Multifunctional Cellulolytic Enzymes Outperform Processive Fungal

Multifunctional Cellulolytic Enzymes Outperform Processive Fungal Cellulases for Coproduction of Nanocellulose and Biofuels John. M. Yarbrough,† Ruoran Zhang,† Ashutosh Mittal, Todd Vander Wall, Yannick J. Bomble, Stephen R. Decker, Michael E. Himmel, and Peter N. Ciesielski* Biosciences Center, National Renewable Energy Lab, 1503 Denver W. Parkway, Golden, Colorado 80401, United States ABSTRACT: Producing fuels, chemicals, and materials from renewable resources to meet societal demands remains an important step in the transition to a sustainable, clean energy economy. The use of cellulolytic enzymes for the production of nanocellulose enables the coproduction of sugars for biofuels production in a format that is largely compatible with the process design employed by modern lignocellulosic (second generation) biorefineries. However, yields of enzymatically produced nanocellulose are typically much lower than those achieved by mineral acid production methods. In this study, we compare the capacity for coproduction of nanocellulose and fermentable sugars using two vastly different cellulase systems: the classical “free enzyme” system of the saprophytic fungus, Trichoderma reesei (T. reesei) and the complexed, multifunctional enzymes produced by the hot springs resident, Caldicellulosiruptor bescii (C. bescii). We demonstrate by comparative digestions that the C. bescii system outperforms the fungal enzyme system in terms of total cellulose conversion, sugar production, and nanocellulose production. In addition, we show by multimodal imaging and dynamic light scattering that the nanocellulose produced by the C. bescii cellulase system is substantially more uniform than that produced by the T. reesei system. These disparities in the yields and characteristics of the nanocellulose produced by these disparate systems can be attributed to the dramatic differences in the mechanisms of action of the dominant enzymes in each system. KEYWORDS: nanocellulose, biofuel, enzymatic hydrolysis, cellulase, multifunctional enzyme

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other classes of nanomaterials are enabling a myriad of emerging applications for CNFs and CNCs.5,6 The inherent nanostructure and macromolecular order of plant cellulose stems from its biological synthesis, in which cellulose is produced as ordered aggregates of several dozen chains of poly(1→4)-β-D-glucan, called elementary fibrils or nanofibrils.7 These structures are then sheathed in hemicelluloses and further aggregate into larger bundles. These filamentous carbohydrate assemblies form the scaffolding for the mesoscale architecture of plant cell walls which also includes pectins, proteins, and lignins.8 The precise nature of cellulose crystallinity has long been a topic of debate among cell wall researchers, but the presence of both highly crystalline domains and less-ordered (sometimes called “amorphous”) domains has been clearly evidenced by X-ray diffraction,9

anocellulose (NC) is rapidly emerging as an impressive class of renewable nanobiomaterials with widespread utility and can be produced from biomass at industrial scales. These materials can be produced from virtually any cellulosic feedstock, from hardwoods,1 to grasses,2 to algae.3 NC can be generally grouped into two classes of nanomaterials, consisting of cellulose nanofibers (CNFs) and cellulose nanocrystals (CNCs). Although considerable ambiguity surrounds the specific definitions of these materials, cellulose nanofibers are generally described as extremely highaspect ratio nanostructures consisting of cellulose microfibrils or bundles thereof with diameters ranging from a few to several hundred nanometers and lengths up to several microns. Nanocrystals are described as shard-like, highly crystalline microfibril fragments with diameters ranging from ∼3 to 10 nm and lengths from 50 to 500 nm, depending on the source and production method.4 The impressive properties of these materials, including high mechanical strength, biocompatibility, biodegradability, and relatively low cost compared to many © 2017 American Chemical Society

Received: January 5, 2017 Accepted: March 7, 2017 Published: March 7, 2017 3101

DOI: 10.1021/acsnano.7b00086 ACS Nano 2017, 11, 3101−3109

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ACS Nano Raman spectroscopy,10 and NMR.11 It is clear that the less ordered regions of cellulosic aggregates are more accessible to water and aqueous solutes, and are thus preferentially hydrolyzed when exposed to acid and enzymes.12 The production of NC is typically accomplished by subjecting delignified cellulosic feedstocks to various extents of mechanical and/or chemo-mechanical processing.13,14 The details of the production process can determine if the NC consists of CNFs, CNCs, or mixtures of both. For example, mechanical refining alone is sufficient to produce CNFs,15 although incorporating other enzymatic or chemical treatments, such as 2,2,6,6-tetramethylpiperidine-1-oxyl (TEMPO) oxidation, can reduce the energy requirements of refining and enhance fibrillation of cellulose bundles.16,17 The dominant production method for CNCs is controlled acid hydrolysis, which is often performed in tandem with mechanical refining.18 Sulfuric acid is the most commonly used hydrolytic reagent due to the charged sulfate groups introduced by esterification of surface hydroxyl groups during the hydrolysis step. These nonnative sulfates help stabilize the resultant CNCs in aqueous suspensions.19 However, the sulfate groups limit the suite of surface modifications available, and the extreme hydrophilicity of the sulfated nanocellulose makes drying and dewatering expensive. In contrast, hydrolysis with hydrochloric acid preserves the native hydroxyl surface functionality, which provides a more versatile platform for subsequent surface functionalization.20 One potential drawback of retaining the native surface chemistry of cellulose in CNC preparations is the propensity of these crystallites to aggregate and eventually “crash out” of aqueous solutions, especially at physiological ionic strength. While this characteristic may be problematic for aqueous-phase processing, it is likely a desirable property when significant dewatering is required to reduce shipping costs or for potential end-use applications that require dry particles. The integrated production of biofuels and value-added coproducts from all the components in biomass has emerged as goal of modern lignocellulosic biorefineries in order to achieve economic self-sustainability.21,22 Today, the primary biofuel production strategies are centered on the fermentation of 5- and 6-carbon sugars liberated from the polysaccharide components of the feedstock.23 Therefore, recovery of fermentable hydrolysate during nanocellulose production is critical for the integrated production of nanocellulose with biofuels in a format that is compatible with the typical downstream biochemical processing employed by biorefineries. For this reason, virtually all of the strong acid hydrolysis nanocellulose production methods previously established are incompatible with biorefinery processes, as the highly acidic hydrolysate is unsuitable for fermentation and requires costly neutralization. We also note that nanocellulose production is fundamentally incompatible with thermochemical conversion technologies (i.e., pyrolysis, gasification, hydrothermal liquefaction) because these processes result in the complete depolymerization and/or transformation of biopolymers to smaller, typically monomeric molecules. The opportunity for coproduction of nanocellulose and biofuels via methods that are largely compatible with second generation biorefinery technology was first reported by Zhu and colleagues at the USDA Forest Product Lab, who realized that enzymatic hydrolysis, rather than strong acid hydrolysis, could provide a sugar stream suitable for downstream fermentation.24 Indeed, enzymatic hydrolysis provides an ideal route to integrate production of nanocellulose and biofuels from

biomass. In the traditional process, hydrolytic enzymes with specific functionalities work in concert to efficiently depolymerize carbohydrates to produce a high-quality stream of sugars that is well-suited for downstream fermentative processing and/ or catalytic upgrading. Additional advantages to this process are afforded by the specificity of enzymes relative to chemical catalysts. Various enzymes target certain regions or functional groups of carbohydrate substrates with high selectivity.25 Reports of enzymatic production of nanocellulose have been sparse in the literature relative to acid hydrolysis methods, although there has been a resurgence of publications on this topic in recent years. In 2006, Janardhnan and co-workers showed that treating kraft pulp with OS1 fungal culture prior to mechanical refining could enhance the yield of microfibrillated cellulose.26 A year later, Henriksson et al.27 and Päak̈ kö et al.28 published investigations wherein endoglucanase treatment was demonstrated to enhance the efficacy of mechanical fibrillation of cellulose in the preparation of microfibrillated cellulose. In 2009, Filson and co-workers published exploratory work wherein endoglucanase was used to produce nanocellulose (both CNFs and CNCs) from recycled softwood pulp.29 These studies employed endoglucanase in order to avoid excessive hydrolysis. This resulted in relatively low yields of soluble sugars which is not a favorable outcome if coproduction of biofuels and NC is desired. These initial studies were followed by several detailed investigations of the integration of enzymatic treatment, using both endo- and exoglucanases, with mechanical refinement and acid hydrolysis for the production of nanocellulose of different sizes and aspect ratios.30,31 Soon thereafter, the opportunity for the coproduction of nanocellulose and biofuels by enzymatic routes was first reported by Zhu and co-workers at the USDA Forest Product Lab.24 Using commercially available cellulases and kraft hardwood pulp, this insightful work provided proof of concept that fermentable sugars and cellulose nanomaterials could be coproduced using enzymes. Since these foundational studies, additional investigations have been published regarding the use of commercially available cellulases to assist in the production of nanocellulose (primarily CNFs) from various feedstocks; including cotton,32 sugar cane bagasse,33 date palm stalks,34 banana peel,35,36 corrugated packaging,37 soybean straw,38 and various kraft pulps.39−41 We posit that a considerable disadvantage concerning the use of many commercially available cellulase formulations for the coproduction of nanocellulose and biofuels is the high content of processive exoglucanases, such as T. reesei Cel7A (cellobiohydrolase 1) and Cel6A (cellobiohydrolase II), which, in the presence of endoglucanases, aggressively depolymerizes crystalline cellulose.42,43 Since the majority of commercial cellulase cocktails are currently optimized for the highest possible conversion of cellulose to sugars, high titers of these processive cellulases are needed to depolymerize the most recalcitrant, highly crystalline cellulose in biomass. However, this functionality is not desirable if nanocellulose, particularly CNC, is a desired coproduct of the digestion. This paradox was recently described by Anderson, Zhu, and colleagues, who screened a series of enzymes with various functionalities for their propensity to produce nanocellulose.44 Their work identified several enzymatic functionalities, particularly cellulases from Aspergillus niger, that produced high quality NC. However, the reported NC yields were still low (∼10%) compared to acid hydrolysis methods. 3102


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Figure 1. Comparison of the structure of the processive exoglucanase, Cel7A (A), from the filamentous fungus, T. reesei (Tr) to that of the multifunctional, complexed bacterial cellulase, CelA (B), from the thermophilic bacterium C. bescii (Cb). Depicted are the respective catalytic domains (TrGH7CD, CbGH48CD, and CbGH9CD) and carbohydrate binding modules (CBM1 and CBM3). Conformation for Cel7A was obtained from Zhong et al.45 with O-glycosylation (shown in gray) added by Beckham et al.46 Structure and conformation of CelA obtained from Brunecky et al.47

Figure 2. Saccharification yields of cellulose conversion (A), soluble sugar production (B), and nanocellulose production (C) at 24, 48, 72, 96, and 120 h.

and more uniform nanocellulose particles produced by the C. bescii system relative to that of T. reesei.

In the present study, we compare the nanocellulose production capabilities of two cellulolytic enzyme systems with notably different functionality: the fungal enzyme system of T. reesei QM6a, and the thermophilic bacterial enzyme system of C. bescii. A structural comparison of the most abundant enzymes in each system, Cel7A from T. reesei and CelA from C. bescii, is presented in Figure 1. The T. reesei system is a classical cellulase system that consists of noncomplexed enzymes,48 the most prolific of which is the processive exoglucanase Cel7A, which contains a single CAZy (carbohydrate-active enzymes) carbohydrate-binding module family 1 (CBM1)49 and a single glycoside hydrolase family 7 (GH7) reducing end-acting cellobiohydrolase catalytic domain.50 This enzyme is particularly effective for the degradation of crystalline cellulose51,52 and is the most abundant and primary workhorse enzyme in commercial cellulase cocktails formulated for biomass conversion. In contrast, the C. bescii system contains multimodular and multifunctional enzyme complexes. The most abundant of these is CelA, which contains a GH9 endo-β-1,4-glucanase catalytic domain, a GH48 exo-β1,4-glucanase catalytic domain, and three family 3 carbohydrate-binding modules (CBM3), all separated by linker peptides. This complex enzyme has been recently shown to exhibit unrivaled activity on cellulose substrates with a cellulolytic activity far greater than that of Cel7A from T. reesei.47 Moreover, it employs a distinctly different degradation mechanism compared to Cel7A, wherein localized hydrolysis (in contrast to the processive, surface ablative hydrolysis of Cel7A) tends to result in “pit digging” into the substrate.47 By comparative digestion experiments, we demonstrate that these differences in degradation mechanisms result in improved overall cellulose conversion, enhanced yields of nanocellulose,

RESULTS AND DISCUSSION Saccharification Yields. The resultant yields of unconverted cellulose, soluble sugars, and nanocellulose produced by both enzyme systems through 120 h of saccharification time are presented in Figure 2. The superior performance of the C. bescii system in terms of overall cellulose conversion is clearly evidenced in Figure 2A. The unconverted cellulose digested by the C. bescii system continually decreases throughout the saccharification time, whereas the T. reesei system achieves a plateau of ∼60% unconverted cellulose and fails to significantly reduce this fraction further throughout the course of the digestion period. The soluble sugar production (shown in Figure 2B) of the C. bescii system similarly outperforms that of the T. reesei system, which is desirable for effective biofuel production. The NC fraction, defined as the mass fraction present in the supernatant following centrifugation less the soluble sugars, is presented in Figure 2C. Both systems displayed similar NC production of ∼30% after 24 h. However, the NC fraction produced by the C. bescii system continued to increase through 72 h of saccharification, whereas that of T. reseei system essentially plateaued after 24 h. The peak nanocellulose yield of 42% is achieved by the C. bescii system at 72 h, after which the NC fraction decreases, indicating that a portion of the NC is further digested to soluble sugars. The total glucan conversion glucan achieved by these digestions was lower than that reported in many studies of enzymatic saccharification in the context of biofuel production where complete conversion of glucan to soluble sugars is desired. We attribute this observation to the relatively low enzyme loadings intentionally employed here in order to avoid unwanted 3103


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with an intensity maximum at Rg of ∼255 nm. The corresponding image data for these samples (Figure 4B/B′ and C/C′) indeed reveal the presence of largely uniform CNCs. The TEM data presented in Figure 4A′−F′ indicate that the diameters of the individual nanofibrils and nanocrystals are largely similar across the digestion products of both enzyme systems, which is likely a resultant property of the elementary fibril diameter of the cellulose feedstock used in this study. Assuming that the cellulose nanoparticles characterized in this study may be treated as spheroids with largely similar minor axes, the radius of gyration increases with increasing aspect ratio.53,54 Therefore, the observation of larger Rg values for the nanocellulose fractions produced by T. reesei is consistent with the assertion that these samples contain larger fractions of high aspect-ratio CNFs relative to the nanocellulose produced by the C. bescii system. These significant changes in the particle size distribution, specifically the disappearance of the bimodal distribution and the emergence of a largely uniform population of particles sized between the two previously observed populations, indicate that the C. bescii system performed two significant modifications to the NC particles during the 24 to 48 h digestion period: first, the smaller CNC fraction initially observed after 24 h is completely digested; and second, the CNFs initially present after 24 h are fragmented into smaller CNC particles with a largely uniform size distribution. This assertion is further supported by the absence of high-aspect-ratio CNFs from the image data obtained from the NC produced by the C. bescii system at 48 h. The saccharification yield data presented in Figure 2 show that the fraction of nanocellulose produced by C. bescii system increases by ∼6% between the 24 and 48 h time points, which suggests that additional CNCs are generated from the unconverted cellulose during this period. The soluble sugar yield also increases by ∼15% during the 24 to 48 h digestion period which indicates that a significant amount of hydrolysis is associated with the CNF fragmentation performed by the C. bescii system. After 72 h of digestion the C. bescii system achieves the peak yield of nanocellulose but the particle size distribution is broadened. This broadening likely results from additional fragmentation of existing CNCs into smaller particles as well as some flocculation of particles into larger, more intimate assemblies. Some degree of flocculation is expected from enzymatically produced NC particles due to the lack of anionic groups on the particle surface.55 In contrast, the T. reesei system consistently produces a heterogeneous distribution of NC comprising mostly CNF, although some population of CNCs is present as evidenced by the DLS and imaging data. These results are qualitatively similar to those reported previously by Zhu and co-workers wherein the primary NC product of enzymatic hydrolysis was CNF using a combination of commercially available cellulase preparations, namely Novozym 476 (a purified endoglucanase) and Genencor Multifect B which contains a high titer of processive exoglucanases.24 We attribute the differences in the size and shape distributions of the NC particles produced by these two enzyme systems to differences in the degradation mechanisms of the most prolific enzyme in each system. This concept is depicted schematically and by TEM micrographs in Figure 5. Cel7A, the most abundant enzyme in the T. reesei system, binds crystalline cellulose and translates processively along the cellulose chain as hydrolysis occurs. This tends to result in fibrillation of larger cellulose bundles into smaller fibrils followed by thinning of the isolated fibrils. In contrast,

hydrolysis of the nanocellulose fraction to soluble sugars. In addition, these digestions were carried out in deionized water rather than typical buffer solutions (as described in the Experimental Section) in order to facilitate solubilization of nanocellulose due to the low ion concentration of the solution. This further limited the activity of the hydrolytic enzymes relative to their performance in buffer conditions optimized for complete hydrolysis of cellulose to soluble sugars. Characterization of the Nanocellulose Saccharification Products. Dynamic light scattering (DLS) studies were performed in tandem with multimodal imaging on the nanocellulose fractions produced by both enzyme systems at various time points to assess the size and shape distributions of the NC particles produced by saccharification. The light scattering intensity spectra for both enzyme systems at 24, 48, and 72 h are presented in Figure 3. After 24 h of digestion, the

Figure 3. Size distribution by dynamic light scattering of nanocellulose produced by cellulases from T. reesei (A) and C. bescii (B) at 24, 48, and 72 h of hydrolysis. Peak intensity values of Rg for each size population are shown under the respective distribution curve.

DLS results indicate several distinct populations of sizes are present in the nanocellulose particles produced by both enzyme systems: a smaller fraction with radii of gyration (Rg) of ∼100 to 200 nm and a larger fraction with Rg of ∼300 to 1000 nm. The AFM and TEM images of these same nanocellulose particles at 24 h (Figure 4A/A′ and D/D′) clearly show a mixture of CNC and CNF. After 48 h of digestion, the nanocellulose particles produced by the T. reesei system display a similar heterogeneous distribution of Rg values, which is consistent with the corresponding with the image data (Figure 4 E/E′ and F/F′) that again shows a mixture of CNCs and CNFs. In contrast, the NC produced by C. bescii after 48 h is substantially more uniform as evidenced by a single DLS peak 3104


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Figure 4. AFM (A−F) and TEM (A′−F′) images of nanocellulose particles produced by T. reesei and C. bescii at 24, 48, and 72 h.

CONCLUSIONS We have compared the nanocellulose production capabilities of the T. reesei QM6a secretome, a classic fungal cellulase system containing predominantly processive exoglucanases, with that of C. bescii, a newly discovered bacterial enzyme system that contains mainly complexed multifunctional enzymes. We have demonstrated by comparative digestions that the C. bescii system outperforms the T. reesei system in terms of total cellulose conversion, sugar production, and NC production. We have also shown by DLS in tandem with multimodal imaging that the NC produced by the C. bescii system is considerably more uniform than that produced by the T. reesei system, particularly after 48 h of digestion at which time largely uniform CNCs were produced. The differences in the NC production capacities of the two enzyme systems are attributed to the distinct functionality of CelA, the dominant cellulase secreted

the multifunctional CelA complex from C. bescii binds cellulose in multiple locations and performs localized hydrolysis which results in the “pit digging” action described previously.47 We postulate that this localized hydrolytic action of CelA promotes fragmentation of CNFs into CNCs and is responsible for the more uniform nanocellulose size and shape distributions produced by the C. bescii system. The presence of periodic defects is widely speculated to exist in cellulosic substrates56 and could give rise to locations at which CelA may initiate hydrolysis. The propensity of this enzyme complex to remain localized once actively engaged, combined with the aforementioned presence of periodic defects in the substrate, could give rise to the largely uniform population of CNCs observed at 48 h of digestion by the C. bescii system. 3105


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Figure 5. Depiction of the differences in degradation mechanisms of Cel7A (T. reesei) and CelA (C. bescii). (A,B) T. reesei Cel7A primarily binds to free reducing ends and proceeds down a single cellulose surface chain with each hydrolytic event. (C) TEM micrograph of showing CNFs of various widths produced by digestion with the T. reesei enzyme system. (D,E) C. bescii CelA tends to bind and “excavate” a single locus, resulting in more intrafibril fragmentation than processive cellullases. (F) TEM micrograph of cellulose partially digested by the C. bescii enzyme system shows some free CNCs and several larger bundles of cellulose fibrils in the process of fragmentation. at a strict 28 °C, and pH controlled at 4.8. The acid and base used for pH control was 2 M HCl and 2 M KOH, respectively. The cell culture was grown for 3 d, after which the entire culture broth was drained, filtered through nylon to remove all cell mass, and concentrated via tangential flow filtration using a 10 000 MWCO Vivaspin 20 diafiltration membrane from Sartorius. The concentrated broth was buffer exchanged into 50 mM sodium acetate buffer pH 6.5 and brought to a final volume of ∼200 mL. A complete description of the enzymatic components produced by T. reesei has been published previously.59 C. bescii Strain Growth Conditions and Exoproteome Isolation. C. bescii strain DSM6725 was obtained from Deutsche Sammlung von Mikroorganismen, Germany. This strain DSM6725 was chosen for this study as it is the most stable C. bescii strain for high production of cellulases in our laboratory. The culture was agitated (150 rpm) for 24 h at 75 °C in a 600 L tower-fermentation vessel with 500 L of the mineral medium60 containing 0.5 wt% crystalline cellulose (Avicel). The pH was maintained at pH 7.2 and gas phase was continuously flushed with N2/CO2 (80/20, v/v). The exoproteome was collected and concentrated using 10 000 MWCO Vivaspin 20 diafiltration membrane from Sartorius to 1.5 L. The exoproteome was buffer exchanged in 20 mM sodium acetate buffer pH 5.5, with 100 mM NaCl and 10 mM CaCl2 using the same diafiltration system mentioned above. Protein concentration was determined by the Bradford method. A complete description of the enzymatic components of the secretome of this strain has been reported previously.61 Cellulose Digestions for Nanocellulose Production. Digestions for nanocellulose production were performed in water due to the flocculation of nanocellulose in buffer. The experiment used 400 mg of SWKP in 40 mL of solution incubated with a total enzyme loading of 7.5 mg of protein per gram of glucan in 50 mL conical Falcon tubes. Incubations were performed at previously determined optimal temperatures for each enzyme system, i.e., 50 °C for the T. reesei system62 and 75 °C for the thermophilic C. bescii system.63 Samples were collected at 24, 48, 72, 96, and 120 h and consisted of a “kill series” by sacrificing individual tubes with aliquots taken for sugar analysis, imaging, and dynamic light scattering. Yields of NC were determined gravimetrically after differential centrifugation to separate NC from unconverted bulk cellulose. Samples collected at each time point were sonicated for 25 min with an amplitude setting of 75 on a Qsonic Q700 microtip sonicator. Following sonication, the suspension was centrifuged at 1600g for 10 min in a swinging bucket rotor. The supernatant was decanted into a new preweighed Falcon tube and the pellet was resuspended in 40 mL of deionized water. This was repeated

by C. bescii, which differs significantly from that of processive cellulases (Cel7A and Cel6A), the primary cellulolytic agents in fungal secretomes. This work also provides general proof of concept that the differing functionalities of various hydrolytic enzymes can affect the yields and characteristics of nanocellulose produced from enzymatic saccharification. Given the vast diversity of naturally occurring cellulases and emerging strategies for enzyme engineering, it is likely that many discoveries remain in this area of research. Furthermore, rational design of enzyme systems for controlled production of nanocellulose with tailored properties will benefit from improved fundamental understanding of mechanisms by which enzymes deconstruct cellulose and other biopolymers. We feel that the nanobiotechnology demonstrated here, as well as similar approaches by which biopolymeric materials may be produced, modified, and controlled by enzymatic systems, hold great potential for future biorefineries that seek to produce fuels, chemicals, and materials from renewable resources. Costeffective integration of biofuels process technology with biomaterials production strategies will also require new sensitivity studies to balance carbon flow between fuels and material products according to market considerations.

EXPERIMENTAL SECTION Compositional Characterization of the Raw Feedstock. The compositional analysis of the softwood bleached Kraft pulp (SBKP) obtained from International Paper (Memphis, TN) was conducted according to standard NREL Laboratory Analytical Procedures (LAPs).57 The compositional analysis of SBKP showed that it contained 79.2 ± 0.5% glucan, 15.3% xylan, and 1.2% lignin. T. reesei Production. The cell culture was streaked on a Potato Dextrose Agar plate and allowed to grow until a well-lawned plate of spores was achieved. A ∼ 0.5 cm plug was extracted from the plate and deposited into 1 L of liquid growth medium in a 2800 mL shake flask. The growth media consisted of Mandel’s Growth Media58 with 2% lactose as the carbon source and 0.5% tryptone added. The culture was grown at 28 °C with agitation for 3 d, after which the entire 1 L was transferred to 7 L of the same media in a bioreactor. The bioreactors were 14 L working volume vessels manufactured by New Brunswick and controlled via New Brunswick’s BioFlo3000 system. The total of 8 L was grown with mixing at 300 rpm via dual down-flow marine impellers, purged with 12 L/min of filtered air (1.5 VVM5VVM), kept 3106


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ACS Nano twice, yielding three supernatant tubes and one final pellet for each sample. Aliquots (500 μL) of supernatant were taken for imaging and dynamic light scattering from the final supernatant; whereas 100 μL samples of all three supernatants were taken for sugar analysis (described below) followed by centrifugation. The combined supernatants were lyophilized and NC yield was calculated from the resultant masses after correcting for free sugar content. To determine the progress of glucan conversion, 100 μL aliquots of the well-mixed slurries were taken at 24, 48, 72, 96, and 120 h. The samples were immediately diluted with 900 μL of deionized water, and the enzymes were inactivated by heating at 95 °C for 12 min. Samples were filtered through Pall Acrodisc nylon 0.2 μm syringe filters (Pall, Port Washington, NY) and refrigerated until HPLC analysis on an Agilent 1100 using a 300 mm × 7.8 mm BioRad Aminex HPX87H Ion Exclusion column maintained at 55 °C. The mobile phase for HPLC was 0.01 N sulfuric acid and the flow rate was 0.6 mL/min. The sample injection volume was 20 μL and the run time was 25 min. Glucan conversion was calculated by adding the total glucose and cellobiose yields (both glucose and cellobiose were converted to glucan equivalent) for each hydrolysis time point. Dynamic Light Scattering. Size distribution experiments were performed with a Zetasizer Nano-ZS (Malvern Instruments Ltd., Britain) at a detection angle of 173°. Samples were first dispersed in water to give 0.1 wt% suspensions. Measurements were carried out at 25 °C in triplicate for error analysis. Atomic Force Microscopy. Samples were diluted to 0.1 wt% solids and drop-cast onto freshly glow-discharged silicon substrates. The samples were then dried under vacuum. The dry samples were mounted at the Multi-Mode scanning probe microscope (SPM) with a NanoScope IV controller (Bruker, Santa Barbara, CA) utilized for all AFM measurements. A customized Nikon optical microscope was used to position scanning area of the sample. Images were obtained in tapping mode using etched silicon probes (TESP, Bruker) with an autotuned resonance frequency range of 250 to 300 kHz at a scan rate of 0.5 to 2 Hz. Images were analyzed with Nanoscope Analysis v1.2 software. Transmission Electron Microscopy. Samples were diluted to 0.1 wt% solids and drop-cast onto freshly glow-discharged carbon-coated copper TEM grids (VWR). After one min, the excess solution was removed and the grid was post-stained with 2 wt% aqueous uranyl acetate (Sigma-Aldrich) for 2 to 3 min. Excess solution was blotted away using filter paper and the grid was allowed to dry prior to imaging. Images were taken with a four mega-pixel GatanUltraScan 1000 camera (Gatan, Pleasanton, CA) on a FEI Tecnai G2 20 Twin 200 kV LaB6 TEM (FEI, Hillsboro, OR) using Digital Micrograph image capture software.

ACKNOWLEDGMENTS Funding for this work was provided by the National Renewable Energy Laboratory’s Laboratory Directed Research and Development (LDRD) program. REFERENCES (1) Wang, Q.; Zhu, J.; Reiner, R.; Verrill, S.; Baxa, U.; McNeil, S. Approaching Zero Cellulose Loss in Cellulose Nanocrystal (Cnc) Production: Recovery and Characterization of Cellulosic Solid Residues (Csr) and Cnc. Cellulose 2012, 19, 2033−2047. (2) Meng, Y.; Wu, Q.; Young, T. M.; Huang, B.; Wang, S.; Li, Y. Analyzing Three-Dimensional Structure and Geometrical Shape of Individual Cellulose Nanocrystal from Switchgrass. Polym. Compos. 2015, DOI: 10.1002/pc.23819. (3) Feng, X.; Meng, X.; Zhao, J.; Miao, M.; Shi, L.; Zhang, S.; Fang, J. Extraction and Preparation of Cellulose Nanocrystals from Dealginate Kelp Residue: Structures and Morphological Characterization. Cellulose 2015, 22, 1763−1772. (4) Moon, R. J.; Martini, A.; Nairn, J.; Simonsen, J.; Youngblood, J. Cellulose Nanomaterials Review: Structure, Properties and Nanocomposites. Chem. Soc. Rev. 2011, 40, 3941−3994. (5) Brinchi, L.; Cotana, F.; Fortunati, E.; Kenny, J. Production of Nanocrystalline Cellulose from Lignocellulosic Biomass: Technology and Applications. Carbohydr. Polym. 2013, 94, 154−169. (6) Zhu, H.; Luo, W.; Ciesielski, P. N.; Fang, Z.; Zhu, J.; Henriksson, G.; Himmel, M. E.; Hu, L. Wood-Derived Materials for Green Electronics, Biological Devices, and Energy Applications. Chem. Rev. 2016, 116, 9305−9374. (7) Nechyporchuk, O.; Belgacem, M. N.; Bras, J. Production of Cellulose Nanofibrils: A Review of Recent Advances. Ind. Crops Prod. 2016, 93, 2−25. (8) Somerville, C. Cellulose Synthesis in Higher Plants. Annu. Rev. Cell Dev. Biol. 2006, 22, 53−78. (9) Wakelin, J. H.; Virgin, H. S.; Crystal, E. Development and Comparison of Two X-Ray Methods for Determining the Crystallinity of Cotton Cellulose. J. Appl. Phys. 1959, 30, 1654−1662. (10) Schenzel, K.; Fischer, S.; Brendler, E. New Method for Determining the Degree of Cellulose I Crystallinity by Means of Ft Raman Spectroscopy. Cellulose 2005, 12, 223−231. (11) Park, S.; Johnson, D. K.; Ishizawa, C. I.; Parilla, P. A.; Davis, M. F. Measuring the Crystallinity Index of Cellulose by Solid State 13c Nuclear Magnetic Resonance. Cellulose 2009, 16, 641−647. (12) Atalla, R.; Crowley, M.; Himmel, M.; Atalla, R. Irreversible Transformations of Native Celluloses, Upon Exposure to Elevated Temperatures. Carbohydr. Polym. 2014, 100, 2−8. (13) Chen, L. H.; Wang, Q. Q.; Hirth, K.; Baez, C.; Agarwal, U. P.; Zhu, J. Y. Tailoring the Yield and Characteristics of Wood Cellulose Nanocrystals (Cnc) Using Concentrated Acid Hydrolysis. Cellulose 2015, 22 (3), 1753−1762. (14) Wang, Q. Q.; Zhao, X. B.; Zhu, J. Y. Kinetics of Strong Acid Hydrolysis of a Bleached Kraft Pulp for Producing Cellulose Nanocrystals (Cncs). Ind. Eng. Chem. Res. 2014, 53, 11007−11014. (15) Zimmermann, T.; Bordeanu, N.; Strub, E. Properties of Nanofibrillated Cellulose from Different Raw Materials and Its Reinforcement Potential. Carbohydr. Polym. 2010, 79, 1086−1093. (16) Qing, Y.; Sabo, R.; Zhu, J. Y.; Agarwal, U.; Cai, Z.; Wu, Y. A Comparative Study of Cellulose Nanofibrils Disintegrated Via Multiple Processing Approaches. Carbohydr. Polym. 2013, 97, 226− 234. (17) Syverud, K.; Chinga-Carrasco, G.; Toledo, J.; Toledo, P. G. A Comparative Study of Eucalyptus and Pinus Radiata Pulp Fibres as Raw Materials for Production of Cellulose Nanofibrils. Carbohydr. Polym. 2011, 84, 1033−1038. (18) Brinchi, L.; Cotana, F.; Fortunati, E.; Kenny, J. M. Production of Nanocrystalline Cellulose from Lignocellulosic Biomass: Technology and Applications. Carbohydr. Polym. 2013, 94, 154−169.

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. ORCID

Peter N. Ciesielski: 0000-0003-3360-9210 Author Contributions †

J.M.Y. and R.Z. contributed equally to this work.

Author Contributions

Project was initially conceived by P.N.C., M.E.H., S.R.D., J.M.Y., A.M., and Y.J.B. Fermentations for enzyme production were performed by T.V.W. Enzymatic saccharification was performed by J.M.Y., A.M., and R.Z. Dynamic light scattering was performed by R.Z. and J.M.Y. Electron and scanning probe microscopies were performed by R.Z. and P.N.C. Article text and figures were prepared by P.N.C. with editorial assistance from all authors. Notes

The authors declare no competing financial interest. 3107


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ACS Nano

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Multifunctional Cellulolytic Enzymes Outperform Processive Fungal

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