This work over the years has been supported by various financial awards that included organizations such as the Pulp and Paper Research Institute of Canada, McGill University Montreal, the Natural Sciences and Engineering Research Council of Canada, National Science Foundation USA, United States Department of agriculture, and the Solvay company.

The following flow chart (Figure 6) outlines the whole experimental protocol to perform a 31P NMR analysis of lignins and tannins.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62696/62696fig06.jpg" /> 
Figure 6: Procedure for the 31P NMR analysis of lignins and tannins. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Sample pretreatment
<ol>
	<li>Dry an aliquot (around 100 mg) of the analyte (lignin or tannin sample) overnight in a vacuum oven set at 40 &#176;C. 
	NOTE: Particular attention is needed on the temperature selection since temperatures higher than 40 &#176;C may chemically alter the sensitive structure of the examined polyphenols.</li>
	<li>After drying, rapidly transfer the sample to an anhydrous calcium sulfate desiccator until it reaches room temperature. This step is compulsory to avoid the sample absorbing humidity from the environment.</li>
</ol>
2. Solvent solution preparation
<ol>
	<li>Prepare a pyridine/deuterated chloroform solvent mixture in a 20 mL sample vial by mixing anhydrous pyridine and deuterated chloroform in a 1.6/1 (v/v) ratio. 
	CAUTION: Pay attention while manipulating pyridine and deuterated chloroform. These compounds are flammable, harmful, and toxic. Prepare and use the solution in a well-ventilated fume-hood using appropriate gloves.</li>
	<li>Add 5-8 g of well-washed and dried activated 5A molecular sieves in 3.2 mm pellets to remove water traces. In addition, the use of a septum cap is highly recommended to prevent air contact and moisture contamination of the solvent system. Store the prepared solution in the dark.</li>
</ol>
3. Internal standard solution (IS) preparation
<ol>
	<li>In a 2 mL Erlenmeyer flask, prepare a 0.1 M solution of chromium (III) acetylacetonate (about 10 mg) and internal standard (around 35.8 mg of NHND or 77.3 mg of cholesterol) in the solvent solution previously prepared. 
	CAUTION: Chromium (III) acetylacetonate is harmful; during its manipulation, wear appropriate gloves.</li>
	<li>Record the exact weight of the IS added in the IS solution.</li>
	<li>Transfer the IS solution in a vial equipped with a sealed cap containing activated molecular sieves (see point 2.2) and store it in the dark at 40 &#176;C.</li>
</ol>
4. NMR sample solution preparation
<ol>
	<li>Accurately weigh ~30 mg of sample in a 2 mL vial equipped with a stirring bar. Seal the vial with a septum cap.</li>
	<li>Add 0.5 mL of the solvent system solution to the sample vial.</li>
	<li>Transfer 100 &#181;L of the IS solution in the sample vial via a micropipette. Magnetically stir the resulting dispersion (500 rpm) until all the lignin or the tannin is dissolved, resulting in a clear solution. 
	NOTE: Since complete sample solubilization is imperative, this step could take up to 12 h.</li>
	<li>Transfer 0.1 mL of TMDP to the sample solution. Place the sample under vigorous magnetic stirring. Keep the sample solution sealed. Use TMDP in a well-ventilated fume-hood while wearing appropriate gloves. 
	CAUTION: TMDP and its vapors are corrosive, harmful, and rapidly interact with water. 
	NOTE: The formation of a yellow precipitate is due to water traces in the sample or the pyridine/chloroform solution. In such a case, the procedure must be repeated by ensuring all possible moisture contamination is avoided.</li>
	<li>Transfer the sample solution into an NMR tube using a Pasteur pipette.</li>
</ol>
5. NMR analysis
<ol>
	<li>Load the tube into the NMR instrument. The spectrometer used to perform this analysis needs a broadband probe.</li>
	<li>Fix the experimental parameters according to the setting shown in Table 11.</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>PULSE PROGRAM</td>
			<td>Inverse gated decoupling pulse (zgig)</td>
		</tr>
		<tr>
			<td>NUCLEOUS</td>
			<td>31P</td>
		</tr>
		<tr>
			<td>SPECTRAL WIDTH</td>
			<td>100 p.p.m.</td>
		</tr>
		<tr>
			<td>ACQUISITION TIME</td>
			<td>&#160;- 0.8 s</td>
		</tr>
		<tr>
			<td>RELAXATION DELAY</td>
			<td>&#160;&#8805; 10 s</td>
		</tr>
		<tr>
			<td>SCANS NUMBER</td>
			<td>&#160;64 or more</td>
		</tr>
		<tr>
			<td>SPECTRUM CENTER</td>
			<td>&#160;140 p.p.m.</td>
		</tr>
	</tbody>
</table>
Table 1: Experimental parameters to record 31P NMR spectra of derivatized lignins or tannins.
<ol start="3">
	<li>Set the spectrometer frequency using the resonance frequency of deuterated chloroform, shim the sample and tune the spectrometer. Then, start the acquisition.</li>
</ol>
6. Spectrum processing and analysis
<ol>
	<li>Process 31P NMR raw data by an appropriate standard software according to the following steps.
	<ol>
		<li>Perform Fourier transformation.</li>
		<li>Adjust Phase by manual phase correction (Processing | Phase Correction | Manual Correction).</li>
		<li>Correct baseline manually, carefully setting zero points (Processing | Baseline | Multipoint Baseline Correction).</li>
	</ol>
	</li>
	<li>Signal calibration.
	<ol>
		<li>Set the signal for the phosphitylated water at the chemical shift value of 132.2 ppm (Analysis | Reference | Reference). 
		NOTE: The presence of a sharp 31P signal at 175 ppm is due to the excess of TMDP. Its presence ensures the complete derivatization of the sample. If this peak is absent, one needs to revisit the whole procedure by providing a thorough sample and solvent drying and adding more TMDP. Once this is guaranteed, the spectrum is zoomed in the spectral range 132 to around 150 ppm (Figure 7).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/62696/62696fig07.jpg" /> 
Figure 7: Check the presence of an excess of TMDP: if it can be seen, the derivatization of the sample is complete. The spectra can then be analyzed. To do that zoom in&#160;the spectral range between 155 and 132 ppm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>Integration
	<ol>
		<li>Normalize integration by setting the internal standard to 1.0 (click on the Peak | Edit Integral | Normalized: 1.00). Perform spectrum integration according to the chemical shifts reported in the following tables. Use Table 2 for lignins and Table 3 for tannins.</li>
	</ol>
	</li>
</ol>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>FUNCTIONAL GROUP</td>
			<td>&#160;CHEMICAL SHIFT (ppm)</td>
		</tr>
		<tr>
			<td>Aliphatic OH</td>
			<td>&#160;149.0-146.0</td>
		</tr>
		<tr>
			<td>Phenolic OH</td>
			<td>&#160;144.0-137.4</td>
		</tr>
		<tr>
			<td>C5 substituted phenolic OH</td>
			<td>&#160;143.0-140.2</td>
		</tr>
		<tr>
			<td>5-5&#8217; phenolic OH</td>
			<td>&#160;141.7-140.2</td>
		</tr>
		<tr>
			<td>Syringyl OH</td>
			<td>&#160;143.2-142.7</td>
		</tr>
		<tr>
			<td>4-O-5&#8217; OH</td>
			<td>&#160;142.8-141.7</td>
		</tr>
		<tr>
			<td>Guaiacyl OH</td>
			<td>&#160;140.2-138.8</td>
		</tr>
		<tr>
			<td>p-hydroxyphenyl OH</td>
			<td>&#160;138.8-137.4</td>
		</tr>
		<tr>
			<td>COOH</td>
			<td>&#160;136.0-133.6</td>
		</tr>
		<tr>
			<td>Tricin</td>
			<td>&#160;137.0-136.0</td>
		</tr>
	</tbody>
</table>
Table 2: 31P NMR chemical shifts for lignin phosphitylated OH-groups.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>FUNCTIONAL GROUP</td>
			<td>&#160;CHEMICAL SHIFT (ppm)</td>
		</tr>
		<tr>
			<td colspan="2">Ring A</td>
		</tr>
		<tr>
			<td>o-unsubstituted phenolic</td>
			<td>&#160;137.9&#8211;137.4</td>
		</tr>
		<tr>
			<td>o-substituted PHENOLIC</td>
			<td>&#160;138.8&#8211;137.9</td>
		</tr>
		<tr>
			<td colspan="2">Ring B</td>
		</tr>
		<tr>
			<td>Catechol OH</td>
			<td>&#160;140.2&#8211;138.8</td>
		</tr>
		<tr>
			<td>Pyrogallol OH</td>
			<td>&#160;144.0&#8211;140.2</td>
		</tr>
		<tr>
			<td colspan="2">Ring C</td>
		</tr>
		<tr>
			<td>AliphatiC OH</td>
			<td>&#160;146.0&#8211;145.0</td>
		</tr>
	</tbody>
</table>
Table 3: 31P NMR chemical shift for tannin phosphitylated OH-groups.
NOTE: Using standard spectral-processing software, it is possible to set pre-defined regions of the chemical shift to be integrated. This opportunity is advantageous when several spectra have to be processed.
7. Functional group quantification
<ol>
	<li>Calculate the concentration of the IS solution. 
	<img alt="Equation 1" src="/files/ftp_upload/62696/62696eq1v2.png" /></li>
	<li>Calculate the equivalent amount of the specific signal: 
	<img alt="Equation 2" src="/files/ftp_upload/62696/62696eq2v2.png" /> 
	<img alt="Equation 3" src="/files/ftp_upload/62696/62696eq3v2.png" /></li>
</ol>

Claudia Crestini and Dimitris S Argyropoulos ensure that all authors (C.C., N.P., and D.S.A.) have no conflicts of interest.

The described method represents the implementation and optimization of the analytical protocol aimed at the qualitative and quantitative characterization of lignins as developed by Argyropoulos37,38,39,40,41,42. Compared to many other techniques available for lignin structural elucidation, the method has been widely accepted as being among the most facile, rapid, and reproducible. The validity of the wet chemical methods (e.g., nitrobenzene, permanganate oxidations, etc.) relies on the good experimental skills of the operator, effectively confining the method to limited operators. Furthermore, it is not uncommon to encounter correction factors in the literature for wet chemical methods to account for several drawbacks. The described 31P NMR protocol does not require advanced experimental skills making this readily applicable, user-friendly, and widely available. Compared to other instrumental analytical methods, 31P NMR is the only technique capable of precisely detecting and quantifying the different hydroxy groups in lignins. For instance, FTIR can be used to identify various hydroxy groups such as 1H NMR. Both techniques, however, suffer since they cannot offer reliable quantitative data due to extensive signal overlap issues. Another widely used technique is UV-Vis spectroscopy, firstly reported by Goldschmid. The approach, however, is limited to a general overall determination of hydroxy groups since it cannot effectively differentiate among aliphatic, aromatic, and carboxylic OHs47.
From an economic point of view, the only limitation of the 31P NMR technique is the price of TMDP, which is a relatively expensive reagent. It costs about 190 USD per gram; consequently, if the cost of analysis would be approximated only to the price of TMDP, excluding those deriving from the pyridine/chloroform mixture and those of the operator time, it would amount to about 24 USD per analysis. To solve this issue, many laboratories resort to synthesizing TMDP, thus, reducing the reagent costs. To do this, pinacol and phosphorous trichloride are reacted in the presence of triethylamine44. Technically, this reaction is relatively easy; however, care in the use of phosphorous trichloride and its work-up, including well-controlled vacuum distillation, is required. More details as to the synthesis of the TMDP can be supplied upon request.
Even though this protocol is among the best in terms of ease, reproducibility, and precision, some critical points need to be highlighted. Firstly, the sample needs to be fully soluble in the identified pyridine/chloroform mixture. This consideration is fundamental because the quantitative phosphitylation reaction of the hydroxyl groups needs to take place under completely homogeneous conditions. If only part of the sample is solubilized, the resulting analysis would be inaccurate. Secondly, the sample to be examined needs to be moisture- and solvent-free since these variables will detrimentally affect the precision and overall success of the analysis. Traces of humidity will react with TMDP giving 2-hydroxy-4,4&#39;-5,5&#39;-tetramethyl-1,3,2-dioxaphospholane. This compound is a pale-yellow flocculating salt, insoluble in the pyridine/chloroform solvent mixture, causing inadequate NMR signal acquisition. Since only a small weight (~30 mg) of a sample is required, it needs to be free of volatiles for its precise weight to be accurately known before the analysis.
Sometimes, sample solvation issues can be promoted (especially for highly oxidized samples) by adding small amounts of a co-solvent (i.e., dimethylformamide), aiding sample dissolution. In principle, every solvent that does not interact with TMDP can be used to help sample dissolution. The election of a co-solvent cannot include co-solvents containing labile hydroxy or amino groups since they react with the reagent, causing misleading final spectra. Notably, dimethylsulfoxide also reacts with TMDP precluding its use as a co-solvent. Pyridine-based ionic liquids, such as 1-allyl-3-butylpyridinium chloride, can be used when solubility issues arise; however, the ionic liquid should be once again dry48. To dissolve lignosulfonates (a lignin type characterized by a high sulfonation degree), a pre-treatment involving the conversion of neutralized groups into their acidic form was demonstrated to be helpful. Lignosulfonates can be conveniently converted into their acidic conditions using acidic exchange resins in aqueous media. The resulting lignosulfonic acids are isolated from the solution by their adsorption on specific resins (e.g., XAD-7) and desorption in ethanol. Evaporation of the ethanolic solutions over reduced pressure at 40 &#176;C allows the isolation of lignosulfonic acids. These lignins can then be characterized by 31P NMR because they are soluble in the pyridine/chloroform mixture proposed by the protocol.
Prolonged vacuum drying at mild temperatures effectively reduces the amount of moisture and other volatiles in each sample. Notably, small quantities of water do not affect the final spectrum because&#160;TMDP is added in excess. In addition, in some cases, a small amount of 2-hydroxy-4,4&#39;-5,5&#39;-tetramethyl-1,3,2-dioxaphospholane may result from the humidity present in the NMR tube or the sample vial. In these cases, stirring is sufficient to dissolve the amount of the formed precipitate completely. If a high amount of 2-hydroxy-4,4&#39;-5,5&#39;-tetramethyl-1,3,2-dioxaphospholane is formed, it is suggested to repeat the sample preparation, improving the drying treatment. For instance, before use, all glassware can be briefly heated with a heat gun.
The spectral range used to record the spectrum is broad compared to the region of interest for the signal regarding the different hydroxyl groups. However, this is compulsory to understand whether the sample derivatization occurred successfully. The confirmation of complete sample derivatization is given by the presence of a strong signal around 174 ppm. This sharp peak is due to the unreacted TMDP, and its existence ensures that the reagent was present in excess, and therefore, all hydroxyl groups have been derivatized. If this peak is absent, the two most probable causes are: (1) the amount of TMDP used is insufficient to perform the complete derivatization of the sample, or (2) a high amount of water is present in the sample. In the first case, using a higher amount of TMDP would likely ensure the sample&#39;s complete derivatization, and the signal at 174 ppm will appear. In the second case, the sample should be dried more extensively. Once an excess of TMDP is ensured, peak integration can be performed. Before this operation, zoom to a narrower window (150 to 132 ppm) that confines the signals of interest.
The amount of sample (~30 mg) to be analyzed, reported in the above experimental protocol, has been selected to collect good quality spectra for a 300 MHz NMR spectrometer or more. Nevertheless, we have observed that it is possible to reduce the sample amount if a 500 MHz or higher field magnet is used. For instance, in Figure 8D, the NMR spectrum (resulting from a 700 MHz instrument) of a sample prepared with 7.2 mg of lignin is shown. Signal integration of this spectrum offers the same results as those obtained when using higher amounts of lignin. This fact amplifies the applicability of this protocol for all the research in which small quantities of products are available.
Overall, this experimental protocol can be applied to many research and development applications when understanding the origin and fate of the various hydroxy groups present in lignins and tannins is required. In particular, when coupled with GPC and HSQC data, the resulting data offers the opportunity to further elaborate and speculate over the structure of lignin or a tannin. In many instances where chemical modifications are applied to the hydroxy groups of lignin or a tannin, quantitative 31P NMR analyses can be extremely valuable to detect whether these modifications occurred and to which degree. For instance, Figure 9 shows two NMR spectra of the same lignin before and after its oxidation. A simple qualitative evaluation shows the reduction of both&#160;aliphatic and aromatic hydroxy groups upon oxidation, thus providing valuable information and guidance.
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/62696/62696fig09.jpg" /> 
Figure 9: Quantitative 31P NMR spectra of the same Organosolv lignin derivatized using TMDP (A) Prior and (B) post its oxidation. The spectra were recorded using a 300 NMR spectrometer. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In conclusion, this technique has all the attributes of being among the most essential and powerful tools when inquiries dealing with polyphenolic, OH bearing lignins, and tannins (and even synthetic polymers)49,50,51 need to be made in a variety of fields, ranging from chemistry to engineering, from biology to polymer, and pharmaceutical applications.

This procedure, which was recently published in Nature Protocols1&#160;has been cited over 3,000 times in the archival literature and has become a routine measurement for lignin and tannin characterization since it provides essential, rapid, and reproducible structural information.
Lignin and tannins 
When Green Chemistry was introduced by Paul T. Anastas and John C. Werner2,3, it drastically changed the general conception of Chemistry. In particular, the importance of employing sustainable materials instead of fossil feedstocks, such as oil and coal, as a starting point is highlighted as a crucial aspect2,3. Among the different types of biomass, lignin is the most abundant aromatic biopolymer and can be seen as a potential source for industrial commodities and high-added value products4.
Lignin is the second most abundant wood-constituent (with cellulose being first and hemicellulose third). Its content in plants varies depending on the plant-type: for example hardwoods characterized by a lower amount of lignin compared to softwoods (20% &#177; 4% vs. 28% &#177; 4%). In addition, lignin distribution within vegetable tissue is not homogeneous: the higher lignin content can be found in the cell wall5,6. Lignin is a polyphenolic material industrially obtained as a by-product of the paper/cellulose industry7.&#160;It is recovered from the wood pulping process, in which wood chips are primarily processed in the presence of OH- and/or OH- + HS- ion conditions to separate cellulose from hemicellulose and lignin (Soda and/or Kraft processes)8,9.
The first attempts to study lignin were made by Payen and Schultze, respectively, in 1838 and 186510. In 1977, Adler summarized all the relevant available knowledge of that time11. It is currently recognized that the lignin building blocks are&#160;three phenyl-propanoid&#160;units: p-coumaryl, coniferyl, and sinapyl alcohols. These monomers, thanks to a&#160;free radical polymerization process, give rise to p-hydroxyphenyl, guaiacyl, and sinapyl units that eventually broadly constitute lignin (Figure 1)12. The lack of a primary structure in lignins implies an inherent difficulty for its structural characterization. Accordingly, the evaluation of the distribution of molecular weight has always been somewhat controversial. Milled wood lignin, the lignin isolated under mild conditions that approximate mostly protolignin10, is composed of oligomers13 which highly interact via supramolecular aggregation processes14,15.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62696/62696fig01.jpg" /> 
Figure 1:&#160;A representative model of softwood lignin in which the different types of bonds are highlighted. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Lignins are commonly classified depending on: (a) the type of wood from which they are derived (e.g., hardwood and softwood), (b) the process used to isolate it. The most crucial industrial lignin types are Kraft, Lignosulfonates, and Organosolv.
The structure of lignin is highly dependent upon its origin and processing chemistry. More specifically, when the rather complex and irregular structure of lignin is compounded with its natural diversity and the complex processing chemistries, a material of extreme variability, diversity, and heterogeneity emerges, limiting its use to low-value applications16. While softwood lignins contain mainly guaiacyl units (G) with negligible amounts of p-hydroxyphenyl groups (G lignin), hardwood lignins are composed by guaiacyl and syringyl subunits (GS lignin) in varying ratios and grass lignins are constituted by guaiacyl, syringyl, and p-hydroxyphenyl (GSH lignin) subunits. The extractive approach used for isolation&#160;dramatically affects the structure of the emerging lignin17. Figure 2 depicts three lignin structures, differing by the isolation approach employed. Some considerations regarding the effect of the extraction method could be highlighted. Firstly, Kraft lignin is a dealkylated, highly fragmented, and condensed lignin, while Organosolv lignin has a structure similar to milled wood lignin (isolated using the Bjorkman approach)18,19,20. Finally, lignosulfonates are characterized by a high degree of sulfonation, depending on the intensity and the conditions of the extractive sulfonation process.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62696/62696fig02.jpg" /> 
Figure 2: Representative structures for technical lignins. In this figure, the differences among the different types of lignin can be seen. (A) Softwood Kraft lignin is highly condensed, (B) lignosulfonates are characterized by sulfonic groups on saturated carbons, and (C) organosolv lignin has a structure similar to the one of milled wood lignin. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Similar to lignins, tannins are polyphenolic compounds that are found in plants. A recent and updated review on tannins&#39; extractive approaches and applications was recently released by Das et al.21. The importance of tannins in everyday life can be highlighted considering two examples: they impart taste and color to wines22;&#160;furthermore their poly-phenolic structure offers antioxidant characteristics and makes them ideal for application in the tanning industry23. Tannins are divided into two classes: hydrolyzable and non-hydrolyzable. Hydrolyzable tannins can be considered a polymer of gallic, di-gallic, and ellagic acid esters (Figure 3). These esters result from the esterification of the phenolic acids with sugar molecules (e.g., glucose, rhamnose, and arabinose).
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62696/62696fig03.jpg" /> 
Figure 3: Typical hydrolysable tannins: tannic acid, vescalgin. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Non-hydrolyzable tannins, also known as condensed tannins, are polymers and oligomers deriving from flavan-3-ols. Among flavan-3-ols, catechins and gallocatechin are the most frequent. They are colorless crystalline compounds (Figure 4). The polymerization creates a polymer characterized by a helicoidal structure. The aromatic hydroxy groups are directed on the exterior of the helix, while the pyran oxygens are in the interior.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62696/62696fig04.jpg" /> 
Figure 4: Proantocyanidin structures: R =H, OH, OCH3.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Characterization of lignins and tannins using NMR 
Two types of information are crucial in lignin or tannin characterization: (a) chemical structure (e.g., hydroxy group content, nature, and frequency of interunit linkages) and (b) molecular weight and polydispersity. Since the early studies on lignin, different techniques have been employed to achieve these goals, and two classes of methods have emerged: chemical and physical methods.
In lignin chemistry, chemical methods, such as alkaline nitrobenzene oxidation, derivatization followed by reductive cleavage, permanganate oxidation, and thioacidolysis, have been historically widely used24,25,26,27,28,29. However, even if the analytical protocols have been implemented and optimized, they are time-demanding, laborious, and require extensive experimental skills30. Alternatively, from the beginning of the instrumental analysis, physical methods have been used to perform lignin and tannin characterizations31. These techniques allow overcoming the problems of classical methods making it easy to characterize lignin structure.
Nuclear Magnetic Resonance (NMR) allows obtaining information about lignin structure and chemical composition among the instrumental techniques. In particular, data from quantitative monodimensional 1H NMR spectra and quantitative 13C NMR spectra can provide information about different types of lignin interunit bondings32,33,34,35. Unfortunately, monodimensional spectra suffer from signal overlap, which can seriously undermine signal integration efforts. Quantitative versions of HSQC (Heteronuclear Single Quantum Coherence), Q-HSQC (Quantitative - Heteronuclear Single Quantum Coherence), have been used to understand lignin structure better, providing helpful information about internal linkages. However, they cannot be fully utilized to determine the various buildings units13,36,37 quantitatively.
To overcome the issues associated with mono- and two-dimensional NMR, substrate derivatization has been considered. Among the advantages of this approach is that specific labels can be introduced within the complex macromolecule and no spectral interference results from the solvent in which the labeled substrates are dissolved1. Verkade was the pioneer in this field, performing 31P NMR analysis of phosphorous derivatives, coal derivatives, and related compounds38. In its publication, a screening of different phosphorus-containing reagents (phospholanes) was performed, and the chemical shift of other labeled compounds was recorded. Argyropoulos&#39; team first introduced derivatization for the quantitative and qualitative analysis of hydroxy groups in lignin in 1991. After studying the derivatization of lignin model compounds using phosphorus-containing reagents, his group paved the way for one of the most daily-used techniques in lignin chemistry, 31P NMR analysis39,40,41,42,43. Among the different phospholanes examined, Argyropoulos arrived at the use of 2-chloro-4,4,5,5-tetramethyl-1,3-2-dioxaphospholane (TMDP) as being the most suitable one to perform lignin analysis44. TMDP selectively reacts with hydroxy groups causing the quantitative formation of phosphorus-containing derivatives characterized by specific 31P NMR chemical shifts (Figure 5).
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62696/62696fig05.jpg" /> 
Figure 5: Lignin and tannin phosphytilation chemistry. Labeling lignin and tannin labile H groups is accomplished by in situ reaction. The labeled polyphenols are characterized by specific 31P NMR bands corresponding to the different type of hydroxy groups. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Sample derivatization is performed in a pyridine/chloroform (1.6:1) mixture; this choice results from an accurate evaluation. Pyridine has two advantages. Firstly, selecting a solvent characterized by a Hildebrand parameter of about 22.1 MPa1/2 simplifies and amplifies lignin solubilization45. Consequently, the addition of pyridine as a solvent, whose Hildebrand parameter equals 21.7, is thus optimal. Secondly, the reaction of TMDP with hydroxy groups is accompanied by the formation of hydrochloric acid (HCl) as a by-product with concomitant negative implications toward the facile formation of lignin-phospholane derivatives. For this reason, the resulting HCl needs to be neutralized. When present in significant excess, the basicity of the pyridine, relative to TMDP, allows for the neutralization of the HCl (via the formation of pyridine hydrochloride).
The use of the recommended pyridine/deuterated chloroform binary solvent system is based on three reasons. Firstly, it favors sample dissolution. Secondly, as pyridine hydrochloride is soluble in chloroform, it can prevent precipitation and deterioration of the final spectrum. Thirdly, deuterated chloroform is chosen for its unique singlet signal, allowing locking of the NMR spectrometer during the acquisition process. Sample derivatization is performed in the presence of an internal standard. In this way, when the sample and the standard are derivatized, the comparison of the integrals of the peaks of the sample and the standard allows the quantification of the amount for each type of hydroxy group present. Various compounds have been considered as internal standards. These compounds are characterized by a single hydroxy group per molecule, offering a single sharp signal in the 31P NMR spectrum after derivatization. The selection of the standard must be made carefully. Its signal should not overlap with those of the derivatized sample. Cholesterol was widely used during the early days. However, a partial overlap with signals arising from aliphatic hydroxy group limits its use. For routine analysis, internal standard solutions of N-hydroxy-5-norbornene-2,3-dicarboximide (NHND) are preferred. However, owing to NHND instability, its standard solutions can be stored only for a few days46.

<table><tbody><tr><td>100 - 1000 &amp;micro;l Eppendorf micropipette</td><td>VWR</td><td>613-0866</td><td></td></tr><tr><td>20 - 200 &amp;micro;l Eppendorf micropipette</td><td>VWR</td><td>613-0865</td><td></td></tr><tr><td>2-chloro-4,4,5,5-tetramethyl-1,3-2-dioxaphospholane, 95%</td><td>Sigma-Aldrich</td><td>447536</td><td></td></tr><tr><td>Analytical balance (sensibility &amp;plusmn; 0.1 mg)</td><td>Precisa</td><td>LX220 A</td><td></td></tr><tr><td>Binder Vacuum Oven</td><td>Binder</td><td>VD53</td><td></td></tr><tr><td>Certified Vial Kit, Low Adsorption (LA), 2 mL, pk of 100</td><td>Sigma-Aldrich</td><td>29651-U</td><td></td></tr><tr><td>Chloroform-d</td><td>Sigma-Aldrich</td><td>151823</td><td></td></tr><tr><td>Cholesterol, Sigma-grade</td><td>Sigma-Aldrich</td><td>C8667</td><td></td></tr><tr><td>Molecular sieves, 4A</td><td>Sigma-Aldrich</td><td>208604</td><td></td></tr><tr><td>N-hydroxy-5-norbornene-2,3-dicarboximide, 97%</td><td>Sigma-Aldrich</td><td>226378</td><td></td></tr><tr><td>NMR spectrometer, 300 MHz</td><td>Bruker</td><td></td><td></td></tr><tr><td>Norell&amp;nbsp;natural quartz 3 mm NMR tubes</td><td>Sigma-Aldrich</td><td>NORS33007</td><td></td></tr><tr><td>Pipette tips, 100-1000 &amp;micro;L UltraFine (blue)</td><td>VWR</td><td>613-0342</td><td></td></tr><tr><td>Pipette tips, 20-200 &amp;micro;L Bevel Point (yellow)</td><td>VWR</td><td>613-0239</td><td></td></tr><tr><td>Pyridine, anhydrous, 99.8%</td><td>Sigma-Aldrich</td><td>270970</td><td></td></tr><tr><td>Stirring bars,micro, 3 mm lenght</td><td>VWR</td><td>442-0360</td><td></td></tr><tr><td>Stirring bars,micro, 6 mm lenght</td><td>VWR</td><td>442-0362</td><td></td></tr><tr><td>Triphenylphospine oxide, 97%</td><td>Sigma-Aldrich</td><td>T84603</td><td></td></tr><tr><td>Vials for environmental analysis, WHEATON,&amp;nbsp; 20.00 mL</td><td>DWK Life Sciences</td><td>WHEAW224609</td><td></td></tr><tr><td>Weighing paper, grade 531</td><td>VWR</td><td>516-0318P</td><td></td></tr></tbody></table>

quantitative 31p nmr analysis of lignins and tannins

The development of sustainable biorefinery products is confronted, among others, with the challenge of lignin and tannin valorization. These abundant, renewable aromatic biopolymers have not been widely exploited due to their inherent structural complexity and high degrees of variability and species diversity. The lack of a defined primary structure for these polyphenols is further compounded with complex chemical alterations induced during processing, eventually imparting a large variety of structural features of extreme significance for any further utilization efforts.
Consequently, a protocol for the rapid, simple, and unequivocal identification and quantification of the various functional groups present in natural polyphenols, is a fundamental prerequisite for understanding and accordingly tailor their reactivity and eventual utility.
Quantitative 31P NMR offers the opportunity to rapidly and reliably identify unsubstituted, o-mono substituted, and o-disubstituted phenols, aliphatic OHs, and carboxylic acid moieties in lignins and tannins with broad application potential.
The methodology consists of an in situ quantitative lignin or tannin labeling procedure using a suitable 31P containing probe, followed by the acquisition of a quantitative 31P NMR spectrum in the presence of an internal standard. The high natural abundance of the 31P nucleus allows for small amounts of the sample (~30 mg) and short NMR acquisition times (~30-120 min) with well-resolved 31P signals that are highly dependent on the surrounding chemical environment of the labeled OH groups.

The described protocol can be applied both for the analysis of lignins and&#160;tannins. In lignin chemistry, this method is fundamental because it allows the detection and quantification of the different types of hydroxy groups. Figure 8A-D show examples of 31P NMR spectra of lignins and tannins acquired with spectrometers working at different frequencies. The spectrum shown in Figure 8A was recorded using a 300 MHz NMR spectrometer, while Figure 8D was recorded with a 700 MHz NMR instrument.
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/62696/62696fig8v2.jpg" /> 
Figure 8: Quantitative 31P NMR spectrum of (A) softwood kraft lignin (spectrum recorded with a 300 MHz spectrometer on 30.8 mg of lignin), (B) softwood lignosulfonic acid (spectrum recorded with a 300 MHz&#160;spectrometer on 30.1 mg of lignin after conservation of lignosulfonate to lignosulfonic acid), (C) Acacia tannin (spectrum recorded with a 300 MHz spectrometer on a 30.3 mg sample) and (D) softwood kraft lignin (spectrum recorded with a 700 MHz spectrometer on 7.2 mg of lignin). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62696/62696fig8v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
These spectra were carefully recorded and processed manually. The typical signals for aliphatic (150-145 ppm), aromatic (145-137 ppm), and carboxylic (136-134 ppm) hydroxy groups are very well resolved and as such readily integrated. If the spectral window is opened (from 95 to 190 ppm, Figure 8), three sharp, strong peaks (175, 144, and 132 ppm) are apparent. Those are due to the excess of TMDP, the internal standard (cholesterol or NHND), and the hydroxylated-TMDP (caused by water traces), respectively.
In contrast to kraft and organosolv lignin, lignosulfonates are insoluble in the pyridine/chloroform mixture. To obtain a reliable 31P NMR spectrum, solubility is mandatory. To overcome this issue, lignosulfonates can be converted to the corresponding lignosulfonic acids before derivatization. Treating lignosulfonate solutions with strong acids (i.e., sulfuric acid), or acid exchange resins (e.g., Dowex 1H, a strong acid cation exchanger) drives the conversion of all the sulfonate groups in their acidic forms. The resulting products can be removed from the acidic solution using selective adsorptive resins (XAD-7, a polar adsorbent used to isolate compounds characterized by molecular weights up to 60,000 u.m.a) analyzed using this protocol. Figure 8B shows the quantitative 31P NMR spectrum of a TMDP derivatized lignosulfonic acid. Even in this case, the different signals of the hydroxy groups are evident. Figure 8C shows a typical quantitative 31P NMR spectrum of a tannin sample derivatized using TMDP. A characteristic signal from the different aliphatic OH (Ring C), pyrogallol, and catechol units in ring B and units in ring A are well visible.

Watch this Scientific Journal Video about Quantitative 31P NMR Analysis of Lignins and Tannins at JoVE.com

Quantitative 31P NMR Analysis of Lignins and Tannins

31P NMR is a powerful tool for the structural elucidation of polyphenols. This fast, easy, precise, quantitative, and highly reproducible analytical procedure, that allows for the quantification and differentiation of the different types of hydroxy, phenolic, and carboxylic groups in lignins and tannins has now become a routine analytical tool.

Quantitative 31P NMR Analysis of Lignins and Tannins

The development of sustainable biorefinery products is confronted, among others, with the challenge of lignin and tannin valorization. These abundant, ...

quantitative-31p-nmr-analysis-of-lignins-and-tannins

Nanyang Technological University

Research

JoVE Journal

Chemistry

12.0K Views.  Ca’ Foscari University of Venezia. 31P NMR is a powerful tool for the structural elucidation of polyphenols. This fast, easy, precise, quantitative, and highly reproducible analytical procedure, that allows for the quantification and differentiation of the different types of hydroxy, phenolic, and carboxylic groups in lignins and tannins has now become a routine analytical tool.

Video: Quantitative 31P NMR Analysis of Lignins and Tannins

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 ...

Environment

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

A Novel Method for the Pentosan Analysis Present in Jute Biomass and Its Conversion into Sugar Monomers Using Acidic Ionic Liquid

Recently, ionic liquids (ILs) are used for biomass valorization into valuable chemicals because of their remarkable properties such as thermal stability, lower vapor pressure, non-flammability, higher heat capacity, and tunable solubility and acidity. Here, we demonstrate a method for the synthesis of C5 sugars (xylose and arabinose) from the pentosan present in jute biomass in a one-pot process by utilizing a catalytic amount of Br&#248;nsted acidic 1-methyl-3-(3-sulfopropyl)-imidazolium hydrogen sulfate IL. The acidic IL is synthesized in the lab and characterized using NMR spectroscopic techniques for understanding its purity. The various properties of BAIL are measured such as acid strength, thermal and hydrothermal stability, which showed that the catalyst is stable at a higher temperature (250 &#176;C) and possesses very high acid strength (Ho 1.57). The acidic IL converts over 90% of pentosan into sugars and furfural. Hence, the presenting method in this study can also be employed for the evaluation of pentosan concentration in other kinds of lignocellulosic biomass.

We present a protocol for the synthesis of C5 sugars (xylose and arabinose) from a renewable non-edible lignocellulosic biomass (i.e., jute) with the presence of Br&#248;nsted acidic ionic liquids (BAILs) as the catalyst in water. The BAILs catalyst exhibited better catalytic performance than conventional mineral acid catalysts (H2SO4 and HCl).

Recently, ionic liquids (ILs) are used for biomass valorization into valuable chemicals because of their remarkable properties such as thermal stability, ...

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 ...

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

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 ...

Quantitative ³¹P NMR Analysis of Lignins and Tannins

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

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

A Novel Method for the Pentosan Analysis Present in Jute Biomass and Its Conversion into Sugar Monomers Using Acidic Ionic Liquid

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

Extraction of Lignin with High β-O-4 Content by Mild Ethanol Extraction and Its Effect on the Depolymerization Yield

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

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

Fractionation of Lignocellulosic Biomass using the OrganoCat Process

Ultrafast Lignin Extraction from Unusual Mediterranean Lignocellulosic Residues

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