, 71 (6), 481−490. (29) Menon, R. S.; MaCkay, A. L.; Hailey, J. R. T.; Bloom, M.; Burgess, A. E.; Swanson, J. S. An NMR determination of the physiological water distribution in wood during drying. J. Appl. Polym. Sci. 1987, 33, 1141−1155. (30) Araujo, C. D.; MacKay, A. L.; Hailey, J. R. T.; Whittall, K. P.; Le, H. Proton magnetic resonance techniques for characterization of water in wood: application to white spruce. Wood Sci. Technol. 1992, 26, 101−113. (31) Song, Y. Q.; Venkataramanan, L.; Hurlimann, M. D.; Flaum, M.; Frulla, P.; Straley, C. T-1-T-2 correlation spectra obtained using a fast two-dimensional Laplace inversion. J. Magn. Reson. 2002, 154, 261− 268. (32) Austin, D. T. R.; Hills, B. P. Two-dimensional NMR relaxation study of the pore structure in silicone hydrogel contact lenses. Appl. Magn. Reson. 2009, 35, 581−591. (33) Sluiter, A.; Hames, B.; Ruiz, R.; Scarlata, C.; Sluiter, J.; Templeton, D. W.; Crocker, D. Determination of Structural Carbohydrates and Lignin in Biomass, NREL/TP-510-42618; National Renewable Energy Laboratory: Golden, CO, 2008. (34) Bloembergen, N.; Purcell, E. M.; Pound, R. V. Relaxation Effects in Nuclear Magnetic Resonance Absorption. Phys. Rev. 1948, 73, 679− 712. (35) Bloch, F. Nuclear Induction. Phys. Rev. 1946, 70, 460−474. (36) Felby, C.; Thygesen, L. G.; Kristensen, J. B.; Jorgensen, H.; Elder, T. Cellulose-water interactions during enzymatic hydrolysis as studied by time domain NMR. Cellulose 2008, 15, 703−710. (37) Snaar, J. E. M.; Van As, H. A method for the simultaneous measurement of NMR spin-lattice and spin-spin relaxation times in compartmentalized systems. J. Magn. Reson. (1969-1992) 1992, 99, 139−148. (38) Fredriksson, M.; Thygesen, L. G. The states of water in Norway spruce (Picea abies (L.) Karst.) studied by low-field nuclear magnetic resonance (LFNMR) relaxometry: assignment of free-water populations based on quantitative wood anatomy. Holzforschung 2017, 71, 77−90. (39) Engelund, E. T.; Thygesen, L. G.; Svensson, S.; Hill, C. A. S. A critical discussion of the physics of wood-water interactions. Wood Sci. Technol. 2013, 47, 141−161. (40) Lindh, E. L.; Bergenstrahle-Wohlert, M.; Terenzi, C.; Salmen, L.; Furo, I. Non-exchanging hydroxyl groups on the surface of cellulose fibrils: The role of interaction with water. Carbohydr. Res. 2016, 434, 136−142. (41) Salmen, L.; Burgert, I. Cell wall features with regard to mechanical performance. A review COST Action E35 2004−2008: Wood machining - micromechanics and fracture. Holzforschung 2009, 63, 121−129. (42) Mackie, K. L.; Brownell, H. H.; West, K. L.; Saddler, J. N. Effect of Sulphur Dioxide and Sulphuric Acid on Steam Explosion of Aspenwood. J. Wood Chem. Technol. 1985, 5, 405−425. (43) Galbe, M.; Zacchi, G. A Review of the Production of Ethanol from Softwood. Appl. Microbiol. Biotechnol. 2002, 59, 618−628. (44) Wingren, A.; Galbe, M.; Zacchi, G. Techno-economic evaluation of producing ethanol from softwood: Comparison of SSF and SHF and identification of bottlenecks. Biotechnol. Prog. 2003, 19, 1109− 1117. (45) Kristensen, J.; Felby, C.; Jorgensen, H. Yield-determining factors in high-solids enzymatic hydrolysis of lignocellulose. Biotechnol. Biofuels 2009, 2, 11. (46) Hodge, D. B.; Karim, M. N.; Schell, D. J.; McMillan, J. D. Model-Based Fed-Batch for High-Solids Enzymatic Cellulose Hydrolysis. Appl. Biochem. Biotechnol. 2009, 152, 88−107. 8794


Research Article

ACS Sustainable Chemistry & Engineering (47) Lin, J. K.; Ladisch, M. R.; Patterson, J. A.; Noller, C. H. Determining pore size distribution in wet cellulose by measuring solute exclusion using a differential refractometer. Biotechnol. Bioeng. 1987, 29, 976−981. (48) Lavenson, D. M.; Tozzi, E. J.; Karuna, N.; Jeoh, T.; Powell, R. L.; McCarthy, M. J. The effect of mixing on the liquefaction and saccharification of cellulosic fibers. Bioresour. Technol. 2012, 111, 240− 247. (49) Cardona, M. J.; Tozzi, E. J.; Karuna, N.; Jeoh, T.; Powell, R. L.; McCarthy, M. J. A process for energy-efficient high-solids fed-batch enzymatic liquefaction of cellulosic biomass. Bioresour. Technol. 2015, 198, 488−496. (50) Laymon, R. A.; Adney, W. S.; Mohagheghi, A.; Himmel, M. E.; Thomas, S. R. Cloning and expression of full-length Trichoderma reesei cellobiohydrolase I cDNAs in Escherichia coli. Appl. Biochem. Biotechnol. 1996, 57−58, 389−397. (51) Yang, B.; Wyman, C. E. BSA treatment to enhance enzymatic hydrolysis of cellulose in lignin containing substrates. Biotechnol. Bioeng. 2006, 94, 611−617. (52) Ko, J. K.; Kim, Y.; Ximenes, E.; Ladisch, M. R. Effect of liquid hot water pretreatment severity on properties of hardwood lignin and enzymatic hydrolysis of cellulose. Biotechnol. Bioeng. 2015, 112, 252− 262. (53) Peciulyte, A.; Karlstrom, K.; Larsson, P.; Olsson, L. Impact of the supramolecular structure of cellulose on the efficiency of enzymatic hydrolysis. Biotechnol. Biofuels 2015, 8, 56.

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