Microfluidic Glycosyl Hydrolase Screening for Biomass-to-Biofuel

Oct 22, 2010 - We consider the separation of labeled glucose and xylose molecules as the limiting case. Glucose and xylose are difficult to separate s...
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Anal. Chem. 2010, 82, 9513–9520

Microfluidic Glycosyl Hydrolase Screening for Biomass-to-Biofuel Conversion Rajiv Bharadwaj,*,†,‡ Zhiwei Chen,‡,§ Supratim Datta,‡,§ Bradley M. Holmes,‡,§ Rajat Sapra,‡,§ Blake A. Simmons,‡,§ Paul D. Adams,†,|,⊥ and Anup K. Singh†,‡ Technology and Deconstruction Divisions, The Joint BioEnergy Institute, Emeryville, California 94608, Sandia National Laboratories, Livermore, California 94551, Department of Bioengineering, University of California, Berkeley, California 94720, and Lawrence Berkeley National Laboratory, Berkeley, California 94720, United States The hydrolysis of biomass to fermentable sugars using glycosyl hydrolases such as cellulases and hemicellulases is a limiting and costly step in the conversion of biomass to biofuels. Enhancement in hydrolysis efficiency is necessary and requires improvement in both enzymes and processing strategies. Advances in both areas in turn strongly depend on the progress in developing highthroughput assays to rapidly and quantitatively screen a large number of enzymes and processing conditions. For example, the characterization of various cellodextrins and xylooligomers produced during the time course of saccharification is important in the design of suitable reactors, enzyme cocktail compositions, and biomass pretreatment schemes. We have developed a microfluidicchip-based assay for rapid and precise characterization of glycans and xylans resulting from biomass hydrolysis. The technique enables multiplexed separation of soluble cellodextrins and xylose oligomers in around 1 min (10fold faster than HPLC). The microfluidic device was used to elucidate the mode of action of Tm_Cel5A, a novel cellulase from hyperthermophile Thermotoga maritima. The results demonstrate that the cellulase is active at 80 °C and effectively hydrolyzes cellodextrins and ionicliquid-pretreated switchgrass and Avicel to glucose, cellobiose, and cellotriose. The proposed microscale approach is ideal for quantitative large-scale screening of enzyme libraries for biomass hydrolysis, for development of energy feedstocks, and for polysaccharide sequencing. Lignocellulosic (LC) biomass is an abundant and potentially carbon-neutral resource for production of biofuels and chemicals.1,2 LC biomass is made of three major componentsscellulose * Corresponding author. E-mail: [email protected]. Phone: (650) 804-5359. Fax: (510) 495-5225. † Technology Division, The Joint BioEnergy Institute. ‡ Sandia National Laboratories. § Deconstruction Division, The Joint BioEnergy Institute. | University of California. ⊥ Lawrence Berkeley National Laboratory. (1) Ragauskas, A. J.; Williams, C. K.; Davison, B. H.; Britovsek, G.; Cairney, J.; Eckert, C. A.; Frederick, W. J., Jr.; Hallett, J. P.; Leak, D. J.; Liotta, C. L.; Mielenz, J. R.; Murphy, R.; Templer, R.; Tschaplinski, T. Science 2006, 311, 484. (2) Tilman, D.; Hill, J.; Lehman, C. Science 2006, 314, 1598. 10.1021/ac102243f  2010 American Chemical Society Published on Web 10/22/2010

(35-50%), hemicellulose (20-35%), and lignin (10-25%).3,4 One of the key steps in the biomass-to-biofuel processes is the saccharification of biomass using glycosyl hydrolases (GHs), such as cellulases and hemicellulases, to generate fermentable monosaccharides. Cellulases hydrolyze cellulose into D-glucose, which is the primary fermentable sugar for biofuel production. The hemicellulases convert the hemicellulose polysaccharide mainly to D-xylose, which is the second most abundant sugar. D-Xylose can also be fermented to produce biofuels, although with less efficiency than glucose currently.4,5 Cellulose is a linear condensation polymer that consists of β-1,4linked D-glucose units with a degree of polymerization (DP) from 100 to 20000.6 In the plant cell walls, the polymeric cellulose chains are held together by extensive networks of hydrogen bonds and van der Waals forces. The cellulosic component of LC is highly crystalline and recalcitrant to enzymatic saccharification. The hydrolysis involves synergistic action by endoglucanases (EC 3.2.1.4), cellobiohydrolases (EC 3.2.1.91), and β-glucosidases (EC 3.2.1.21) that convert cellulose into D-glucose. Endoglucanases randomly hydrolyze accessible intramolecular β-1,4-glucosidic bonds of cellulose chains and produce new chain ends. The exoglucanases or cellobiohydrolases (CBHs) cleave cellulose chains at the ends to release soluble cellobiose or glucose. Finally, β-glucosidases hydrolyze cellobiose to glucose. Similarly, endoxylanase and β-xylosidase break down the hemicellulose component of biomass during the saccharification process.7 The cost of cellulases and hemicellulases is a major contributor to the overall process economics, and significant improvements in specific activity on industrially relevant feedstocks is necessary to develop economically feasible biorefineries.6,8 Improvements in the specific activity can be achieved through enzyme engineering techniques such as rational design or directed evolution.9 These enzyme engineering activities result in vast libraries of enzymes, and thus, high-throughput (HTP) enzyme assays are needed for rapid and cost-effective selection of enzymes with the desired properties. (3) Lynd, L. R.; Weimer, P. J.; van Zyl, W. H.; Pretorius, I. S. Microbiol. Mol. Biol. Rev. 2002, 66, 506. (4) Alper, H.; Stephanopoulos, G. Nat. Rev. Microbiol. 2009, 7, 715. (5) Jeffries, T. W. Curr. Opin. Biotechnol. 2006, 17, 320. (6) Zhang, Y.-H.; Himmel, M.; Mielenz, J. Biotechnol. Adv. 2006, 24, 452. (7) Gao, D. H.; Chundawat, S. P. S.; Krishnan, C.; Balan, V.; Dale, B. E. Bioresour. Technol. 2010, 101, 2770. (8) Moreira, N. Sci. News 2005, 168, 218. (9) Heinzelman, P.; Snow, C. D.; Wu, I.; Nguyen, C.; Villalobos, A.; Govindarajan, S.; Minshull, J.; Arnold, F. H. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 5610.

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There are two main classes of conventional sugar assays. The first class is the colorimetric/fluorometric sugar assays such as the dinitrosalicylic acid (DNS) method, Nelson-Somogyi method, and enzymatic sugar assays.6,10 These assays are high-throughput but do not provide detailed information about the nature of the various carbohydrates present in the hydrolysate. For example, either the total reducing sugar concentration can be measured (e.g., DNS assay) or a single analyte can be quantified (e.g., glucose-hexokinase assay). The second class of assays uses chromatographic approaches such as HPLC and TLC to obtain precise information about the soluble oligosaccharides produced as a result of GH activity.6 In addition, there are few examples of enzymatic fingerprinting of plant carbohydrates using electrophoretic separations in polyacrylamide gels or glass capillaries.11-16 However, these methods are typically slow (∼10 min per assay) and difficult to implement for high-throughput screening of large cellulase libraries. In this study, we have developed a microfluidic capillary electrophoresis device for rapid, precise, and high-throughput characterization of plant oligomers for GH screening. In the next section, we develop a mathematical model to rationally select the system parameters for optimized separation of labeled oligomers. Next, we describe multiplexed separation of cellodextrins (soluble cellulose oligomers with DP up to 6) and xylose oligomers using the microfluidic device. Finally, we apply the technique to elucidate the mode of action of a novel thermostable endoglucanase, Tm_Cel5A, from a hyperthermophilic microorganism, Thermotoga maritima MSB8. Tm_Cel5A enzyme is highly thermostable and is an excellent candidate for use in the degradation of biomass at temperatures relevant to biorefinery processes. THEORY A vast majority of plant mono- and oligo-saccharides do not have readily ionizable functional groups. Moreover, the carbohydrates lack chromophoric and/or fluorogenic function. Therefore, preseparation derivatization with reagents that contain ionizable and fluorogenic/chromophoric groups is necessary for electrophoretic separation. Several carbohydrate tags or labels and associated conjugation protocols have been described in the literature.11-15 The size range of the available labels is between 200 and 800 Da, and the valence varies between -1 and -3. The molecular sizes of the label and the carbohydrate sample are often comparable. Hence, the choice of the labeling molecule can have a profound impact on the separation resolution and efficiency of the conjugated carbohydrates. In this section we derive a simple scaling-based mathematical model to enable rational design of the electrophoresis of labeled oligosaccharides. We develop the model by considering the separation of two labeled oligosaccharides. We assume that the electroosmotic flow (10) Sharrock, K. R. J. Biochem. Biophys. Methods 1988, 17, 81. (11) Jackson, P. Biochem. J. 1990, 270, 705. (12) Goubet, F.; Strom, A.; Quemener, B.; Stephens, E.; Williams, M. A. K.; Dupree, P. Glycobiology 2006, 16, 29. (13) O’Shea, M. G.; Samuel, M. S.; Konik, C. M.; Morell, M. K. Carbohydr. Res. 1998, 307, 1. (14) Barton, C. J.; Tailford, L. E.; Welchman, H.; Zhang, Z. N.; Gilbert, H. J.; Dupree, P.; Goubet, F. Planta 2006, 224, 163. (15) Dahlman, O.; Jacobs, A.; Liljenberg, A.; Olsson, A. I. J. Chromatogr., A 2000, 891, 157. (16) Khandurina, J.; Blum, D. L.; Stege, J. T.; Guttman, A. Electrophoresis 2004, 25, 2326.

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(EOF) is negligible to enable rapid separation of negatively charged labeled carbohydrates. The molecular weight of the label is mL, and those of the unconjugated oligosaccharides are m1 and m2. The dependence of the electrophoretic mobility on the molecular size of the carbohydrate analytes and the properties of the separation matrix is very complex, and quantitative relations are not readily available in the literature.17 However, the electrophoretic mobility can be estimated by balancing the electrostatic force with the Stokes drag: v)

q 6πµr

(1)

where q is the net charge, µ is the viscosity of the solvent, and r is the radius of gyration. The radius of gyration can be estimated from the molecular weight as r ≈ Mn

(2)

where M = m + mL is the molecular weight of the labeled saccharide. The exponent n is typically less than 1, and for a perfectly spherical molecule the exponent is expected to be 1/3. However, for nonspherical molecules the exponent is determined by fitting the above relation to the experimental data. Combining eqs 1 and 2 leads to v)C

( ) z Mn

(3)

where C is the proportionality constant and z is the valence of the label. Next, the separation resolution for a given separation length, L, is given by

R≈

t2 - t1 4σ

(4)

where t ) L/(Ev) is the arrival time and σ is the standard deviation of the peaks in the time domain. The standard deviation can be estimated by

σ≈



σinj2 + 2D1t (Ev1)2

(5)

where σinj is the injection width, D1 is the molecular diffusion coefficient, E is the electric field, and v1 is the electrophoretic mobility. The above relation assumes that molecular diffusion and the initial peak width are the main sources of peak broadening. Appropriate relations can be developed in cases where analyte absorption, advective dispersion, or thermal dispersion are important. We note that since the two analytes are similar in molecular size, the diffusion coefficient and the mobility are interchangeable for estimating the peak width. Substituting eqs 5 and 3 into eq 4 leads to R≈

L(f - 1) 4√

σinj2

+ 2D1LM1n/(ECz)

(17) Chiesa, C.; Horvath, C. J. Chromatogr., A 1993, 645, 337.

(6)

where L is the separation length and f ) (M2/M1)n is the ratio of the mobility of the two analytes. The above relation can be used to predict the separation performance of labeled carbohydrates as a function of various system parameters. EXPERIMENTAL SECTION Microchip System. An inverted epifluorescence microscope (Olympus IX71) equipped with 10× (Olympus, NA ) 0.4) and 40× (Olympus, NA ) 0.65) objectives was used for full-field imaging and for the capture of electropherograms. Illumination from a mercury lamp was spectrally filtered at absorption and emission wavelengths of 485 and 535 nm, respectively. Images were captured using a frame transfer back-illuminated electron-multiplying CCD camera (iXon+897, Andor Technology, South Windsor, CT) with a 512 × 512 CCD pixel array and 16-bit digitization. A photomultiplier tube (H5784-20, Hamamatsu Corp., Bridgewater, NJ) was used to capture the electropherograms. A low-fluorescence Borofloat glass microchip with a cross-channel geometry was used for all the experiments (Figure S-1, Supporting Information). The microchannel width is 80 µm, and the centerline depth of the channels is 30 µm. The channels have the characteristic shape of an isotropic wet etch. The lengths of the sample, sample waste, buffer, and buffer waste channels measured from the intersection point are 8.4, 17.34, 28.9, and 11.1 mm, respectively. The microchip was flushed with acidified poly(ethylene oxide) (PEO) solution to suppress EOF as described previously.18 After this surface treatment step, the microchip was flushed with HEPES (100 mM, pH 7.5) buffer for 5 min at a flow rate of 20 µL h-1 to remove PEO solution. A small amount of PEO (0.01% (w/v)) was added to the running buffer to ensure stability of the surface treatment. A custom high-voltage power supply was used to control the platinum electrode potential mated to the microchip reservoirs. A pinched-injection scheme was used for the separation experiments. The voltage was applied by using platinum electrodes, which were inserted into reagent reservoirs made from polystyrene pipet tips cut with a razor blade. The reservoirs (∼20 µL) were press-fit into the chip wells. For the sample-loading step (30 s duration) the potentials applied to the sample (S), sample waste (SW), buffer (B), and buffer waste (BW) reservoirs were -850, 0, -800, and -700 V, respectively. For the separation step (60 s duration), the potentials were switched to -950, -950, 0, and -2000 V, respectively (Figure S-1). Ionic Liquid Pretreatment. We used switchgrass (MPV2 cultivar provided by the laboratory of Dr. Ken Vogel) and microcrystalline cellulose (Avicel) as the representative insoluble substrates. Switchgrass was milled with a Thomas-Wiley minimill fitted with a 40-mesh screen (model 3383-L10, Arthur H. Thomas Co., Philadelphia, PA). The ionic liquid (IL) solvent was 1-ethyl3-methylimidazolium acetate ([C2mim][OAc]) (Sigma-Aldrich, St. Louis, MO). The pretreatment was performed by incubating a slurry of milled biomass and [C2mim][OAc] in an oven (Thelco laboratory oven, Thermo Fisher Scientific Inc., Waltham, MA) at 120 °C for 3 h. The pretreated cellulose was regenerated by slowly adding deionized water (antisolvent) into the stirred biomass/ [C2mim][OAc] solution as described previously.19 The supernatant containing ionic liquid was removed, and the precipitate was (18) Preisler, J.; Yeung, E. S. Anal. Chem. 1996, 68, 2885.

washed extensively to ensure that excess ionic liquid had been removed. The recovered product was lyophilized at -50 °C for 24 h before enzyme assays. Enzymatic Saccharification. The enzymatic saccharification experiments were carried out off-chip in Eppendorf tubes using a thermomixer (Thermomixer R, Eppendorf, New York). We used a commercial cellulase cocktail as well as a thermophilic endoglucanase, Tm_Cel5A, for the hydrolysis experiments. The commercial cellulase cocktail (Worthington Biochemical Corp., Lakewood, NJ) was used at a loading of 12.5 IU/mL and augmented with 5.1 IU/mL β-glucosidase (NS50010, Novozyme, Davis, CA). The reaction was performed in 50 mM sodium acetate buffer (pH 4.8) at 50 °C and 1000 rpm. The gene Tm_Cel5A (NP_229549) was synthesized and the codon optimized by Genscript Corp. (Piscataway, NJ). The gene was cloned into an expression vector, pCDF2 LIC/Ek, by a ligation-independent cloning method (Novagen). Tm_Cel5A was expressed in BL21 (DE3) using LB autoinduction media (Overnight Express Autoinduction System 1, Novagen) at 37 °C. Tm_Cel5A with a C-terminal His tag was purified by a Histrap FF 1 mL column as described previously.20 The hydrolysis reactions were performed at 80 °C since it corresponds to the optimal activity for Tm_Cel5A at pH 4.8. For the microfluidic electrophoretic analysis, the carbohydrates present in the hydrolysate were fluorescently labeled using 8-aminopyrene-1,3,6-trisulfonic acid (APTS) fluorophore via the reductive amination method reported previously.21 Briefly, the labeling reaction was performed by adding 3 µL of APTS (0.1 M solution) and 3 µL of sodium cyanoborohydride (1 M solution) to 5 µL of the hydrolysate. The reductive amination was performed in the dark at room temperature for 24 h. The reaction was quenched by adding 100 µL of DI water to the reaction solution. The total amount of reducing sugars for Avicel and switchgrass samples was also measured using the DNS assay with D-glucose as the standard.22 RESULTS AND DISCUSSION Microfluidic Analysis of Plant Oligosaccharides. The scaling-based model from the Theory section can be used to develop guidelines for rational selection of the labeling reagents and the separation parameters. We consider the separation of labeled glucose and xylose molecules as the limiting case. Glucose and xylose are difficult to separate since the molecular weight difference is small. Further, the relatively large molecular weight of the common fluorogenic labels makes the separations of the conjugated monosaccharides even more challenging. As shown in Figure 1, the presented model can be used to predict the separation performance as a function of the molecular weight and valence of the label for a fixed separation length. We used n ) 2/3 for the mathematical model. The numerical value of the exponent was confirmed using experimental data as discussed later in this section. The constant C in eq 3 was estimated by the measured APTS mobility. (19) Li, C.; Knierim, B.; Manisseri, C.; Arora, R.; Scheller, H. V.; Auer, M.; Vogel, K. P.; Simmons, B. A.; Singh, S. Bioresour. Technol. 2010, 101, 4900. (20) Datta, S.; Holmes, B.; Park, J. I.; Chen, Z.; Dibble, D. C.; Hadi, M.; Blanch, H.; Simmons, B.; Sapra, R. Green Chem. 2010, 12, 338. (21) Olajos, M.; Hajos, P.; Bonn, G. K.; Guttman, A. Anal. Chem. 2008, 80, 4241. (22) Miller, G. L. Anal. Chem. 1959, 31, 426.

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Figure 1. Model predictions of the separation resolution (A) and separation time (B) of labeled glucose and xylose molecules as a function of the charge and mass of the labeling reagent. The performance characteristics of commonly used labeling reagents are also indicated. Key: (I) 5-aminonaphthalene-2-sulfonic acid, (II) 7-aminonaphthalene-1,3-disulfonic acid, (III) 9-aminopyrene-1,4,6trisulfonic acid, (IV) 8-aminonaphthalene-1,3,6-trisulfonic acid. The parameters for the model predictions are D ) 5.5E-10 m2/s, σinj ) 170 µm, E ) 400 V/cm, and L ) 22 mm.

For a given valence, the resolution decreases rapidly with increasing molecular weight of the label. The trade-off between resolution and separation time can be quantitatively analyzed using our model. This information can be used to rationally select the fluorescent label for rapid and high-resolution separation. In practice, the sensitivity requirements and the availability of detection hardware also influence the choice of the fluorophore. In Figure 1, we consider the performance of four carbohydrate labels with different valence and molecular weight characteristics: 5-aminonaphthalene-2-sulfonic acid (ANSA), 7-aminonaphthalene1,3-disulfonic acid (ANDSA), 9-aminopyrene-1,4,6-trisulfonic acid (APTS), and 8-aminonaphthalene-1,3,6-trisulfonic acid (ANTS). As shown in Figure 1, all four labels provide satisfactory separation resolution (R > 1.25).23 However, on the basis of the speed of analysis, ANDSA and ANSA were eliminated as potential fluorophore labels. Both ANTS and APTS enable rapid (∼20s) separation, with ANTS being slightly faster. Finally, we chose APTS over ANTS as the label for the carbohydrate separations on the basis of the availability of the detection optics in our laboratory. The APTS-derivatized sugars have significant absorption at 488 nm compared to APTS itself. Although a laser light source was not required for our experiments, it has been shown that, by using a 488 nm argon ion laser for excitation and a narrow band filter at 520 nm for fluorescence emission, the APTS-derivatized sugars can be selectively detected while the signal from excess APTS reagent is significantly suppressed.24 In Figure 2a, the separation of cellulose and xylose oligosaccharides is described. The oligosaccharide sample contained equal concentrations (1.1 µM) of all analytes. To enable rapid separations, the EOF was suppressed using an acidified PEO dynamic (23) Jimidar, M.; Bourguignon, B.; Massart, D. L. J. Chromatogr., A 1996, 740, 109. (24) Vanhooren, V.; Laroy, W.; Libert, C.; Chen, C. Y. Biogerontology 2008, 9, 351.

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Figure 2. (a) Separation of a mixture of cellodextrins (C1-C6) and xylose oligomers (X1-X6) using microfluidic electrophoresis. The concentration of each oligosaccharide was 1.1 µM. The assay time includes a 30 s sample-loading step. (b) Relation between the measured electrophoretic mobility of APTS-derivatized oligosaccharides and the molecular weight. (c) Comparison of measured and predicted separation resolutions for xylose and glucose oligomers of different degrees of polymerization.

coating. The residual EOF mobility was measured (vEOF ) 1.90 × 10-9 m2/(V s)) using the current monitoring technique and was around an order of magnitude smaller than the electrophoretic mobility of APTS. The dynamic coating was stable for at least 50 consecutive separations on the basis of the reproducibility of the peak arrival times (4 times the transit time of DP6 from the sample well to the injection cross) and did not observe any change in the variability of the fluorescence signal of the various oligosaccharides. This suggests that the nonuniformity in the peak areas is likely due to the variability in the labeling efficiency and not due to injection bias. Differential labeling efficiencies have been documented in the literature for various monosaccharides and oligosaccharides.13,25-29 For example, Bui et al.25 observed that the peak areas (normalized to (25) Bui, A.; Kocsis, B.; Kilar, F. J. Biochem. Biophys. Methods 2008, 70, 1313.

that of mannose) of various monosaccharide-APTS conjugates varied between 0.06 and 1.72. O’Shea et al.13 compared the APTS labeling efficiency of maltooligosaccharides using radiolabeled synthetic maltooligosaccharides and found that maltose (88%) and glucose (95%) were labeled more efficiently than the oligosaccharides (80%). As a result of the variability of the labeling efficiency, the electropherograms presented in this study represent qualitative profiling of hydrolysis products. To obtain quantitative information from the electropherograms, it is necessary to generate accurate calibration curves that relate the peak area to the analyte concentration. The quantitative analysis is described later in the paper (Table 1). We note that, recently, Vanderschaeghe et al.28 have described a comprehensive strategy to optimize the APTS labeling for N-glycans by varying the incubation time, temperature, and concentration of APTS. Similar approaches could be undertaken to optimize the labeling yields for lignocellulosic oligosaccharides. In Figure 2b, the relation between the measured mobilities and molecular weight is summarized. The mobilities are successfully correlated to the molecular mass using eq 3 with n ) 2/3. A similar relation was experimentally determined for derivatized maltooligosaccharides17,30 as well as for small peptides and proteins.31,32 The correlation suggests that the frictional drag in eq 1 is proportional to the surface area of a sphere with a volume that is proportional to the molecular weight: 4 V ) πr3 ≈ MW 3 drag ≈ A ) 4πr2 ≈ (MW)2/3

(7)

In Figure 2c, the separation resolution between consecutive glucose and xylose oligomers is seen to increase as the degree of polymerization increases. The model predictions match closely the experimental data. The injection width in eq 6 was measured from the CCD images of the pinched injection-separation process (Figure S-1, Supporting Information). The molecular diffusion coefficient of APTS-xylose was used as a fitting parameter. The diffusion coefficient of the other oligosaccharides was estimated by assuming that D ≈ (MW)-1/3.33 The separation resolution initially increases linearly since for a given fluorophore the size difference between the oligomers increases with the degree of polymerization. The separation resolution starts to taper off with a further increase in DP due to the power-law nature of the relation between mobility and molecular weight. The model predictions compare favorably with the experimental data. The model slightly overpredicts the separation resolution for larger oligomers. The model does not account for surface adsorption (26) Chen, F. T. A.; Dobashi, T. S.; Evangelista, R. A. Glycobiology 1998, 8, 1045. (27) Evangelista, R. A.; Guttman, A.; Chen, F. T. A. Electrophoresis 1996, 17, 347. (28) Vanderschaeghe, D.; Szekrenyes, A.; Wenz, C.; Gassmann, M.; Naik, N.; Bynum, M.; Yin, H. F.; Delanghe, J.; Guttman, A.; Callewaert, N. Anal. Chem. 2010, 82, 7408. (29) Khandurina, J.; Anderson, A. A.; Olson, N. A.; Stege, J. T.; Guttman, A. Electrophoresis 2004, 25, 3122. (30) Chiesa, C.; Oneill, R. A. Electrophoresis 1994, 15, 1132. (31) Offord, R. E. Nature 1966, 211, 591. (32) Rickard, E. C.; Strohl, M. M.; Nielsen, R. G. Anal. Biochem. 1991, 197, 197. (33) Young, M. E.; Carroad, P. A.; Bell, R. L. Biotechnol. Bioeng. 1980, 22, 947.

Figure 3. Simultaneous detection of cellulolytic and xylolytic activities of the cellulase cocktail. A mixture of xylotetraose and cellotetraose was used as the substrate. The enzymatic reaction was carried out for 2 h at 50 °C. The total substrate loading was 1 mM.

effects that may be important for higher molecular weight oligomers. The surface absorption will lead to higher peak broadening and hence lower separation resolution. The optimized microfluidic assay can be used for rapid and multianalyte characterization and is an attractive analytical tool for multiplexed screening of substrate-cellulase interactions. For example, in Figure 3, the microfluidic assay was used to simultaneously measure the cellulolytic and xylolytic activities of a commercial cellulase cocktail. The cellulase cocktail hydrolyzes a mixture of cellotetraose and xylotetraose substrates into glucose, xylose, xylobiose, and xylotriose. A small amount of unhydrolyzed xylotetraose was also observed, which indicates that the cellulase cocktail has higher cellulolytic activity than xylolytic activity. In the next two sections, we provide two examples to demonstrate the utility of the proposed platform for the analysis of industrially relevant enzymes (thermophilic endoglucanase) and substrates (ionic-liquid-pretreated biomass). Mode of Action of Thermophilic Cellulase. In this section, we use the microfluidic device to analyze the mode of action of Tm_Cel5A, a thermostable cellulase, on both soluble and insoluble substrates. Tm_Cel5A, which belongs to family 5 of the glycosyl hydrolases (GH5), is an extremely stable enzyme among the endoacting glycosidases present in the hyperthermophilic organism T. maritima. The crystal structure of Tm_Cel5A has been recently solved and reported.34 However, the mode of action of the cellulase has not been studied in detail. We performed a series of experiments both to determine the substrate specificity and to qualitatively identify the fingerprint of the hydrolysis products (Figure 4). First, we investigated the hydrolysis of the six individual soluble cellodextrins. The hydrolysis of well-defined soluble substrates is a useful tool for understanding the mechanism of cellulolytic enzymes. The Tm_Cel5A enzyme was found to be inactive on cellobiose and cellotriose substrates. The enzyme readily hydrolyzed cellotetraose, cellopentaose, and cellohexaose, and the evolution of the hydrolysis products is summarized in Figure 4. Cellotetraose is hydrolyzed to cellobiose, cellotriose, and (34) Pereira, J. H.; Chen, Z.; McAndrew, R. P.; Sapra, R.; Chahabra, S. R.; Sale, K. L.; Simmons, B. A.; Adams, P. D. J. Struct. Biol., in press.

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cellotetraose and cellotriose. In addition to the impurities that are inherent to the oligosaccharide manufacturing process, the mechanical vortexing during the dissolution of higher DP cellodextrins may also lead to generation of degradation products. While these degradation products are not desirable for quantitative analyses, the presence of multiple substrates enables qualitative assessment of substrate preference. The data in Figure 4b suggest that the dominant first step is the hydrolysis to cellobiose and cellotriose: DP5 f DP2 + DP3

(9)

Small amounts of glucose and cellotetraose are also produced. We note that the enzyme preferentially acts on cellopentaose rather than on cellotetrose that is present in small quantity in the starting sample. The subsequent degradation of cellotetraose proceeds in accordance to eq 8. The cellohexaose hydrolysis is described in Figure 4c. Again, we observed cellopentaose and cellotetraose as the degradation products in the control sample. The cellulase rapidly hydrolyzes cellohexaose and the small amount of cellopentaose in the starting sample, the dominant reactions being DP6 f DP2 + DP4 V DP3 + DP3

Figure 4. Analysis of the time evolution of substrates and products during hydrolysis of cellotetraose (a), cellopentaose (b), and cellohexaose (c) using thermophilic enzyme Tm_Cel5A. The electropherograms for three representative time points are overlaid to highlight the mechanism of cellulase on cellodextrin substrates. APTS-labeled D-xylose was added as an internal standard to enable peak alignment and identification. The hydrolysis reaction was carried out for 1 h at 80 °C. The enzyme loading was 10 µg, and the substrate concentration was 4 mM.

glucose (Figure 4a and Table 1). This suggests that the dominant reaction steps are DP4 f DP1 + DP3 V DP2 + DP2

(8)

where DP stands for the degree of polymerization and the numerals refer to the number of glucose units in the oligomers. The hydrolysis of cellopentaose is described in Figure 4b. The control (no enzyme) case shows that the cellopentaose sample contains a small amount of degradation products, primarily 9518

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(10)

The data clearly show that initially there is accumulation of DP4 while DP5 and DP6 substrates are hydrolyzed, and subsequently, the DP4 substrate is hydrolyzed according to eq 8. The hydrolysis of cellotetraose is therefore the rate-limiting step for the complete hydrolysis of cellodextrins into cellobiose, cellotriose, and glucose. The major hydrolysis products of the Tm_Cel5A endoglucanase are cellobiose and cellotriose. This is in agreement with the published data on similar bacterial endoglucanases.35 Saccharification of Ionic-Liquid-Pretreated Biomass. Next, we consider the enzymatic hydrolysis of lignocellulosic feedstocks for generation of fermentable sugars for biofuel production (Figure 5a). The recalcitrance of plant cell walls to enzymatic hydrolysis is a significant challenge toward cost-effective generation of fermentable sugars for biofuel production. There are two main reasons for biomass recalcitrance. The crystallinity of the cellulosic substrates renders the substrates inaccessible to cellulases. In addition, hemicellulose, pectin, and lignin coat the cellulose fibril and generate a physical barrier that further decreases the accessibility of cellulose to cellulases. Therefore, a thermochemical pretreatment of biomass is necessary to improve the enzymatic saccharification efficiency. We employed an ionic-liquid-based pretreatment using [C2mim][OAc]. Ionic liquids are a promising class of green solvents that decrease the crystallinity of cellulose and increase the accessible surface area for enzymatic hydrolysis under milder reaction conditions than conventional pretreatment processes such as ammonia fiber expansion, steam explosion, and dilute acid.36-38 In Figure 5b, we consider the hydrolysis of Avicel and switchgrass. Avicel is a model heterogeneous substrate that consists of microcrystalline cellulose. Switchgrass is a lignocel(35) Chhabra, S. R.; Kelly, R. M. FEBS Lett. 2002, 531, 375. (36) Singh, S.; Simmons, B. A. Biotechnol. Bioeng. 2009, 68.

Figure 5. (a) Schematic of the biochemical process for converting biomass to fermentable sugars. (b) Hydrolysis of ionic-liquid-pretreated Avicel (A) and switchgrass substrates using Tm_Cel5A enzyme. The biomass loading was 1 mg, and the enzyme loading was 10 µg. The reaction was performed at 80 °C for 24 h.

lulosic nonfood bioenergy crop that can be grown on marginal lands with relatively low water and nutrient requirements.39 The main hydrolysis products of the pretreated Avicel and switchgrass are glucose, cellobiose, and cellotriose. The data also indicate the presence of a peak overlapping the DP3 peak. The overlapping peak is more pronounced for switchgrass than Avicel. As opposed to Avicel, which only consists of cellulose, switchgrass is a more complex substrate that consists of cellulose, hemicellulose, and lignin. We are currently using mass spectrometry techniques to further understand the mechanism of action of Tm_Cel5A on complex lignocellulosic feedstocks. However, the overall product profile of the insoluble IL-pretreated substrates is similar to that of the model soluble cellodextrin substrates. Small amounts of higher cellodextrins (DP > 3) are observed for switchgrass, which indicates incomplete hydrolysis. The contaminant peak between the glucose and cellobiose was also observed in the control (no enzyme) samples. Finally, in Table 1, we consider the quantitative aspects of the microfluidic assay. A series of standard curves (Figure S-2, Supporting Information) were used to obtain the analyte concentration from the peak areas in the electropherograms. The standard curves were obtained by labeling a mixture of pure cellodextrins at known concentrations in the range of 0.5-1 mM. The standard curves were found to be linear for the concentration range. The total reducing sugar concentration for Avicel, switchgrass, and cellotetraose obtained using the microfluidic assay was (37) Swatloski, R. P.; Spear, S. K.; Holbrey, J. D.; Rogers, R. D. J. Am. Chem. Soc. 2002, 124, 4974. (38) Zhao, H.; Jones Cecil, L.; Baker Gary, A.; Xia, S.; Olubajo, O.; Person Vernecia, N. J. Biotechnol. 2009, 139, 47. (39) Sanderson, M. A.; Adler, P. R.; Boateng, A. A.; Casler, M. D.; Sarath, G. Can. J. Plant Sci. 2006, 86, 1315.

Table 1. Concentration (mM) of the Hydrolysis Products of IL-Pretreated Switchgrass and Avicel Using Tm_Cel5A Cellulasea

glucose cellobiose cellotriose total reducing sugars DNS assay

Avicel

switchgrass

cellotetraose

0.14 ± 0.03 0.57 ± 0.08 0.18 ± 0.04 0.89 ± 0.15 0.84 ± 0.09

0.14 ± 0.01 0.56 ± 0.05 0.32 ± 0.05 1.02 ± 0.12 1.15 ± 0.08

0.32 ± 0.02 1.51 ± 0.18 0.27 ± 0.04 2.1 ± 0.24 2.38 ± 0.11

a Also shown for comparison are the hydrolysis data for 1 mM cellotetraose. The data represent an average over three replicates for each case. The standard deviation is used as an estimate of the error in the measurements.

found to be in close agreement with the traditional DNS assay results. Further, the microfluidic assay is at least 10-fold faster than HPLC-based carbohydrate analysis (Figure S-3, Supporting Information). Therefore, microfluidic electrophoresis is an attractive technique for rapid and quantitative analysis of multiple hydrolysis products of complex real-world biomass samples. CONCLUSIONS The conventional approaches for analyzing cellulolytic activities are either throughput limiting (e.g., HPLC) or nonspecific (e.g., DNS assay). To overcome these limitations, we have developed a microfluidic electrophoresis system for cellulase assays. Compared to conventional approaches, microfluidic electrophoresis is a robust, rapid, multianalyte, and quantitative approach for analysis of complex biomass samples for biofuel production. We have developed an experimentally validated model to predict the separation resolution of biomass carbohydrate oligomers as a function of various system parameters including the separation Analytical Chemistry, Vol. 82, No. 22, November 15, 2010

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length, electric field, and fluorophore label characteristics. We demonstrated a multiplexed separation of both cellodextrins and xylose oligomers in around 1 min. The presented system can be used for large-scale characterization and optimization of cellulase cocktail compositions for real-world lignocellulosic substrates. In addition, the proposed system can be used to characterize glycosyltransferases for understanding and engineering bioenergy feedstocks. ACKNOWLEDGMENT Switchgrass (MPV2) was kindly provided by Dr. Ken Vogel of the U.S. Department of Agriculture, Agricultural Research Service, Lincoln, NE. We gratefully thank April Wong for her assistance with the electrophoresis assays. Special thanks go to Ujvalla Gupta for the stimulating discussions and encouragement. This work was part of the Department of Energy Joint BioEnergy Institute (http://www.jbei.org) supported by the U.S. Department

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of Energy, Office of Science, Office of Biological and Environmental Research, through Contract DE-AC02-05CH11231 between the Lawrence Berkeley National Laboratory and the U.S. Department of Energy. Sandia is a multiprogram laboratory operated by Sandia Corp., a Lockheed Martin company, for the U.S. Department of Energy’s Nuclear Security Administration under Contract DE-AC04-94AL85000. SUPPORTING INFORMATION AVAILABLE Schematic of the microfluidic chip along with CCD images of the sample introduction and separation process, measured calibration data for quantitative analyses, and HPLC data. This material is available free of charge via the Internet at http://pubs.acs.org. Received for review August 26, 2010. Accepted October 11, 2010. AC102243F