Name: estimation of crystalline cellulose content of plant biomass using the updegraff method
Uploaded: 2021-05-15T14:00:00.000+00:00
Duration: 12 min 34 s
Description: Watch this Scientific Journal Video about Estimation of Crystalline Cellulose Content of Plant Biomass using the Updegraff Method at JoVE.com

We thank the Department of Plant &#38; Soil Science and Cotton Inc. for their partial support of this study.

1. Experimental preparation
<ol>
	<li>Grind dried plant material into a fine powder.</li>
	<li>Protein Solubilization Buffer (PSB): Prepare stock solutions of 1 M Tris (pH 8.8), 0.5 M ethylenediaminetetraacetic acid (EDTA) (pH 8.0) and autoclave them. Make fresh PSB buffer from these stock solutions with final concentrations of 50 mM Tris, 0.5 mM EDTA and 10% sodium dodecyl sulfate (SDS) in sterile water.</li>
	<li>Prepare 100 mL of 70% ethanol (v/v): 70 mL of 100% ethanol and 30 mL of sterile water.</li>
	<li>Prepare 100 mL of methanol: chloroform in a 1:1 ratio (50 mL methanol and 50 mL chloroform).</li>
	<li>Prepare 82.5 mL of Updegraff reagent. Add 75 mL of 80% acetic acid to 7.5 mL of nitric acid so that the ultimate ratio of water: acetic acid: nitric acid is in 2:8:1 (v/v). To prepare 80% acetic acid, dissolve 80 mL of glacial acetic acid in 20 mL of sterile water (v/v).</li>
	<li>Prepare a fresh stock of 1 mg/mL glucose solution. Dissolve 10 mg of glucose in 10 mL of water (w/v).</li>
	<li>To prepare 100 mL of 67% sulfuric acid (v/v), add 67 mL of concentrated sulfuric acid to 33 mL of water. Always use a glass bottle and add acid slowly to the water. This step is exothermic (releases heat). Hence, prepare this solution on ice and cool in the refrigerator for at least 2 hours before use.</li>
	<li>Prepare fresh 0.2% anthrone (w/v) for each batch of experimental samples. Weigh 0.2 g of anthrone and dissolve in 100 mL of pre-chilled concentrated sulfuric acid in a glass bottle wrapped with aluminum foil. Keep in refrigerator for 1-2 hours before use. 
	&#8203;NOTE: Pre-chilling concentrated sulfuric acid in the refrigerator on the day of the experiment, and fresh preparation of anthrone is highly recommended for accurate estimation of glucose content.</li>
</ol>
2. Preparation of plant biomass material
<ol>
	<li>Collect plant biomass samples from 2-month-old cotton experimental lines grown in the greenhouse with the same growth conditions, the same development stage, the same position of the plants and the same type of tissue (leaf/stem/root). 
	NOTE: Collect a minimum of three biological replicates for each sample. Wash them thoroughly with water to remove all the dirt from the root tissue.</li>
	<li>Air-dry root tissue for 2 days on paper towels at room temperature to remove the moisture content (Figure 1). 
	NOTE: Air-dry desired tissue for cellulose estimation to prevent any fungal contamination.</li>
	<li>Place the root samples into individual containers. Then label and dry them in the incubator at 49 &#176;C for 10 days (Table of Materials). 
	NOTE: Alternatively, a freeze dryer can be used to dry the plant tissue in less time (1 or 2 days) without causing any chemical changes to the plant biomass material.</li>
	<li>Cut the dried samples into small pieces, freeze them in liquid nitrogen, and grind into uniform fine powder by using a mortar and pestle, a freezer mill, or a biomass grinder (Figure 1). 
	NOTE: The freezer mill was used at a rate of 10 cps for 3 cycles.</li>
	<li>Collect the grounded tissue (Figure 2) and proceed with the cell wall extraction. 
	&#8203;NOTE: The process can be paused at this point by storing the samples in airtight containers at room temperature.</li>
</ol>
3. Extraction of crude cell walls from plant biomass
NOTE: The plant cell walls contain cellulose, lignin, non-cellulose components, pectin, matrix proteins, phenolic compounds, and water26. Since cellulose is present in cell walls, the first step is to separate cell wall component from non-cell wall components of the plant biomass26.
<ol>
	<li>Label and weigh individual blank 2 mL tubes before starting the cell wall extraction process.</li>
	<li>Make a note of empty tube weights in a lab notebook before proceeding further.</li>
	<li>Weigh 20 mg of powdered tissue from step 2.5 and transfer it to pre-weighed 2 mL tubes and label them.</li>
	<li>Add 1 mL of protein solubilization buffer (PSB) (50 mM Tris hydrochloride (HCl) buffer pH 8.8, 0.5 mM EDTA, and 10% sodium dodecyl sulfate (SDS) to solubilize proteins. Vortex and centrifuge at 25,200 x g for 5 min at room temperature (RT). After centrifugation, discard the supernatant and save the pellet. 
	NOTE: The supernatant can be saved if protein component needs to be analyzed.</li>
	<li>Repeat step 3.4 two more times.</li>
	<li>To the retained pellet, add 1 mL of distilled water and vortex. Centrifuge at 25,200 x g for 5 min at room temperature (RT) and remove the supernatant.</li>
	<li>Repeat step 3.6 two more times.</li>
	<li>Add 1 mL of 70% ethanol to the saved pellet, vortex and heat it at 70 &#176;C for 1 h in a water bath/heat block to remove soluble components and starch from the samples. Vortex and centrifuge at 25,200 x g for 5 min at RT. Discard the supernatant and save the pellet.</li>
	<li>Repeat step 3.8 one more time.</li>
	<li>To the pellet add 1 mL of 100% methanol and vortex. Centrifuge at 25,200 x g for 5 min at room temperature (RT) and remove the supernatant.</li>
	<li>Add 1 mL of chloroform/methanol (chloroform and methanol in 1:1 ratio) to the pellet and vortex. Centrifuge at 25,200 x g for 5 min at RT and remove the supernatant. The addition of methanol and chloroform solvent solubilizes and removes the lipid fraction from the biomass27.</li>
	<li>To the pellet, add 1 mL of 100% acetone and vortex. Incubate at room temperature for 5 min. Centrifuge at 25,200 x g for 5 min at RT and remove the supernatant. Acetone removes pigments such as chlorophyll and free fatty acids from the biomass28,29.</li>
	<li>Dry the pellet at 37 &#176;C overnight or proceed further by vacuum drying. 
	&#8203;NOTE: The dried pellet is the crude cell wall used for crystalline cellulose estimation. The process can be paused at this point or proceed further using vacuum drier for drying samples.</li>
</ol>
4. Treatment with Updegraff reagent (acetic and nitric acid) to remove non-cellulosic components
NOTE: The protocol involves use of acids and other chemicals. Wear personal protective equipment (PPE) throughout the process.
<ol>
	<li>Measure 5 mg of the dried crude cell wall pellet into a fresh 2 mL screw capped tube. Note the exact weight (Figure 3)30.</li>
	<li>Include a positive control at this point. Use 2 mg of filter paper (Table of Materials) as a positive control that yields 80% cellulose content.</li>
	<li>Add 1.5 mL of the Updegraff reagent to the weighed 5 mg of cell wall extract and positive control. Mix by vortexing. 
	NOTE: The positive control should be processed in the same way as the experimental samples from this step onwards. This step should be carried out in the fume hood with proper personal protective equipment (PPE). 
	CAUTION: This step should be carried out in screw cap tubes to prevent splashing of sample and popping of the tubes. Three biological replicates of each sample along with three replicates for positive control should be included.</li>
	<li>Heat the suspension at 100 &#176;C for 30 min in a boiling water bath and cool on a bench for 10 min. Centrifuge at 25,200 x g for 10 min at RT. Remove the supernatant by centrifugation and save the pellet. 
	NOTE: The waste generated should be collected separately for organic solvents, sulfuric acid, and Updegraff reagent (acetic acid and nitric acid). Waste with acetic acid and nitric acid should be kept at cool temperatures with ventilated caps and never mixed with any other organic solvents and other acids to prevent any explosion.</li>
	<li>Add 1 mL of water to the pellet. Centrifuge at 25,200 x g&#160;for 10 min at RT. Remove 500 &#181;L of the supernatant and add 1 mL of acetone to the tube.</li>
	<li>Centrifuge at 25,200 x g for 5 min at RT. Remove 1 mL of supernatant, and add 1 mL of acetone to the tube and centrifuge at 25,200 x g&#160;for 5 min at RT.</li>
	<li>After centrifugation, remove all the supernatant and suspend the pellet in 1 mL of acetone. Incubate the tubes at room temperature for 5 min and spin at 25,200 x g for 5 min at RT.</li>
	<li>Discard the supernatant and save the pellet. Dry the pellet at 37 &#176;C overnight (Figure 3). 
	NOTE: The process can be paused at this point or continue further using vacuum drier for drying and proceed to next step.</li>
</ol>
5. Hydrolysis of cellulose by acid to produce glucose monomer units
<ol>
	<li>Add 1 mL of 67% sulfuric acid to the dried pellet. Vortex to mix the pellet completely in the acid.</li>
	<li>Shake tubes at room temperature for 1 h to dissolve the cellulose pellet in 67% sulfuric acid. 
	NOTE: As an alternative, solubilization of the cellulose pellet in 67% sulfuric acid can be improved by sonication of each sample for 10 min after 30 min of incubation in the shaker. After sonication, samples can be re-incubated in the shaker at RT. However, we observed complete solubility of cellulose pellet without sonication when we start with 5 mg of crude cell wall extract (Figure 3). This procedure worked well for various plant biomass samples3.</li>
</ol>
6. Measuring glucose content by the anthrone assay and estimation of cellulose content
<ol>
	<li>At this stage, the cellulose is in the form of free glucose monomers. Determine the amount of cellulose by measuring the amount of glucose present in the sample by spectrophotometry.</li>
	<li>Take 10 &#181;L of the sample from the step 5 and add it to 490 &#181;L of sterile distilled water to make a dilution of each sample to 500 &#181;L. Vortex the diluted mixture of sample and water for 10 seconds.</li>
	<li>Add 1 mL of 0.2% freshly prepared anthrone reagent to each tube and mix immediately by vortexing.</li>
	<li>Boil samples at 100 &#176;C for 10 min and cool the tubes on ice for 5 min. Transfer 200 &#181;L of each sample in three wells of a 96 well plate. Load the 96 well plate into a spectrophotometer to measure the absorbance at 620 nm. 
	&#8203;NOTE: For boiling at high temperature, use screw-capped tubes to prevent splashing of harmful chemicals and to avoid loss of samples.</li>
</ol>
7. Preparation of glucose standard curve
<ol>
	<li>To prepare glucose standard, make a fresh stock of 1 mg/mL glucose by dissolving 10 mg of glucose in 10 mL of sterile distilled water. Add this stock solution of 1 mg/mL in 20 &#181;L increments (0, 20, 40, 60, 80,100,120,140, 160, 180 &#181;L) to sterile distilled water (total volume 1 mL) to prepare glucose standards ranging from 0 &#181;g to 180 &#181;g/mL concentrations (Figure 3). Vortex each glucose standard after adding sterile water.</li>
	<li>From these 10 different concentrations, aliquot 500 &#181;L of each glucose concentration into a fresh 2 mL screw capped tube.</li>
	<li>Add 1 mL of freshly prepared 0.2% anthrone to each tube and mix immediately by vortexing. Incubate on ice and boil at 100 &#176;C for 10 min followed by incubation on ice for 5 min23.</li>
	<li>Transfer 200 &#181;L of each standard to three wells in a 96 well micro titer plate for three technical replicates and measure the absorbance at 620 nm (Figure 3).</li>
	<li>Develop a standard glucose curve by plotting different glucose concentrations (0 to 180 &#181;g/mL) against normalized absorbance values at 620 nm (Figure 7). Use the regression line Y= mx+c generated from the absorbance values to calculate the cellulose content in the prepared samples. 
	NOTE: Glucose standards must be freshly prepared for each set of experiment. The concentration of the glucose should be increased if the OD values are too high/ low so that these values fall within the range of the standard curve absorbance values. The regression line generates different m and c values based on the glucose standards used for each batch of samples. Mix tubes immediately after the addition of anthrone23. Anthrone, diluted samples and prepared standards were always kept cold until the anthrone assay was performed because of the exothermic nature of this reaction.</li>
</ol>

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The authors declare that they have no conflict of interest.

Cotton fibers are natural fibers produced from the cottonseed. Cotton fiber is a single cell with ~95% cellulose content2 with high crystalline cellulose content with extensive applications in textile industry31. As, cotton fiber contains ~95% cellulose, we have used cotton root tissues for demonstration of the estimation of crystalline cellulose content. Cotton root tissues are moderately rich in crystalline cellulose content and represents a commonly available plant bioma...

Cellulose is the primary load-bearing component of cell walls, which is present in both primary and secondary cell walls. The cell wall is an extracellular matrix that surrounds plant cells and is primarily composed of cellulose, lignin, hemicellulose, pectin, and matrix proteins. Approximately one third of plants biomass is cellulose1 and it plays significant roles in plant growth and development by providing strength, rigidity, rate and direction of cell growth, cell shape maintenance, and protection from biotic and abiotic stressors. Cotton fiber contains 95% cellulose2 content, while trees contain 40% to 50% of cellulose depending on the plant species and organ types3. The cellulose is composed of repeating units of cellobiose, a disaccharide of glucose residues connected by &#946;-1,4 glycosidic bonds4. Cellulosic ethanol is produced from the glucose derived from the cellulose present in the plant cell walls5. Cellulosic fiber is made up of several micro fibrils in which each micro fibril acts as core unit with 500-15000 glucose monomers1,6. The cellulose homopolymer is synthesized by plasma membrane embedded cellulose synthase complexes (CSC&#39;s)1,7. Individual cellulose synthase A (CESA) proteins synthesize glucan chains and the adjacent glucan chains are connected by hydrogen bonds to form crystalline cellulose1,8. Cellulose exists in several crystalline forms with two predominant forms, cellulose I&#945; and cellulose I&#946; as native forms9. In higher plants, cellulose exists in cellulose I&#946; form while lower plant cellulose exists in I&#945; form10,11. Overall, the cellulose plays a significant role in imparting strength and rigidity to the plant cell walls.
First generation biofuels are primarily produced from corn starch, cane sugars, and beet sugars, which are food sources, while second-generation biofuels are focusing on the biofuel production from non-food plant biomass cell wall material12. Accurate estimation of crystalline cellulose content is not only important for fundamental research on cellulose biosynthesis and cell wall dynamics but also for applied biofuel and bio products research. Various methods have been developed and optimized for estimation of cellulose in the plant biomass, and the Updegraff method is the most widely used method for cellulose estimation. The first reported method for cellulose estimation was by Cross and Bevan in 190813. The method was based on the principle of alternate chlorination and extraction by sodium sulphate. However, the cellulose obtained by the original as well as modified protocols of Cross and Bevan method showed contamination of small fractions of lignin in addition to a substantial amount of xylans and mannans14. Despite several modifications to remove lignin and hemicelluloses from the cellulose fraction, the Cross-Bevan method retained a considerable amount of mannans along with cellulose. Later, Kurschner&#39;s method was developed by employing nitric acid and ethanol to extract cellulose15. This method stated that total lignin and 75% of pentosans were removed but the true cellulose results were the same as those estimated by chlorination method of Cross and Bevan. Another method (Norman and Jenkins) was developed by employing methanol-benzene, sodium sulphate, and sodium hypochlorite to extract cellulose16. This method also retained some fraction of lignin (3%) and significant amounts of pentosans leading to in accurate estimation of cellulose. Later, Kiesel and Semiganowsky used a different approach to hydrolyze cellulose using 80% concentrated sulfuric acid, and the hydrolyzed reduced sugars were estimated by Bertrand&#39;s method17. The two methods, Waksman&#39;s and Stevens18 and Salo14,19 which were developed based on Kiesel and Semiganowsky&#39;s method, also yielded 4-5% less cellulose content compared to earlier methods20.
The Updegraff method is the most widely used method for the estimation of crystalline cellulose content. This method was first described by Updegraff for the measurement of cellulose in 196921. The Updegraff method integrates the Kurschner method (use of nitric acid), Kiesel and Seminowsky methods (hydrolysis of cellulose into glucose monomers using sulfuric acid) with some modifications, and the anthrone assay of Viles and Silverman for simple colorimetric estimation of glucose and crystalline cellulose content22. The principle of this method is the use of acetic acid and nitric acid (Updegraff reagent) to eliminate hemicellulose and lignin from the homogenized plant tissues, which leaves acetic/nitric acid resistant cellulose for further processing and estimation15. The acetic/nitric acid resistant cellulose is treated with 67% sulfuric acid to break the cellulose into glucose monomers and the released glucose monomers are estimated by anthrone assay21,23. Several modifications of the original Updegraff method were used to simplify the procedure and cellulose estimation by anthrone assay24. Broadly, this method can be divided into five phases. In the first phase, the plant material is prepared. In the second phase, the crude cell wall is separated from the total biomass, as cellulose is the key component of plant cell walls. Later, in the third phase, the cellulose is separated from the non-cellulosic cell wall components by treating with Updegraff reagent. In the fourth phase, the acetic/nitric acid resistant cellulose is broken into glucose monomers by sulfuric acid treatment. Sulfuric acid treatment of cellulose results in formation of 5-hydroxymethylfurfural compounds from the reaction of glucose monomers with sulfuric acid. Finally, in the last phase, the anthrone generates a greenish blue complex by boiling with the furfural compound generated in the previous phase25. This anthrone based colorimetric method was first used in 1942 by Dreywood. Anthrone is a dye that identifies furfural compounds of pentose and hexose dehydrated products such as 5-hydroxymethylfurfural, under acidic conditions. Reaction with hexose produces an intense color and better response compared to pentoses25. The amount of bound glucose is measured by spectrophotometer absorbance at 620 nm and the intensity of the greenish blue complex is directly proportional to the amount of sugar in the sample. The measured absorbance values were compared with a glucose standard curve regression line to calculate the glucose concentration of the sample. The measured glucose content was used to estimate the cellulose content of the plant biomass.

<table><tbody><tr><td>Acetone</td><td>Fisher Chemical</td><td>A18-500</td><td>Used in the protocol</td></tr><tr><td>Anthrone</td><td>Sigma Aldrich</td><td>90-44-8</td><td>For colorimetric assay</td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5424</td><td>For centrifugation</td></tr><tr><td>Chloroform</td><td>Mallinckrodt</td><td>67-66-3</td><td>Used in the protocol</td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Sigma Aldrich</td><td>6381-92-6</td><td>Used in the protocol</td></tr><tr><td>Ethanol</td><td>Millipore Sigma</td><td>EM-EX0276-4S</td><td>Used in the protocol</td></tr><tr><td>Filter paper</td><td>Whatman</td><td>1004-090</td><td>Positive control</td></tr><tr><td>Glacial acetic acid</td><td>Sigma</td><td>SKU A6283</td><td>Used in the protocol</td></tr><tr><td>Heat block/ ThermoMixer F1.5</td><td>Eppendorf</td><td>13527550</td><td>For controlled temperatures</td></tr><tr><td>Incubator</td><td>Fisherbrand</td><td>150152633</td><td>Used for drying plant sample</td></tr><tr><td>Measuring Scale</td><td>Mettler Toledo</td><td>30243386</td><td>For specific quantities</td></tr><tr><td>Methanol 100 %</td><td>Fisher Chemical</td><td>A412-500</td><td>Used in the protocol</td></tr><tr><td>Microplate (96 well)</td><td>Evergreen Scientific</td><td>222-8030-01F</td><td>For anthrone assay</td></tr><tr><td>Nitric acid</td><td>Sigma Aldrich</td><td>695041</td><td>Used in the protocol</td></tr><tr><td>Polypropylene Microvials (2 mL) / screw capped tubes</td><td>BioSpec Products</td><td>10831</td><td>For high temperatures</td></tr><tr><td>Spectrophotometer(Multimode Detector)</td><td>Beckmancoulter DTX880</td><td>1000814</td><td>For measuring absorbances</td></tr><tr><td>Spex SamplePrep 6870 Freezer / Mill</td><td>Spex Sample Prep</td><td>68-701-15</td><td>For grinding plant tissues into fine powder</td></tr><tr><td>Sulphuric acid</td><td>J.T.Baker</td><td>02-004-382</td><td>Used in the protocol</td></tr><tr><td>Sodium dodecyl sulfate (SDS)</td><td>Sigma Aldrich</td><td>151-21-3</td><td>Used in the PSB buffer</td></tr><tr><td>Tubes (2 mL)</td><td>Fisher Scientific</td><td>05-408-138</td><td>Used in the protocol</td></tr><tr><td>Tris Hydrochloride</td><td>Sigma Aldrich</td><td>&amp;nbsp;1185-53-1</td><td>Used in the PSB buffer</td></tr><tr><td>Ultrapure distilled water</td><td>Invitrogen</td><td>10977</td><td>Used in the protocol</td></tr><tr><td>Vacuum dryer (vacufuge plus)</td><td>Eppendorf</td><td>22820001</td><td>For drying samples</td></tr><tr><td>Vortex mixer</td><td>Fisherbrand</td><td>14-955-151</td><td>For mixing</td></tr><tr><td>Waterbath</td><td>Thermoscientific</td><td>TSGP02PM05</td><td>For temperature controlled conditions at specific steps</td></tr><tr><td>Weighing Paper</td><td>Fisher Scientific</td><td>09-898-12A</td><td>Used in the protocol</td></tr></tbody></table>

estimation of crystalline cellulose content of plant biomass using the updegraff method

Cellulose is the most abundant polymer on Earth generated by photosynthesis and the main load-bearing component of cell walls. The cell wall plays a significant role in plant growth and development by providing strength, rigidity, rate and direction of cell growth, cell shape maintenance, and protection from biotic and abiotic stressors. The cell wall is primarily composed of cellulose, lignin, hemicellulose and pectin. Recently plant cell walls have been targeted for the second-generation biofuel and bioenergy production. Specifically, the cellulose component of the plant cell wall is used for the production of cellulosic ethanol. Estimation of cellulose content of biomass is critical for fundamental and applied cell wall research. The Updegraff method is simple, robust, and the most widely used method for the estimation of crystalline cellulose content of plant biomass. The alcohol insoluble crude cell wall fraction upon treatment with Updegraff reagent eliminates the hemicellulose and lignin fractions. Later, the Updegraff reagent resistant cellulose fraction is subjected to sulfuric acid treatment to hydrolyze the cellulose homopolymer into monomeric glucose units. A regression line is developed using various concentrations of glucose and used to estimate the amount of the glucose released upon cellulose hydrolysis in the experimental samples. Finally, the cellulose content is estimated based on the amount of glucose monomers by colorimetric anthrone assay.

Cotton plants grown in the green house were selected for this study. Two different experimental lines of cotton were selected for comparative analysis of cellulose content. For each experimental line, the root tissue was collected from three biological replicates. A total of 500 mg of tissue was homogenized and 20 mg of it was used for crude cell wall extraction. Later, 5 mg of crude cell wall extract was used for Updegraff reagent treatment to remove hemicellulose and lignin from cellulose. The purified cellulose was hy...

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Estimation of Crystalline Cellulose Content of Plant Biomass using the Updegraff Method

The Updegraff method is the most widely used method for the cellulose estimation. The main purpose of this demonstration is to provide a detailed Updegraff protocol for estimation of cellulose content in plant biomass samples.

Cellulose is the most abundant polymer on Earth generated by photosynthesis and the main load-bearing component of cell walls. The cell wall plays a ...

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Research

JoVE Journal

Environment

13.0K Views.  Texas Tech University. Hier demonstreren we de Updegraff-methode, 's werelds meest gebruikte methode voor de schatting van het kristallijne cellulosegehalte. Cellulose bestaat in principe uit glucosemonomeren verbonden door bèta-1, 4 glycosidische bindingen. We hebben dit protocol in vijf fasen verdeeld.In de eerste fase bereiden we het biomassamateriaal van de installatie voor. In de tweede fase halen we de ruwe celwand eruit. In de derde fase gebruiken we een Updegraff-behandeling waarbij Updegraff bestaa...

Video: Estimation of Crystalline Cellulose Content of Plant Biomass using the Updegraff Method

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.

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 ...

Combining Raman Imaging and Multivariate Analysis to Visualize Lignin, Cellulose, and Hemicellulose in the Plant Cell Wall

The application of Raman imaging to plant biomass is increasing because it can offer spatial and compositional information on aqueous solutions. The analysis does not usually require extensive sample preparation; structural and chemical information can be obtained without labeling. However, each Raman image contains thousands of spectra; this raises difficulties when extracting hidden information, especially for components with similar chemical structures. This work introduces a multivariate analysis to address this issue. The protocol establishes a general method to visualize the main components, including lignin, cellulose, and hemicellulose within the plant cell wall. In this protocol, procedures for sample preparation, spectral acquisition, and data processing are described. It is highly dependent upon operator skill at sample preparation and data analysis. By using this approach, a Raman investigation can be performed by a non-specialist user to acquire high-quality data and meaningful results for plant cell wall analysis.

This protocol aims to present a general method to visualize lignin, cellulose, and hemicellulose in plant cell walls using Raman imaging and multivariate analysis.

The application of Raman imaging to plant biomass is increasing because it can offer spatial and compositional information on aqueous solutions. The ...

Biochemistry

Determination of the Glycogen Content in Cyanobacteria

Cyanobacteria accumulate glycogen as a major intracellular carbon and energy storage&#160;during photosynthesis. Recent developments in research have highlighted complex mechanisms of glycogen metabolism, including the diel cycle of biosynthesis and catabolism, redox regulation, and the involvement of non-coding RNA. At the same time, efforts are being made to redirect carbon from glycogen to desirable products in genetically engineered cyanobacteria to enhance product yields. Several methods are used to determine the glycogen contents in cyanobacteria, with variable accuracies and technical complexities. Here, we provide a detailed protocol for the reliable determination of the glycogen content in cyanobacteria that can be performed in a standard life science laboratory. The protocol entails the selective precipitation of glycogen from the cell lysate and the enzymatic depolymerization of glycogen to generate glucose monomers, which are detected by a glucose oxidase-peroxidase (GOD-POD) enzyme coupled assay. The method has been applied to Synechocystis sp. PCC 6803 and Synechococcus sp. PCC 7002, two model cyanobacterial species that are widely used in metabolic engineering. Moreover, the method successfully showed differences in the glycogen contents between the wildtype and mutants defective in regulatory elements or glycogen biosynthetic genes.

Here, we present a reliable and easy assay to measure the glycogen content in cyanobacterial cells. The procedure entails precipitation, selectable depolymerization, and the detection of glucose residues. This method is suitable for both wildtype and genetically engineered strains and can facilitate the metabolic engineering of cyanobacteria.

Cyanobacteria accumulate glycogen as a major intracellular carbon and energy storage&#160;during photosynthesis. Recent developments in research have ...

Structural Characterization of Mannan Cell Wall Polysaccharides in Plants Using PACE

Plant cell wall polysaccharides are notoriously difficult to analyze, and most methods require expensive equipment, skilled operators, and large amounts of purified material. Here, we describe a simple method for gaining detailed polysaccharide structural information, including resolution of structural isomers. For polysaccharide analysis by gel electrophoresis (PACE), plant cell wall material is hydrolyzed with glycosyl hydrolases specific to the polysaccharide of interest (e.g., mannanases for mannan). Large format polyacrylamide gels are then used to separate the released oligosaccharides, which have been fluorescently labeled. Gels can be visualized with a modified gel imaging system (see Table of Materials). The resulting oligosaccharide fingerprint can either be compared qualitatively or, with replication, quantitatively. Linkage and branching information can be established using additional glycosyl hydrolases (e.g., mannosidases and galactosidases). Whilst this protocol describes a method for analyzing glucomannan structure, it can be applied to any polysaccharide for which characterized glycosyl hydrolases exist. Alternatively, it can be used to characterize novel glycosyl hydrolases using defined polysaccharide substrates.

A protocol for the structural analysis of polysaccharides by gel electrophoresis (PACE), using mannan as an example, is described.

Plant cell wall polysaccharides are notoriously difficult to analyze, and most methods require expensive equipment, skilled operators, and large amounts ...

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

Estimation of Plant Biomass Lignin Content using Thioglycolic Acid (TGA)

Lignin is a natural polymer that is the second most abundant polymer on Earth after cellulose. Lignin is mainly deposited in plant secondary cell walls and is an aromatic heteropolymer primarily composed of three monolignols with significant industrial importance. Lignin plays an important role in plant growth and development, protects&#160;from biotic and abiotic stresses, and in the quality of animal fodder, the wood, and industrial lignin products. Accurate estimation of lignin content is essential for both fundamental understanding of the lignin biosynthesis and industrial applications of biomass. The thioglycolic acid (TGA) method is a highly reliable method of estimating the total lignin content in the plant biomass. This method estimates the lignin content by forming thioethers with the benzyl alcohol groups of lignin, which are soluble in alkaline conditions and insoluble in acidic conditions. The total lignin content is estimated using a standard curve generated from commercial bamboo lignin.

Estimation of Plant Biomass Lignin Content using Thioglycolic Acid TGA

Here, we present a modified TGA method for estimation of lignin content in herbaceous plant biomass. This method estimates the lignin content by forming specific thioether bonds with lignin and presents an advantage over the Klason method, as it requires a relatively small sample for lignin content estimation.

Lignin is a natural polymer that is the second most abundant polymer on Earth after cellulose. Lignin is mainly deposited in plant secondary cell walls ...

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

Comprehensive Compositional Analysis of Plant Cell Walls (Lignocellulosic biomass) Part II: Carbohydrates

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 transportation2 and other products such as biocomposite materials6. 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, and the polyphenol lignin5 and thus sometimes termed lignocellulosics. However, plant cell walls have evolved to be recalcitrant to degradation as walls contribute extensively to the strength and structural integrity of the entire plant. Despite its necessary rigidity, the cell wall is a highly dynamic entity that is metabolically active and plays crucial roles in numerous cell activities such as plant growth and differentiation5. 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 depolymerisation 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 (Figure 1). The method starts of with preparing destarched cell wall material. The resulting lignocellulosics are then split up to determine its monosaccharide composition of the hemicelluloses and other matrix polysaccharides1, and its content of crystalline cellulose7. The protocol for analyzing the lignin components in lignocellulosic biomass is discussed in Part I3.

Comprehensive Compositional Analysis of Plant Cell Walls Lignocellulosic biomass Part II: Carbohydrates

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 wall derived carbohydrates.

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 ...

Chemistry

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 ...

Estimation of Crystalline Cellulose Content of Plant Biomass using the Updegraff Method

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