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/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/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/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/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/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/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/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/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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100 - 1000 &#181;l Eppendorf micropipette</td>
			<td>VWR</td>
			<td>613-0866</td>
			<td></td>
		</tr>
		<tr>
			<td>20 - 200 &#181;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 &#177; 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&#160;natural quartz 3 mm NMR tubes</td>
			<td>Sigma-Aldrich</td>
			<td>NORS33007</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips, 100-1000 &#181;L UltraFine (blue)</td>
			<td>VWR</td>
			<td>613-0342</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips, 20-200 &#181;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,&#160; 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/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

Universidad San Sebastian

Research

JoVE Journal

Chemistry

11.9K 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

Structure and Coordination Determination of Peptide-metal Complexes Using 1D and 2D 1H NMR

Copper (I) binding by metallochaperone transport proteins prevents copper oxidation and release of the toxic ions that may participate in harmful redox reactions. The Cu (I) complex of the peptide model of a Cu (I) binding metallochaperone protein, which includes the sequence MTCSGCSRPG (underlined is conserved), was determined in solution under inert conditions by NMR spectroscopy.
NMR is a widely accepted technique for the determination of solution structures of proteins and peptides. Due to difficulty in crystallization to provide single crystals suitable for X-ray crystallography, the NMR technique is extremely valuable, especially as it provides information on the solution state rather than the solid state. Herein we describe all steps that are required for full three-dimensional structure determinations by NMR. The protocol includes sample preparation in an NMR tube, 1D and 2D data collection and processing, peak assignment and integration, molecular mechanics calculations, and structure analysis. Importantly, the analysis was first conducted without any preset metal-ligand bonds, to assure a reliable structure determination in an unbiased manner.

Structure and Coordination Determination of Peptide-metal Complexes Using 1D and 2D 1H NMR

The NMR-solution structure of a metallochaperone model peptide with Cu (I) was determined, and a detailed protocol from sample preparation and 1D and 2D data collection to a three-dimensional structure is described.

Copper (I) binding by metallochaperone transport proteins prevents copper oxidation and release of the toxic ions that may participate in harmful redox ...

Synthesis and Purification of Iodoaziridines Involving Quantitative Selection of the Optimal Stationary Phase for Chromatography

The highly diastereoselective preparation of cis-N-Ts-iodoaziridines through reaction of diiodomethyllithium with N-Ts aldimines is described. Diiodomethyllithium is prepared by the deprotonation of diiodomethane with LiHMDS, in a THF/diethyl ether mixture, at -78 &#176;C in the dark. These conditions are essential for the stability of the LiCHI2 reagent generated. The subsequent dropwise addition of N-Ts aldimines to the preformed diiodomethyllithium solution affords an amino-diiodide intermediate, which is not isolated. Rapid warming of the reaction mixture to 0 &#176;C promotes cyclization to afford iodoaziridines with exclusive cis-diastereoselectivity. The addition and cyclization stages of the reaction are mediated in one reaction flask by careful temperature control.
Due to the sensitivity of the iodoaziridines to purification, assessment of suitable methods of purification is required. A protocol to assess the stability of sensitive compounds to stationary phases for column chromatography is described. This method is suitable to apply to new iodoaziridines, or other potentially sensitive novel compounds. Consequently this method may find application in range of synthetic projects. The procedure involves firstly the assessment of the reaction yield, prior to purification, by 1H NMR spectroscopy with comparison to an internal standard. Portions of impure product mixture are then exposed to slurries of various stationary phases appropriate for chromatography, in a solvent system suitable as the eluent in flash chromatography. After stirring for 30 min to mimic chromatography, followed by filtering, the samples are analyzed by 1H NMR spectroscopy. Calculated yields for each stationary phase are then compared to that initially obtained from the crude reaction mixture. The results obtained provide a quantitative assessment of the stability of the compound to the different stationary phases; hence the optimal can be selected. The choice of basic alumina, modified to activity IV, as a suitable stationary phase has allowed isolation of certain iodoaziridines in excellent yield and purity.

A protocol for the diastereoselective one-pot preparation of cis-N-Ts-iodoaziridines is described. The generation of diiodomethyllithium, addition to N-Ts aldimines and cyclization of the amino gem-diiodide intermediate to iodoaziridines is demonstrated. Also included is a protocol to rapidly and quantitatively assess the most appropriate stationary phase for purification by chromatography.

The highly diastereoselective preparation of cis-N-Ts-iodoaziridines through reaction of diiodomethyllithium with N-Ts aldimines is described. ...

A Quantitative Glycomics and Proteomics Combined Purification Strategy

There is a growing desire in the biological and clinical sciences to integrate and correlate multiple classes of biomolecules to unravel biology, define pathways, improve treatment, understand disease, and aid biomarker discovery. N-linked glycosylation is one of the most important and robust post-translational modifications on proteins and regulates critical cell functions such as signaling, adhesion, and enzymatic function. Analytical techniques to purify and analyze N-glycans have remained relatively static over the last decade. While accurate and effective, they commonly require significant expertise and resources. Though some high-throughput purification schemes have been developed, they have yet to find widespread adoption and often rely on the enrichment of glycopeptides. One promising method, developed by Thomas-Oates et al., filter aided N-glycan separation (FANGS), was qualitatively demonstrated on tissues. Herein, we adapted FANGS to plasma and coupled it to the individuality normalization when labeling with glycan hydrazide tags strategy in order to achieve accurate relative quantification by liquid chromatography mass spectrometry and enhanced electrospray ionization. Furthermore, we designed new functionality to the protocol by achieving tandem, shotgun proteomics and glycosylation site analysis on hen plasma. We showed that N-glycans purified on filter and derivatized by hydrophobic hydrazide tags were comparable in terms of abundance and class to those by solid phase extraction (SPE); the latter is considered a gold standard in the field. Importantly, the variability in the two protocols was not statistically different. Proteomic data that was collected in-line with glycomic data had the same depth compared to a standard trypsin digest. Peptide deamidation is minimized in the protocol, limiting non-specific deamidation detected at glycosylation motifs. This allowed for direct glycosylation site analysis, though the protocol can accommodate 18O site labeling as well. Overall, we demonstrated a new in-line high-throughput, unbiased, filter based protocol for quantitative glycomics and proteomics analysis.

A high-throughput protocol was developed for combined proteomics and glycomics purification and LC-MS/MS quantification in plasma. Deamidation analysis of N-linked glycosylation motifs was specific to deglycosylated sites. Accurate quantitation of N-glycans was achieved by coupling filter aided N-glycan separation to the individuality normalization when labeling with glycan hydrazide tags strategy.

There is a growing desire in the biological and clinical sciences to integrate and correlate multiple classes of biomolecules to unravel biology, define ...

Standards for Quantitative Metalloproteomic Analysis Using Size Exclusion ICP-MS

Metals are essential for protein function as cofactors to catalyze chemical reactions. Disruption of metal homeostasis is implicated in a number of diseases including Alzheimer's and Parkinson's disease, but the exact role these metals play is yet to be fully elucidated. Identification of metalloproteins encounters many challenges and difficulties. Here we report an approach that allows metalloproteins in complex samples to be quantified. This is achieved using size exclusion chromatography coupled with inductively coupled plasma - mass spectrometry (SEC-ICP-MS). Using six known metalloproteins, the size exclusion column can be calibrated and the respective trace elements (iron, copper, zinc, cobalt, iodine) can be used for quantification. SEC-ICP-MS traces of human brain and plasma are presented. The use of these metalloprotein standards provides the means to quantitatively compare metalloprotein abundances between biological samples. This technique is poised to help shed light on the role of metalloproteins in neurodegenerative disease as well as other diseases where imbalances in trace elements are implicated.

Size exclusion chromatography hyphenated with inductively coupled plasma - mass spectrometry (ICP-MS) is a powerful tool to measure changes in the abundance of metalloproteins directly from biological samples. Here we describe a set of metalloprotein standards used to estimate molecular mass and the amount of metal associated with unknown proteins.

Metals are essential for protein function as cofactors to catalyze chemical reactions. Disruption of metal homeostasis is implicated in a number of ...

Spin Saturation Transfer Difference NMR (SSTD NMR): A New Tool to Obtain Kinetic Parameters of Chemical Exchange Processes

This detailed protocol describes the new Spin Saturation Transfer Difference Nuclear Magnetic Resonance protocol (SSTD NMR), recently developed in our group to study processes of mutual-site chemical exchange that are difficult to analyze by traditional methods. As the name suggests, this method combines the Spin Saturation Transfer method used for small molecules, with the Saturation Transfer Difference (STD) NMR method employed for the study of protein-ligand interactions, by measuring transient spin saturation transfer along increasing saturation times (build-up curves) in small organic and organometallic molecules undergoing chemical exchange.
Advantages of this method over existing ones are: there is no need to reach coalescence of the exchanging signals; the method can be applied as long as one signal of the exchanging sites is isolated; there is no need to measure T1 or reach steady state saturation; rate constant values are measured directly, and T1 values are obtained in the same experiment, using only one set of experiments.
To test the method, we have studied the dynamics of the hindered rotation of N,N-dimethylamides, for which much data is available for comparison. The thermodynamic parameters obtained using SSTD are very similar to the reported ones (spin-saturation transfer techniques and line-shape analysis). The method can be applied to more challenging substrates that cannot be studied by previous methods.
We envisage that the simple experimental set up and the wide applicability of the method to a great variety of substrates will make this a common technique amongst organic and organometallic chemists without extensive expertise in NMR.

Spin Saturation Transfer Difference NMR SSTD NMR: A New Tool to Obtain Kinetic Parameters of Chemical Exchange Processes

A detailed protocol describing the SSTD NMR method is presented here to help new users apply this new method to obtain the kinetic parameters of their own systems undergoing chemical exchange.

This detailed protocol describes the new Spin Saturation Transfer Difference Nuclear Magnetic Resonance protocol (SSTD NMR), recently developed in our ...

Application and Methodology of the Non-destructive 19F Time-domain NMR Technique to Measure the Content in Fluorine-containing Drug Products

Here, we describe a protocol developed by our group that uses low-field fluorine-19 (19F) time-domain (TD) nuclear magnetic resonance (NMR) to measure the average content of fluorinated drugs in their formulated drug product forms: tablets or capsules. This method is specific to fluorinated drugs because it detects only the content of fluorine, avoiding interference from the excipients that lack fluorine. The advantages of measuring the active content of fluorinated drugs using low-field 19F TD-NMR versus high-field 19F solid-state (SS) NMR are the simplicity of the method; the low cost; and the non-destructive nature of the technique, with all samples recoverable in intact forms (e.g., powders, tablets, and capsules), making this technique affordable for any laboratory.
We have tested the method with three fluorinated drug products available on the market - cinacalcet, lansoprazole, and ciprofloxacin - with doses ranging from 15 to 500 mg. The results of the analyses, measured by low-field 19F TD-NMR, supported the reported label claims for the average drug content.
Based on the simplicity and reproducibility of the analysis, we envision this methodology being implemented in any laboratory, including manufacturing plants, as a process analytical technology (PAT) tool in the pharmaceutical industry.

Application and Methodology of the Non-destructive 19F Time-domain NMR Technique to Measure the Content in Fluorine-containing Drug Products

A simple and non-destructive technique that measures the average content of drug substances in formulated drug products containing fluorine using low-field fluorine-19 (19F) time-domain (TD) nuclear magnetic resonance (NMR) is presented here. The technique can be applied to the development and manufacturing of drugs in the pharmaceutical industry.

Here, we describe a protocol developed by our group that uses low-field fluorine-19 (19F) time-domain (TD) nuclear magnetic resonance (NMR) to measure the ...

NMR-Based Activity Assays for Determining Compound Inhibition, IC50 Values, Artifactual Activity, and Whole-Cell Activity of Nucleoside Ribohydrolases

NMR spectroscopy is often used for the identification and characterization of enzyme inhibitors in drug discovery, particularly in the context of fragment screening. NMR-based activity assays are ideally suited to work at the higher concentrations of test compounds required to detect these weaker inhibitors. The dynamic range and chemical shift dispersion in an NMR experiment can easily resolve resonances from substrate, product, and test compounds. This contrasts with spectrophotometric assays, in which read-out interference problems often arise from compounds with overlapping UV-vis absorption profiles. In addition, since they lack reporter enzymes, the single-enzyme NMR assays are not prone to coupled-assay false positives. This attribute makes them useful as orthogonal assays, complementing traditional high throughput screening assays and benchtop triage assays. Detailed protocols are provided for initial compound assays at 500 &#956;M and 250 &#956;M, dose-response assays for determining IC50 values, detergent counter screen assays, jump-dilution counter screen assays, and assays in E. coli whole cells. The methods are demonstrated using two nucleoside ribohydrolase enzymes. The use of 1H NMR is shown for the purine-specific enzyme, while 19F NMR is shown for the pyrimidine-specific enzyme. The protocols are generally applicable to any enzyme where substrate and product resonances can be observed and distinguished by NMR spectroscopy. To be the most useful in the context of drug discovery, the final concentration of substrate should be no more than 2-3x its Km value. The choice of NMR experiment depends on the enzyme reaction and substrates available as well as available NMR instrumentation.

NMR-Based Activity Assays for Determining Compound Inhibition, IC50 Values, Artifactual Activity, and Whole-Cell Activity of Nucleoside Ribohydrolases

NMR-based activity assays have been developed to identify and characterize inhibitors of two nucleoside ribohydrolase enzymes. Protocols are provided for initial compound assays at 500 &#956;M and 250 &#956;M, dose-response assays for determining IC50 values, detergent counter screen assays, jump-dilution counter screen assays, and assays in E. coli whole cells.

NMR spectroscopy is often used for the identification and characterization of enzyme inhibitors in drug discovery, particularly in the context of fragment ...

Synthesis of Aptamer-PEI-g-PEG Modified Gold Nanoparticles Loaded with Doxorubicin for Targeted Drug Delivery

Due to drug resistance and toxicity in healthy cells, use of doxorubicin (DOX) has been limited in clinical cancer therapy. This protocol describes the designing of poly(ethylenimine) grafted with polyethylene glycol (PEI-g-PEG) copolymer functionalized gold nanoparticles (AuNPs) with loaded aptamer (AS1411) and DOX through amide reactions. AS1411 is specifically bonded with targeted nucleolin receptors on cancer cells so that DOX targets cancer cells instead of healthy cells. First, PEG is carboxylated, then grafted to branched PEI to obtain a PEI-g-PEG copolymer, which is confirmed by 1H NMR analysis. Next, PEI-g-PEG copolymer coated gold nanoparticles (PEI-g-PEG@AuNPs) are synthesized, and DOX and AS1411 are covalently bonded to AuNPs gradually via amide reactions. The diameter of the prepared AS1411-g-DOX-g-PEI-g-PEG@AuNPs is ~39.9 nm, with a zeta potential of -29.3 mV, indicating that the nanoparticles are stable in water and cell medium. Cell cytotoxicity assays show that the newly designed DOX loaded AuNPs are able to kill cancer cells (A549). This synthesis demonstrates the delicate arrangement of PEI-g-PEG copolymers, aptamers, and DOX on AuNPs that are achieved by sequential amide reactions. Such aptamer-PEI-g-PEG functionalized AuNPs provide a promising platform for targeted drug delivery in cancer therapy.

In this protocol, doxorubicin-loaded AS1411-g-PEI-g-PEG modified gold nanoparticles are synthesized via three-step amide reactions. Then, doxorubicin is loaded and delivered to target cancer cells for cancer therapy.

Due to drug resistance and toxicity in healthy cells, use of doxorubicin (DOX) has been limited in clinical cancer therapy. This protocol describes the ...

Preparation of Nanoparticles for ToF-SIMS and XPS Analysis

Nanoparticles have gained increasing attention in recent years due to their potential and application in different fields including medicine, cosmetics, chemistry, and their potential to enable advanced materials. To effectively understand and regulate the physico-chemical properties and potential adverse effects of nanoparticles, validated measurement procedures for the various properties of nanoparticles need to be developed. While procedures for measuring nanoparticle size and size distribution are already established, standardized methods for analysis of their surface chemistry are not yet in place, although the influence of the surface chemistry on nanoparticle properties is undisputed. In particular, storage and preparation of nanoparticles for surface analysis strongly influences the analytical results from various methods, and in order to obtain consistent results, sample preparation must be both optimized and standardized. In this contribution, we present, in detail, some standard procedures for preparing nanoparticles for surface analytics. In principle, nanoparticles can be deposited on a suitable substrate from suspension or as a powder. Silicon (Si) wafers are commonly used as substrate, however, their cleaning is critical to the process. For sample preparation from suspension, we will discuss drop-casting and spin-coating, where not only the cleanliness of the substrate and purity of the suspension but also its concentration play important roles for the success of the preparation methodology. For nanoparticles with sensitive ligand shells or coatings, deposition as powders is more suitable, although this method requires particular care in fixing the sample.

A number of different procedures for preparing nanoparticles for surface analysis are presented (drop casting, spin coating, deposition from powders, and cryofixation). We discuss the challenges, opportunities, and possible applications of each method, particularly regarding the changes in the surface properties caused by the different preparation methods.

Nanoparticles have gained increasing attention in recent years due to their potential and application in different fields including medicine, cosmetics, ...

Quantitative Atomic-Site Analysis of Functional Dopants/Point Defects in Crystalline Materials by Electron-Channeling-Enhanced Microanalysis

A novel elemental and chemical analysis scheme based on electron-channeling phenomena in crystalline materials is introduced, where the incident high-energy electron beam is rocked with the submicrometric pivot point fixed on a specimen. This method enables us to quantitatively derive the site occupancies and site-dependent chemical information of impurities or intentionally doped functional elements in a specimen, using energy-dispersive X-ray spectroscopy and electron energy-loss spectroscopy attached to a scanning transmission electron microscope, which is of significant interest to current materials science, particularly related to nanotechnologies. This scheme is applicable to any combination of elements even when the conventional Rietveld analysis by X-ray or neutron diffraction occasionally fails to provide the desired results because of limited sample sizes and close scattering factors of neighboring elements in the periodic table. In this methodological article, we demonstrate the basic experimental procedure and analysis method of the present beam-rocking microanalysis.

We provide a general outline of quantitative microanalysis methods for estimating the site occupancies of impurities and their chemical states by taking advantage of electron-channeling phenomena under incident electron beam-rocking conditions, which reliably extract information from minority species, light elements, oxygen vacancies, and other point/line/planar defects.