The authors would like to thank intern Evelyn Von Neida who provided paramount insights regarding the preparation of biomass samples for both of the high-throughput pipelines discussed in this manuscript. Support for the development of this work and manuscript was provided by the BioEnergy Science Center. The BioEnergy Science Center is a U.S. Department of Energy Bioenergy Research Center supported by the Office of Biological and Environmental Research in the DOE Office of Science. The National Renewable Energy Laboratory (NREL) is a national laboratory of the US DOE Office of Energy Efficiency and Renewable Energy, operated for DOE by the Alliance for Sustainable Energy, LLC. This work was supported by the U.S. Department of Energy under Contract No. DE-AC36-08-GO28308 with the National Renewable Energy Laboratory.

1. High-throughput Determination of Glucose and Xylose Yields Following Enzymatic Saccharification9,29
<ol>
	<li>Sample Preparation (Grinding, De-starching, Extraction, Pretreatment)
	<ol>
		<li>Grind at least 300 mg of each biomass sample using a Wiley mill, such that the particles pass through a 20 mesh (850 &#181;m) screen. Transfer to anti-static zip-top bags (typically bar-coded) and record sample information to the barcode database.</li>
		<li>Add approximately 250 mg or more of the ground biomass from anti-static bag to a numbered (with pencil, do not use ink or marker) tea-bag, carefully roll up the teabags, being sure to fold the ends over the biomass to prevent loss during de-starching and extraction.</li>
		<li>Wrap the teabag closed using tin-coated copper wire. Record the teabag number for each barcoded sample.</li>
		<li>Prepare de-starching enzyme solution from commercial enzymes (typically, 0.25% (v/v) glucoamylase (~1,600 AGU/L) and 1.5% (v/v) alpha-amylase (~2,900 KNU-S/L) in 0.1 M sodium acetate (pH 5.0)). Prepare 16 ml per g of bulk biomass or 500 ml per 120 teabag samples. Note: Loadings capable of removing starch from ground biomass should be determined empirically by testing for starch after digestion with a range of enzyme loadings and ratios.</li>
		<li>In a plastic container, add 16 ml of de-starching enzyme solution per 1 g of bulk ground biomass. For a batch de-starching protocol, add 120 teabags to 500 ml of the buffer-enzyme solution.</li>
		<li>Incubate in a shaker at 55&#160;&#176;C for 24 (&#177; 4) hr, at 120 rpm to remove possible starch.</li>
		<li>After the de-starching incubation, rinse and soak the biomass in several liters of deionized water for 30 min. Repeat this process two additional times to remove buffer salts that can cause bumping (formation of gas bubbles in the solution that can result in a sudden and violent rise in the level of the solution, causing hot liquid ethanol to cascade out of the reaction vessel) during the extraction.</li>
		<li>Following the exhaustive rinsing of the de-starched biomass, place the teabags into the Soxhlet column for extraction. No thimble is required for this step.</li>
		<li>Set up the Soxhlet reflux, using 95% ethanol and extract the samples for 24 hr.</li>
		<li>Remove the teabags from the Soxhlet reactor and spread out in a single layer on a flat tray.</li>
		<li>Allow the samples to dry overnight at room temperature in a fume hood.</li>
		<li>Unroll the teabags and return the dried biomass to original barcoded anti-static bags. 
		Note: The anti-static bags are efficient at reducing static electricity in the biomass, improving the handling characteristics. If other storage options are used, additional biomass may be needed due to the biomass clinging to the side of the storage container.</li>
		<li>Transfer at least 50 mg of the dried biomass to a barcoded hopper (using more material is better for accurate dispensing). Scan barcodes of anti-static bags and the receiving hopper to ensure accurate sample tracking.</li>
		<li>Load hoppers onto solids weighing robot, paying close attention to the order of the samples in the racks. Load enough reactor plates to contain all the samples and select the dispensing protocol based on the number of plates used.</li>
		<li>Robotically weigh 5 mg of the dried samples (&#177; 0.3 mg) into acid-resistant stainless steel 96-well plates. Weigh out 3 replicates of each sample distributed around the plate to minimize any localized variations.</li>
		<li>Include 8 control biomass samples of a well-characterized biomass standard material in order to track assay performance. For multiple plates, the use of multiple standard biomass hoppers will minimize particle size drift caused by sieving in the hoppers over repeated dispensing cycles. 
		Note: In each plate, there should be 24 samples with 3 replicates, 4 blanks consisting of deionized water and enzyme, and 8 controls, using the previously characterized standard biomass. The 3 corner wells of each of the 4 corners of the plate are reserved for the sugar standards.</li>
		<li>Check blank and sugar standard wells for errant biomass particles and remove if present.</li>
		<li>Add 250 &#181;l of deionized water to each well, and seal the plates with silicone adhesive-backed polytetrafluoroethylene (PTFE) tape.</li>
		<li>Using a 1/8&#8221; soldering iron tip, pierce the PTFE tape at each steam port (117 total) for each plate. Use an empty plate stacked on sealed reactor plate as a guide and to restrict PTFE film movement.</li>
		<li>Clamp the plates tightly with 0.031&#8221; thick glass-reinforced PTFE gaskets (pre-punched with holes for steam ports) between plates and empty plates on top and bottom of stack. Pretreat the samples using a steam reactor set to 180&#160;&#176;C for 17.5 min, or other temperature/time combinations based on desired severity.</li>
		<li>Cool reactor plates to 50&#160;&#176;C by flooding with deionized water.</li>
	</ol>
	</li>
	<li>Enzymatic Saccharification
	<ol>
		<li>Prepare an enzymatic saccharification solution consisting of 8% (v/v) enzyme solution in 1.0 M sodium citrate, pH 5.0. Prepare 5 ml per reactor plate. Note: Dilution required should be determined based on activity and protein content of the specific stock enzyme solution.</li>
		<li>When the plates are cool, centrifuge them in a swinging bucket rotor at 1,500 x g for 20 min. Remove sealing film. Note: These plates are heavy and the centrifuge specifications should be checked for compatibility.</li>
		<li>Add 40 &#181;l of the 8% enzyme stock solution to each well (70 mg enzyme per g biomass).</li>
		<li>Reseal with new PTFE tape. Place sealed plate into magnetic plate clamp.</li>
		<li>Gently mix the samples by inversion (at least 15 times), and incubate at 50&#160;&#176;C for 70 hr.</li>
		<li>When the saccharification has concluded, mix by inversion and centrifuge the plates at 1,500 x g for 20 min.</li>
	</ol>
	</li>
	<li>Sugar Assay
	<ol>
		<li>Prepare a set of 6 glucose and xylose combined calibration standards in 0.014 M citrate buffer (pH 5.0) ranging from 0 to 0.750 mg/ml (0, 0.2, 0.3, 0.45, 0.65, and 0.75 mg/ml recommended) and 0 to 0.600 mg/ml (0, 0.1, 0.2, 0.35, 0.45, and 0.6 mg/ml recommended) for glucose and xylose, respectively.</li>
		<li>Prepare the glucose oxidase/peroxidase (GOPOD) and xylose dehydrogenase (XDH) reagents according to instructions in the kit.</li>
		<li>Using a pipette, remove 200 &#181;l of liquid from the 4 corner wells and edge wells adjacent to the corner wells (12 total wells) of the reactor plate.</li>
		<li>Using a pipette, dispense 180 &#181;l deionized water to each well of a 96-well polystyrene flat-bottom dilution plate. Do not add water to the 12 sugar standard and corner wells (remove those tips if using a 96-channel head).</li>
		<li>Using a pipette, dispense 180 &#181;l GOPOD reagent to each well of the glucose assay plate.</li>
		<li>Using a pipette, dispense 180 &#181;l XDH reagent to each well of the xylose assay plate.</li>
		<li>Using a pipette, transfer 20 &#181;l hydrolysate aliquots from the 96-well reactor plate to the dilution plate. Pipet from the upper section of the wells to avoid biomass solids and residual liquid in corner wells. Mix by trituration for at least 10 cycles.</li>
		<li>Using a pipette, transfer 110 &#181;l of sugar standards, in duplicate, to the corner wells of the dilution plate.</li>
		<li>Using a pipette, transfer 20 &#181;l aliquots from the dilution plate to the glucose and xylose assay plates. Mix by trituration.</li>
		<li>Incubate the glucose and xylose assay plates at room temperature for 30 min. Carefully break any surface bubbles before reading. A heat gun briefly passed over the surface of the plate works well.</li>
		<li>Using an ultraviolet/visible 96-well plate reader, set the measured wavelength to 510 nm and record the absorbance against a reagent blank. This measurement monitors the formation of quinonimine, which is proportional to glucose concentration. The glucose concentration is calculated from the calibration curve based on the calibration standards prepared in 1.3.1.</li>
		<li>Using an ultraviolet/visible 96-well plate reader, set the measured wavelength to 340 nm and record the absorbance. This measurement monitors the reduction of NAD+ to NADH, which is proportional to xylose concentration. The xylose concentration is calculated from the calibration curve based on the calibration standards prepared in 1.3.1.</li>
	</ol>
	</li>
</ol>
2. High-throughput Determination of Total Lignin and Lignin Monomeric Content Using pyMBMS 28
<ol>
	<li>Sample Preparation
	<ol>
		<li>Grind and extract the biomass using the methods described in steps 1.1.1, and 1.1.6-1.1.8. This includes grinding and extracting of a set of standards to use as experimental controls. 
		Note: The pyMBMS measurements are not particle-size dependent, so if only using the pyMBMS protocol, biomass preparation should be carried out to allow the sample to fit in the sample holder, &#60;4 mm.</li>
		<li>Using a small spatula dispense approximately 4 mg (3 to 5 mg preferred) of the prepared biomass into an 80 &#181;l stainless steel cup designed for the auto-sampler.</li>
		<li>Ensure that at least 10% of the samples being analyzed are control standards such as those available from the National Institute of Standards and Technology (sugarcane bagasse-8491; poplar-8492; pine-8493; or wheat straw-8494). The standard can also be any species analogous to the samples to be analyzed that have already been characterized using standard methods.</li>
		<li>Randomly load the samples into the auto-sampler cups using tweezers to avoid bias due to possible spectrometer drift over time. Note: A typical randomization of the samples would include the measurement of all samples once, followed by a re-randomization of the order of the samples for a duplicate measurement. Randomization programs are available online.</li>
		<li>Using a standard hole-punch, manually produce glass filter discs from type A/D glass fiber sheets, with no binder. Hold the round glass fiber filter with tweezers, center it over the sample cup, and push into the sample using a 3.5 mm Allen wrench to confine the material in each cup during the experiment.</li>
	</ol>
	</li>
	<li>Instrumental Protocol
	<ol>
		<li>Calibrate the mass spectrometer using a known standard that has peak intensities over the entire range of compounds that may exist for the experimental samples. For typical biomass samples, use perfluorotributylamine (PFTBA).</li>
		<li>Set the helium carrier gas flow rate to 0.9 L/min using a gas flow meter.</li>
		<li>Set the auto-sampler furnace to a pyrolysis temperature of 500&#160;&#176;C and the interface temperature to 350&#160;&#176;C using the auto-sampler software. Note: A 1/8&#8221; stainless steel heated transfer line wrapped in heat tape that connects the auto-sampler to the mass spectrometer is controlled using a heat controller at 250&#160;&#176;C.</li>
		<li>Begin data acquisition on the mass spectrometer and wait at least 60 sec to obtain sufficient data for background spectra collection.</li>
		<li>Start the automated auto-sampler method with the specifications from 2.2.2. Note: The auto-sampler drops each sample individually into the auto-sampler furnace. The total data acquisition time is approximately 1.5 min; however, pyrolysis of a typical 4 mg sample is complete after 30 sec.</li>
		<li>Record total ion content (TIC) of each sample using mass spectrometer software at 0.5 sec scan rate. Record intensities between m/z 30 to 450. 
		Note: Typical biomass compounds use soft ionization at 17 eV. The instrument can record larger intervals from m/z 1 to 1,000; however, scan rate will be limited by computer CPU power.</li>
		<li>Remove background from the spectra using the manual enhance feature in the software. Note: A 60 scan portion of the baseline at the beginning of data collection is used to calculate an average background value. This average background spectra is removed from the experimental sample spectra automatically in the mass spectrometer software.</li>
		<li>Import the single column text file created by the mass spectrometer software containing the spectral data for each sample, into a database program and combine all the samples into one database. Add any applicable metadata to the spreadsheet. Import the formatted data (spreadsheet/CSV file) into a statistical software package and mean normalize the spectra to account for variation in the pyrolyzed sample masses.</li>
		<li>Use a statistical software package to perform a principal component analysis (PCA) using the spectral data to analyze grouping of the replicated standard samples used in the measurements, as well as to evaluate which peaks are integral to the classification of chemical compounds in the loadings plot28. 
		Note: PCA groups the samples based on the similarity of their spectra, and allows a check of the standards to gauge experimental error due to instrument drift during the run.</li>
		<li>To calculate the lignin syringyl (S)/guaiacyl (G) ratio, sum the areas of the S peaks (m/z = 154, 167, 168, 182, 194, 208, and 210) and divide by the sum of G peaks at 124, 137, 138, 150, 164, and 178.</li>
		<li>To calculate the total lignin content, sum the lignin peaks with m/z = 120, 124, 137, 138, 150, 152, 154, 164, 167, 178, 180, 182, 194, and 210.</li>
		<li>Calculate a correction factor for scaling the pyMBMS measurement to a standard method for estimating total lignin, such as Klason lignin. Divide the Klason lignin value of the individual standard by the total lignin content measured for that standard sample using pyMBMS.</li>
		<li>Apply this correction factor to all like-species in the data set. Repeat for each type of biomass analyzed. Note: The correction factor can vary significantly based on the S/G of the biomass analyzed.</li>
	</ol>
	</li>
</ol>

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R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ch.+12.+High-Throughput+Screening+of+Plant+Cell-Wall+Composition+Using+Pyrolysis+Molecular+Beam+Mass+Spectroscopy.">Ch. 12. High-Throughput Screening of Plant Cell-Wall Composition Using Pyrolysis Molecular Beam Mass Spectroscopy.</a> Biofuels: Methods and Protocols. 581, 169-183 (2009).</li><li>Decker, S., Himmel, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ch.+17.+Reducing+the+effect+of+variable+starch+levels+in+biomass+recalcitrance+screening).">Ch. 17. Reducing the effect of variable starch levels in biomass recalcitrance screening).</a> Biomass Conversion. 908, 181-195 (2012).</li><li>Evans, R. J., Milne, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+characterization+of+the+pyrolysis+of+biomass.">Molecular characterization of the pyrolysis of biomass.</a> Energy Fuels. 1 (2), 123-137 (1987).</li></ol>

The authors have nothing to disclose.

The key sample preparation steps for obtaining accurate and reproducible data when conducting high-throughput screening experiments are as follows:
Sugar Release Assay:
In general, samples are prepared in lots ranging from a few dozen to several thousand at a time. Each major step is typically carried out for all samples prior to moving forward in order to minimize variations in preparation between samples. De-starching was not originally part of the protocol and ca...

As the global supply of non-renewable fuels and their associated products declines, scientists have been challenged to create similar fuels and chemicals from plant-derived sources1. A key aspect of this work is determining which species of plants may be suitable for the production of biofuels and biomaterials2,3. Typically, these feedstocks are evaluated for lignin, cellulose, and hemicellulose content; as well as their susceptibility to deconstruction (recalcitrance) through thermal, mechanical, and/or chemical pretreatment with or without subsequent enzyme saccharification. More detailed analyses are used to determine the specific composition of the lignin and hemicellulose fractions as well as optimal enzyme activities needed. Transgenic modifications of plants that do not intrinsically possess ideal traits for biochemical or thermochemical conversion to desired commodities have provided researchers with a greatly expanded source of potential feedstocks4. The standard analytical methods for quantifying the chemical traits of a plant, while quite useful for small sample sets, are unsuited for the rapid screening of hundreds or thousands of samples5-7. The HTP methods described herein have been developed to rapidly and efficiently evaluate large numbers of biomass variants for changes in cell wall recalcitrance to thermochemical and/or enzymatic degradation.
It is critical to understand that the HTP screening assays described herein have not been designed to maximize conversion or yield. The objective is to determine relative differences in the intrinsic recalcitrance of related biomass samples. As a result, many of the analysis steps differ from the &#8220;typical&#8221; biomass conversion assays, where the objective is to obtain maximum conversion rate or extent. For example, lower pretreatment severities and shorter enzyme hydrolysis times are used to maximize differences between samples. In most cases, relatively high enzyme loadings are used to reduce differences due to experimental variation in enzyme activity, which could skew the results significantly.
Rapid techniques for determining the composition of plant cell-walls and the monomeric sugars liberated following enzymatic saccharification include robotics, customized, thermochemically compatible 96-well plates, and modifications of standard laboratory methods8-11 and instrumental protocols, such as vibrational spectroscopy (infrared (IR), near-infrared (NIR), or Raman) and nuclear magnetic resonance (NMR)12-17. These methodologies are key to isolating feedstocks with high cellulose or low lignin contents, or those expected to yield the highest glucose, xylose, ethanol, etc. These methods have enabled downscaled analyses that employ smaller quantities of biomass and consumables, leading to reductions in experimental expense18. Another feature of this methodological approach is that various experimental conditions can be rapidly, and in some cases simultaneously, evaluated. For example, a variety of different pretreatment strategies or enzyme cocktails can be tested, allowing the most optimal experimental parameters to be quickly identified and employed. Popular feedstocks, such as corn stover9, poplar8,10, sugarcane bagasse8, and switchgrass8 have been successfully evaluated using these HTP methods.
Total lignin and lignin monomeric composition are also commonly quantified biomass traits. Reductions in lignin content have been shown to increase the enzymatic digestibility of polysaccharides19,20. The role that the lignin monomeric ratio (often reported as syringyl/guaiacyl (S/G) content) plays in the deconstruction of the plant cell wall is still under investigation. Some reports have indicated that reductions in the S/G ratio led to increased glucose yields following hydrolysis21, while other studies unveil the opposite trend19,22. High throughput methods for evaluating lignin and its monomers include vibrational spectroscopy (IR, NIR, and Raman23-26) coupled with multivariate analysis, and pyrolysis molecular beam mass spectrometry (pyMBMS)27,28.
When developing HTP methods for screening biomass, several integral considerations need to be kept in mind. One key aspect is the complexity of the method. What is the required skill level for the technique? Chemometric analyses, for example, require specific skills for constructing, evaluating, and maintaining predictive models. The standard methods exhibit undesirable preparatory or data analysis steps or employ toxic reagents. Development of the models is an ongoing process where new data is incorporated into the model over time to increase the model&#8217;s robustness. Another consideration is the cost-savings and decreased experimental analysis times of the proposed high-throughput methods. If the method is quite rapid, but very costly, it may not be a feasible technique for many labs to adopt. The methods illustrated in this manuscript are variants of standardized techniques, modified to amplify the throughput capabilities. These protocols quantitatively measure the biomass traits of interest without necessitating the development of predictive models. This is a key attribute of these techniques, since predictive methods, while exhibiting strong correlations with the standard analyses used to develop the models, are not as accurate as actually measuring the quantity of interest for the samples. Whereas the methods used are essentially scaled down versions of standard bench-scale analytical methods, accuracy and precision are traded for speed and throughput. Mostly, this outcome is due to higher errors in small volume pipetting and weighing; as well as increased sample heterogeneity as sample size is decreased. While large sample sets can be screened and compared, great care must be exercised when making comparisons between separate campaigns and to bench-scale results.
The most time-consuming steps involve the physical manipulation of the biomass. Grinding samples may take several min per sample, including cleaning out the mill between samples. Manually loading, unloading, and cleaning hoppers and filling and emptying tea bags and sample bags is also very labor intensive. While each step may take a minute or more, doing thousands of samples may take many hours or even days. The robots can load a typical reactor plate with biomass in about 3 to 4 hr or 6 to 8 plates day-1 robot-1. This situation depends on the precision parameters used as well as the type and amount of biomass to be tested. Filling reactor plates with water, dilute acid, or enzyme is quickly done using a liquid handling robot. Pretreatment of a plate stack (1 to 20 reactor plates) takes between 1 and 3 hr when assembly, cool down, and disassembly is included. Enzyme hydrolysis takes 3 days and the sugar analysis requires about 1 hr of prep time plus 10 min per reactor plate to complete the assay and read the results. A weekly schedule of set pretreatment and analysis days accommodates a reasonable work schedule, minimizing odd-hour and weekend efforts for the human component of the assay and allows for processing ~800 to 1,000 samples per week on an ongoing basis. The maximum throughput depends on several factors, mainly how much hardware (robots, reactors plates, etc.) and how much &#8220;software&#8221; (i.e., staffing) are available to do the manual work. The practical upper limit is 2,500 to 3,000 samples/week; however, that output requires 7 day-a-week operation and multiple student interns and technicians. In comparison, 3,000 samples by HPLC would require approximately 125 days of sample analysis plus the additional labor of manually weighing samples into reactors and filtering samples prior to analysis.

<table><tbody><tr><td>Biomek FX Automated Workstation</td><td>Beckman Coulter</td><td>Biomek FX</td><td>Automated Liquid Handler</td></tr><tr><td>Wiley mill</td><td>Thomas Scientific</td><td>3375E15 (Model 4), or 3383L20 (Mini-mill)</td><td></td></tr><tr><td>anti-static bags</td><td>Minigrip*</td><td>MGST4P02503</td><td>2.5x3&quot;, multiple suppliers available</td></tr><tr><td>tin-coated copper wire</td><td>McMaster-Carr</td><td>8871K84</td><td>0.016&quot; diameter, bend-and-stay wire</td></tr><tr><td>tea-bags</td><td>Herbco</td><td>press n&#039; brew teabags</td><td>3.5x5 inches</td></tr><tr><td>gluco-amylase</td><td>Novozymes</td><td>Spirizyme Fuel&amp;nbsp;</td><td></td></tr><tr><td>alpha-amylase</td><td>Novozymes</td><td>Liquozyme SC DS</td><td></td></tr><tr><td>sodium acetate trihydrate</td><td>any chemical supplier</td><td></td><td>reagent grade</td></tr><tr><td>acetic acid</td><td>any chemical supplier</td><td></td><td>reagent grade</td></tr><tr><td>190 proof (95%) ethanol</td><td>any chemical supplier</td><td></td><td>reagent grade</td></tr><tr><td>hoppers</td><td>Freeslate</td><td></td><td></td></tr><tr><td>96-well C-276 Hastelloy plates</td><td>Aspen Machining (Lafayette, Colorado)</td><td>N/A (custom built)</td><td></td></tr><tr><td>1/8&amp;rdquo; soldering iron tip</td><td>Sears</td><td></td><td></td></tr><tr><td>silicone-adhesive backed Teflon tape</td><td>3M</td><td>5180</td><td>3&quot; wide (36-yard rolls)</td></tr><tr><td>enzyme solution</td><td>Novozymes</td><td></td><td>Cellic CTec2</td></tr><tr><td>citric acid monohydrate</td><td>any chemical supplier</td><td></td><td></td></tr><tr><td>trisodium citrate dihydrate</td><td>any chemical supplier</td><td></td><td></td></tr><tr><td>disposable, polystyrene 96-well plates</td><td>Greiner Bio-One</td><td>655101</td><td>or equivalent; multiple suppliers available</td></tr><tr><td>glucose oxidase/peroxidase&amp;nbsp;</td><td>Megazyme</td><td>K-Gluc</td><td>Megazyme D-glucose assay kit</td></tr><tr><td>xylose dehydrogenase</td><td>Megazyme</td><td>K-Xylose</td><td>Megazyme D-xylose assay kit</td></tr><tr><td>glucose standard solution</td><td>Megazyme</td><td>K-Gluc</td><td>Megazyme D-glucose assay kit</td></tr><tr><td>xylose standard solution</td><td>Megazyme</td><td>K-Xylose</td><td>Megazyme D-xylose assay kit</td></tr><tr><td>stainless steel sample cups</td><td>Frontier Laboratories</td><td>PY1-EC80F</td><td></td></tr><tr><td>glass fiber sheets</td><td>Pall</td><td>66227</td><td>8x10&quot; sheets; circles punched with standard hole punch</td></tr><tr><td>Sugarcane Bagasse Whole Biomass Feedstock</td><td>NIST</td><td>8491</td><td></td></tr><tr><td>Eastern Cottonwood (poplar) Whole Biomass Feedstock</td><td>NIST</td><td>8492</td><td></td></tr><tr><td>Monterey Pine Whole Biomass Feedstock</td><td>NIST</td><td>8493</td><td></td></tr><tr><td>Wheat Straw Whole Biomass Feedstock</td><td>NIST</td><td>8494</td><td></td></tr></tbody></table>

high-throughput screening of recalcitrance variations in lignocellulosic biomass: total lignin, lignin monomers, and enzymatic sugar release

The conversion of lignocellulosic biomass to fuels, chemicals, and other commodities has been explored as one possible pathway toward reductions in the use of non-renewable energy sources. In order to identify which plants, out of a diverse pool, have the desired chemical traits for downstream applications, attributes, such as cellulose and lignin content, or monomeric sugar release following an enzymatic saccharification, must be compared. The experimental and data analysis protocols of the standard methods of analysis can be time-consuming, thereby limiting the number of samples that can be measured. High-throughput (HTP) methods alleviate the shortcomings of the standard methods, and permit the rapid screening of available samples to isolate those possessing the desired traits. This study illustrates the HTP sugar release and pyrolysis-molecular beam mass spectrometry pipelines employed at the National Renewable Energy Lab. These pipelines have enabled the efficient assessment of thousands of plants while decreasing experimental time and costs through reductions in labor and consumables.

The combined effect of the thermochemical pretreatment and subsequent enzyme saccharification is measured as a function of the mass of glucose and xylose released at the end of the assay. The results are reported in terms of milligrams of glucose and xylose released per gram of biomass. This is in stark contrast to data reported from bench-scale assays, which is usually reported as percent theoretical yield based on compositional analysis of the starting material. As it is not yet practical to carry out compositional ana...

Watch this Scientific Journal Video about High-throughput Screening of Recalcitrance Variations in Lignocellulosic Biomass: Total Lignin, Lignin Monomers, and Enzymatic Sugar Release at JoVE.com

High-throughput Screening of Recalcitrance Variations in Lignocellulosic Biomass: Total Lignin, Lignin Monomers, and Enzymatic Sugar Release

Plant cell wall structure and chemistry traits are evaluated to identify ideal feedstocks for biofuels and bio-materials. Standard methods have limitations when applied to large data sets. These high-throughput pretreatment, enzyme saccharification, and pyrolysis-molecular beam mass spectrometry methods compare large numbers of biomass samples with decreased experimental time and cost.

The conversion of lignocellulosic biomass to fuels, chemicals, and other commodities has been explored as one possible pathway toward reductions in the ...

high-throughput-screening-recalcitrance-variations-lignocellulosic

Universidad de Cartagena UDEC

Research

JoVE Journal

Environment

9.9K Views.  National Renewable Energy Laboratory. Plant cell wall structure and chemistry traits are evaluated to identify ideal feedstocks for biofuels and bio-materials. Standard methods have limitations when applied to large data sets. These high-throughput pretreatment, enzyme saccharification, and pyrolysis-molecular beam mass spectrometry methods compare large numbers of biomass samples with decreased experimental time and cost.

Video: High-throughput Screening of Recalcitrance Variations in Lignocellulosic Biomass: Total Lignin, Lignin Monomers, and Enzymatic Sugar Release

High-throughput Screening of Carbohydrate-degrading Enzymes Using Novel Insoluble Chromogenic Substrate Assay Kits

Carbohydrates active enzymes (CAZymes) have multiple roles in vivo and are widely used for industrial processing in the biofuel, textile, detergent, paper and food industries. A deeper understanding of CAZymes is important from both fundamental biology and industrial standpoints. Vast numbers of CAZymes exist in nature (especially in microorganisms) and hundreds of thousands have been cataloged and described in the carbohydrate active enzyme database (CAZy). However, the rate of discovery of putative enzymes has outstripped our ability to biochemically characterize their activities. One reason for this is that advances in genome and transcriptome sequencing, together with associated bioinformatics tools allow for rapid identification of candidate CAZymes, but technology for determining an enzyme's biochemical characteristics has advanced more slowly. To address this technology gap, a novel high-throughput assay kit based on insoluble chromogenic substrates is described here. Two distinct substrate types were produced: Chromogenic Polymer Hydrogel (CPH) substrates (made from purified polysaccharides and proteins) and Insoluble Chromogenic Biomass (ICB) substrates (made from complex biomass materials). Both CPH and ICB substrates are provided in a 96-well high-throughput assay system. The CPH substrates can be made in four different colors, enabling them to be mixed together and thus increasing assay throughput. The protocol describes a 96-well plate assay and illustrates how this assay can be used for screening the activities of enzymes, enzyme cocktails, and broths.

A high-throughput assay for enzyme screening is described. This multiplexed ready-to-use assay kit comprises of pre-chosen Chromogenic Polymer Hydrogel (CPH) substrates and complex Insoluble Chromogenic Biomass (ICB) substrates. Target enzymes are polysaccharide degrading endo-enzymes and proteases.

Carbohydrates active enzymes (CAZymes) have multiple roles in vivo and are widely used for industrial processing in the biofuel, textile, detergent, paper ...

Biochemistry

Lignin Down-regulation of Zea mays via dsRNAi and Klason Lignin Analysis

To facilitate the use of lignocellulosic biomass as an alternative bioenergy resource, during biological conversion processes, a pretreatment step is needed to open up the structure of the plant cell wall, increasing the accessibility of the cell wall carbohydrates. Lignin, a polyphenolic material present in many cell wall types, is known to be a significant hindrance to enzyme access. Reduction in lignin content to a level that does not interfere with the structural integrity and defense system of the plant might be a valuable step to reduce the costs of bioethanol production. In this study, we have genetically down-regulated one of the lignin biosynthesis-related genes, cinnamoyl-CoA reductase (ZmCCR1) via a double stranded RNA interference technique. The ZmCCR1_RNAi construct was integrated into the maize genome using the particle bombardment method. Transgenic maize plants grew normally as compared to the wild-type control plants without interfering with biomass growth or defense mechanisms, with the exception of displaying of brown-coloration in transgenic plants leaf mid-ribs, husks, and stems. The microscopic analyses, in conjunction with the histological assay, revealed that the leaf sclerenchyma fibers were thinned but the structure and size of other major vascular system components was not altered. The lignin content in the transgenic maize was reduced by 7-8.7%, the crystalline cellulose content was increased in response to lignin reduction, and hemicelluloses remained unchanged. The analyses may indicate that carbon flow might have been shifted from lignin biosynthesis to cellulose biosynthesis. This article delineates the procedures used to down-regulate the lignin content in maize via RNAi technology, and the cell wall compositional analyses used to verify the effect of the modifications on the cell wall structure.

Lignin Down-regulation of Zea mays via dsRNAi and Klason Lignin Analysis

A double stranded RNA interference (dsRNAi) technique is employed to down-regulate the maize cinnamoyl coenzyme A reductase (ZmCCR1) gene to lower plant lignin content. Lignin down-regulation from the cell wall is visualized by microscopic analyses and quantified by the Klason method. Compositional changes in hemicellulose and crystalline cellulose are analyzed.

To facilitate the use of lignocellulosic biomass as an alternative bioenergy resource, during biological conversion processes, a pretreatment step is ...

Bioengineering

Extraction of Lignin with High &#946;-O-4 Content by Mild Ethanol Extraction and Its Effect on the Depolymerization Yield

Lignin valorization strategies are a key factor for achieving more economically competitive biorefineries based on lignocellulosic biomass. Most of the emerging elegant procedures to obtain specific aromatic products rely on the lignin substrate having a high content of the readily cleavable &#946;-O-4 linkage as present in the native lignin structure. This provides a miss-match with typical technical lignins that are highly degraded and therefore are low in &#946;-O-4 linkages. Therefore, the extraction yields, and the quality of the obtained lignin are of utmost importance to access new lignin valorization pathways. In this manuscript, a simple protocol is presented to obtain lignins with high &#946;-O-4 content by relatively mild ethanol extraction that can be applied to different lignocellulose sources. Furthermore, analysis procedures to determine the quality of the lignins are presented together with a depolymerization protocol that yields specific phenolic 2-arylmethyl-1,3-dioxolanes, which can be used to evaluate the obtained lignins. The presented results demonstrate the link between lignin quality and potential for the lignins to be depolymerized into specific monomeric aromatic chemicals. Overall, the extraction and depolymerization demonstrates a trade-off between the lignin extraction yield and the retention of the native aryl-ether structure and thus the potential of the lignin to be used as the substrate for the production of chemicals for higher-value applications.

Here, we present a protocol to perform ethanol extraction of lignin from several biomass sources. The effect of the extraction conditions on the lignin yield and &#946;-O-4 content are presented. Selective depolymerization is performed on the obtained lignins to obtain high aromatic monomer products.

Lignin valorization strategies are a key factor for achieving more economically competitive biorefineries based on lignocellulosic biomass. Most of the ...

Chemistry

Techniques for the Evolution of Robust Pentose-fermenting Yeast for Bioconversion of Lignocellulose to Ethanol

Lignocellulosic biomass is an abundant, renewable feedstock useful for production of fuel-grade ethanol and other bio-products. Pretreatment and enzyme saccharification processes release sugars that can be fermented by yeast. Traditional industrial yeasts do not ferment xylose (comprising up to 40% of plant sugars) and are not able to function in concentrated hydrolyzates. Concentrated hydrolyzates are needed to support economical ethanol recovery, but they are laden with toxic byproducts generated during pretreatment. While detoxification methods can render hydrolyzates fermentable, they are costly and generate waste disposal liabilities. Here, adaptive evolution and isolation techniques are described and demonstrated to yield derivatives of the native Scheffersomyces stipitis strain NRRL Y-7124 that are able to efficiently convert hydrolyzates to economically recoverable ethanol despite adverse culture conditions. Improved individuals are enriched in an evolving population using multiple selection pressures reliant on natural genetic diversity of the S. stipitis population and mutations induced by exposures to two diverse hydrolyzates, ethanol or UV radiation. Final evolution cultures are dilution plated to harvest predominant isolates, while intermediate populations, frozen in glycerol at various stages of evolution, are enriched on selective media using appropriate stress gradients to recover most promising isolates through dilution plating. Isolates are screened on various hydrolyzate types and ranked using a novel procedure involving dimensionless relative performance index (RPI) transformations of the xylose uptake rate and ethanol yield data. Using the RPI statistical parameter, an overall relative performance average is calculated to rank isolates based on multiple factors, including culture conditions (varying in nutrients and inhibitors) and kinetic characteristics. Through application of these techniques, derivatives of the parent strain had the following improved features in enzyme saccharified hydrolyzates at pH 5-6: reduced initial lag phase preceding growth, reduced diauxic lag during glucose-xylose transition, significantly enhanced fermentation rates, improved ethanol tolerance and accumulation to 40 g/L.

Adaptive evolution and isolation techniques are described and demonstrated to yield derivatives of Scheffersomyces stipitis strain NRRL Y-7124 that are able to rapidly consume hexose and pentose mixed sugars in enzyme saccharified undetoxified hydrolyzates and to accumulate over 40 g/L ethanol.

Lignocellulosic biomass is an abundant, renewable feedstock useful for production of fuel-grade ethanol and other bio-products. Pretreatment and enzyme ...

Comprehensive Compositional Analysis of Plant Cell Walls (Lignocellulosic biomass) Part I: Lignin

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st century. This has rekindled interest in the use of plant products as industrial raw materials for the production of liquid fuels for transportation1 and other products such as biocomposite materials7. Plant biomass remains one of the greatest untapped reserves on the planet4. It is mostly comprised of cell walls that are composed of energy rich polymers including cellulose, various hemicelluloses (matrix polysaccharides, and the polyphenol lignin6 and thus sometimes termed lignocellulosics. However, plant cell walls have evolved to be recalcitrant to degradation as walls provide tensile strength to cells and the entire plants, ward off pathogens, and allow water to be transported throughout the plant; in the case of trees up to more the 100 m above ground level. Due to the various functions of walls, there is an immense structural diversity within the walls of different plant species and cell types within a single plant4. Hence, depending of what crop species, crop variety, or plant tissue is used for a biorefinery, the processing steps for depolymerization by chemical/enzymatic processes and subsequent fermentation of the various sugars to liquid biofuels need to be adjusted and optimized. This fact underpins the need for a thorough characterization of plant biomass feedstocks. Here we describe a comprehensive analytical methodology that enables the determination of the composition of lignocellulosics and is amenable to a medium to high-throughput analysis. In this first part we focus on the analysis of the polyphenol lignin (Figure 1). The method starts of with preparing destarched cell wall material. The resulting lignocellulosics are then split up to determine its lignin content by acetylbromide solubilization3, and its lignin composition in terms of its syringyl, guaiacyl- and p-hydroxyphenyl units5. The protocol for analyzing the carbohydrates in lignocellulosic biomass including cellulose content and matrix polysaccharide composition is discussed in Part II2.

Comprehensive Compositional Analysis of Plant Cell Walls Lignocellulosic biomass Part I: Lignin

Plant biomass is a major carbon-neutral renewable resource that could be used for the production of biofuels. Plant biomass consists mainly of cell walls, a structurally complex composite material termed lignocellulosics. Here we describe a protocol for a comprehensive analysis of the content and composition of the polyphenolic lignin.

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st ...

Biology

Fractionation of Lignocellulosic Biomass using the OrganoCat Process

The shift from a petroleum-based to a more sustainable and bio-based economy requires the development of new refinery concepts to maintain the supply of raw materials and energy. For these novel and sustainable biorefinery concepts, it is important to use catalysts and solvents that are aligned with the principles of Green Chemistry. Therefore, the implementation of biogenic alternatives can be a promising solution. The lignocellulose pretreatment and fractionation process presented herein-OrganoCat-is an integrated fractionation of lignocellulose into its main components using biogenic acids such as 2,5-furandicarboxylic acid as catalyst. Hemicelluloses and other non-cellulosic polysaccharides are selectively depolymerized by the diluted acid and dissolved, while the crystalline cellulose remains in the solid pulp. In the presence of a second organic phase consisting of biogenic 2-methyltetrahydrofuran, disentangled lignin is extracted in situ. The process allows for the efficient fractionation of the three main components-lignin, cellulose, and non-cellulosic sugars. This helps to focus on the quality of the lignin, the improvement of enzymatic hydrolysis of the cellulose-enriched pulp, and the mild non-cellulosic sugar extraction with low degradation.

OrganoCat is a method for the pretreatment and fractionation of lignocellulose under mild conditions into lignin, fermentable sugars, and cellulose pulp. In a biogenic, biphasic solvent system of water and 2-methyltetrahydrofuran with 2,5-furancarboxylic acid as catalyst, the OrganoCat products are separated in situ for straightforward product recovery.

The shift from a petroleum-based to a more sustainable and bio-based economy requires the development of new refinery concepts to maintain the supply of ...

Ultrafast Lignin Extraction from Unusual Mediterranean Lignocellulosic Residues

Pretreatment is still the most expensive step in lignocellulosic biorefinery processes. It must be made cost-effective by minimizing chemical requirements as well as power and heat consumption and by using environment-friendly solvents. Deep eutectic solvents (DESs) are key, green, and low-cost solvents in sustainable biorefineries. They are transparent mixtures characterized by low freezing points resulting from at least one hydrogen bond donor and one hydrogen bond acceptor. Although DESs are promising solvents, it is necessary to combine them with an economic heating technology, such as microwave irradiation, for competitive profitability. Microwave irradiation is a promising strategy to shorten the heating time and boost fractionation because it can rapidly attain the appropriate temperature. The aim of this study was to develop a one-step, rapid method for biomass fractionation and lignin extraction using a low-cost and biodegradable solvent. 
In this study, a microwave-assisted DES pretreatment was conducted for 60 s at 800 W, using three kinds of DESs. The DES mixtures were facilely prepared from choline chloride (ChCl) and three hydrogen-bond donors (HBDs): a monocarboxylic acid (lactic acid), a dicarboxylic acid (oxalic acid), and urea. This pretreatment was used for biomass fractionation and lignin recovery from marine residues (Posidonia leaves and aegagropile), agri-food byproducts (almond shells and olive pomace), forest residues (pinecones), and perennial lignocellulosic grasses (Stipa tenacissima). Further analyses were conducted to determine the yield, purity, and molecular weight distribution of the recovered lignin. In addition, the effect of DESs on the chemical functional groups in the extracted lignin was determined by Fourier-transform infrared (FTIR) spectroscopy. The results indicate that the ChCl-oxalic acid mixture affords the highest lignin purity and the lowest yield. The present study demonstrates that the DES-microwave process is an ultrafast, efficient, and cost-competitive technology for lignocellulosic biomass fractionation.

Deep eutectic solvent-based, microwave-assisted pretreatment is a green, fast, and efficient process for lignocellulosic fractionation and high-purity lignin recovery.

Pretreatment is still the most expensive step in lignocellulosic biorefinery processes. It must be made cost-effective by minimizing chemical requirements ...

Label-free in situ Imaging of Lignification in Plant Cell Walls

Meeting growing energy demands safely and efficiently is a pressing global challenge. Therefore, research into biofuels production that seeks to find cost-effective and sustainable solutions has become a topical and critical task. Lignocellulosic biomass is poised to become the primary source of biomass for the conversion to liquid biofuels1-6. However, the recalcitrance of these plant cell wall materials to cost-effective and efficient degradation presents a major impediment for their use in the production of biofuels and chemicals4. In particular, lignin, a complex and irregular poly-phenylpropanoid heteropolymer, becomes problematic to the postharvest deconstruction of lignocellulosic biomass. For example in biomass conversion for biofuels, it inhibits saccharification in processes aimed at producing simple sugars for fermentation7. The effective use of plant biomass for industrial purposes is in fact largely dependent on the extent to which the plant cell wall is lignified. The removal of lignin is a costly and limiting factor8 and lignin has therefore become a key plant breeding and genetic engineering target in order to improve cell wall conversion.
Analytical tools that permit the accurate rapid characterization of lignification of plant cell walls become increasingly important for evaluating a large number of breeding populations. Extractive procedures for the isolation of native components such as lignin are inevitably destructive, bringing about significant chemical and structural modifications9-11. Analytical chemical in situ methods are thus invaluable tools for the compositional and structural characterization of lignocellulosic materials. Raman microscopy is a technique that relies on inelastic or Raman scattering of monochromatic light, like that from a laser, where the shift in energy of the laser photons is related to molecular vibrations and presents an intrinsic label-free molecular "fingerprint" of the sample. Raman microscopy can afford non-destructive and comparatively inexpensive measurements with minimal sample preparation, giving insights into chemical composition and molecular structure in a close to native state. Chemical imaging by confocal Raman microscopy has been previously used for the visualization of the spatial distribution of cellulose and lignin in wood cell walls12-14. Based on these earlier results, we have recently adopted this method to compare lignification in wild type and lignin-deficient transgenic Populus trichocarpa (black cottonwood) stem wood15. Analyzing the lignin Raman bands16,17 in the spectral region between 1,600 and 1,700 cm-1, lignin signal intensity and localization were mapped in situ. Our approach visualized differences in lignin content, localization, and chemical composition. Most recently, we demonstrated Raman imaging of cell wall polymers in Arabidopsis thaliana with lateral resolution that is sub-&mu;m18. Here, this method is presented affording visualization of lignin in plant cell walls and comparison of lignification in different tissues, samples or species without staining or labeling of the tissues.

Label-free in situ Imaging of Lignification in Plant Cell Walls

A method based on confocal Raman microscopy is presented that affords label-free visualization of lignin in plant cell walls and comparison of lignification in different tissues, samples or species.

Meeting growing energy demands safely and efficiently is a pressing global challenge. Therefore, research into biofuels production that seeks to find ...

A High Throughput Screen for Biomining Cellulase Activity from Metagenomic Libraries

Cellulose, the most abundant source of organic carbon on the planet, has wide-ranging industrial applications with increasing emphasis on biofuel production 1. Chemical methods to modify or degrade cellulose typically require strong acids and high temperatures. As such, enzymatic methods have become prominent in the bioconversion process. While the identification of active cellulases from bacterial and fungal isolates has been somewhat effective, the vast majority of microbes in nature resist laboratory cultivation. Environmental genomic, also known as metagenomic, screening approaches have great promise in bridging the cultivation gap in the search for novel bioconversion enzymes. Metagenomic screening approaches have successfully recovered novel cellulases from environments as varied as soils 2, buffalo rumen 3 and the termite hind-gut 4 using carboxymethylcellulose (CMC) agar plates stained with congo red dye (based on the method of Teather and Wood 5). However, the CMC method is limited in throughput, is not quantitative and manifests a low signal to noise ratio 6. Other methods have been reported 7,8 but each use an agar plate-based assay, which is undesirable for high-throughput screening of large insert genomic libraries. Here we present a solution-based screen for cellulase activity using a chromogenic dinitrophenol (DNP)-cellobioside substrate 9. Our library was cloned into the pCC1 copy control fosmid to increase assay sensitivity through copy number induction 10. The method uses one-pot chemistry in 384-well microplates with the final readout provided as an absorbance measurement. This readout is quantitative, sensitive and automated with a throughput of up to 100X 384-well plates per day using a liquid handler and plate reader with attached stacking system.

This protocol describes a high throughput screen for cellulolytic activity from a metagenomic library expressed in Escherichia coli. The screen is solution based and highly automated, and uses one-pot chemistry in 384 well microplates with the final readout as an absorbance measurement.

Cellulose, the most abundant source of organic carbon on the planet, has wide-ranging industrial applications with increasing emphasis on biofuel ...

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

Cellulase enzymes (endoglucanases, cellobiohydrolases, and &beta;-glucosidases) hydrolyze cellulose into component sugars, which in turn can be converted into fuel alcohols1. The potential for enzymatic hydrolysis of cellulosic biomass to provide renewable energy has intensified efforts to engineer cellulases for economical fuel production2. Of particular interest are fungal cellulases3-8, which are already being used industrially for foods and textiles processing.
Identifying active variants among a library of mutant cellulases is critical to the engineering process; active mutants can be further tested for improved properties and/or subjected to additional mutagenesis. Efficient engineering of fungal cellulases has been hampered by a lack of genetic tools for native organisms and by difficulties in expressing the enzymes in heterologous hosts. Recently, Morikawa and coworkers developed a method for expressing in E. coli the catalytic domains of endoglucanases from H. jecorina3,9, an important industrial fungus with the capacity to secrete cellulases in large quantities. Functional E. coli expression has also been reported for cellulases from other fungi, including Macrophomina phaseolina10 and Phanerochaete chrysosporium11-12. 
We present a method for high throughput screening of fungal endoglucanase activity in E. coli. (Fig 1) This method uses the common microbial dye Congo Red (CR) to visualize enzymatic degradation of carboxymethyl cellulose (CMC) by cells growing on solid medium. The activity assay requires inexpensive reagents, minimal manipulation, and gives unambiguous results as zones of degradation (&#x201C;halos&#x201D;) at the colony site. Although a quantitative measure of enzymatic activity cannot be determined by this method, we have found that halo size correlates with total enzymatic activity in the cell. Further characterization of individual positive clones will determine , relative protein fitness. 
Traditional bacterial whole cell CMC/CR activity assays13 involve pouring agar containing CMC onto colonies, which is subject to cross-contamination, or incubating cultures in CMC agar wells, which is less amenable to large-scale experimentation. Here we report an improved protocol that modifies existing wash methods14 for cellulase activity: cells grown on CMC agar plates are removed prior to CR staining. Our protocol significantly reduces cross-contamination and is highly scalable, allowing the rapid screening of thousands of clones. In addition to H. jecorina enzymes, we have expressed and screened endoglucanase variants from the Thermoascus aurantiacus and Penicillium decumbens (shown in Figure 2), suggesting that this protocol is applicable to enzymes from a range of organisms.

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

We describe a low cost, high throughput method to screen for fungal endoglucanase activity in E. coli. The method relies on a simple visual readout of substrate degradation, does not require enzyme purification, and is highly scalable. This allows for the rapid screening of large libraries of enzyme variants.

Cellulase enzymes (endoglucanases, cellobiohydrolases, and &beta;-glucosidases) hydrolyze cellulose into component sugars, which in turn can be converted ...

High-throughput Screening of Recalcitrance Variations in Lignocellulosic Biomass: Total Lignin, Lignin Monomers, and Enzymatic Sugar Release

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Label-free in situ Imaging of Lignification in Plant Cell Walls

A High Throughput Screen for Biomining Cellulase Activity from Metagenomic Libraries

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli