The PACE method was developed and optimized by various members of the Dupree group (University of Cambridge, UK) over the years, and we appreciate all their contributions. This work was funded as part of the DOE Joint BioEnergy Institute (http://www.jbei.org) supported by the U. S. Department of Energy, Office of Science, Office of Biological and Environmental Research, through contract DE-AC02-05CH11231 between Lawrence Berkeley National Laboratory and the U. S. Department of Energy. We also thank Thea Pick and Vy Ngo for help with preparing the csla9 samples.

1. Harvesting of Plant Material
<ol>
	<li>Harvest fresh plant material (~20 mg) and immediately submerge it in 96% (v/v) ethanol and incubate at 70 &#176;C for 30 min; this deactivates any cell wall degrading enzymes present. For dry material, start at step 2. 
	CAUTION: Ethanol is flammable. 
	Note: A single stem or rosette leaf will provide enough material for analysis. However, fewer errors arise if a larger amount of tissue is pooled and analyzed since this is easier to weigh out and handle.</li>
	<li>Carefully record tissue type and developmental stage, as polysaccharide structure varies with both. For example, with Arabidopsis thaliana (used here), stage the tissue according to the methods from Boyes et al.15 
	Note: In this protocol, we have used the lower half of the inflorescence stem, where the first silique is fully elongated but not yellowing.</li>
</ol>
2. Preparation of Alcohol Insoluble Residue (AIR)
<ol>
	<li>Prepare AIR, following the method of Mortimer et al.9 or another similar method, in order to produce a powder lacking soluble sugars, such as sucrose and starch.</li>
</ol>
3. Aliquoting and Pre-treatment of AIR
Note: The exact quantity of AIR needed for each sample will depend upon (a) the abundance of the polysaccharide of interest in the material and (b) the average size of oligosaccharides released by the GH. The method adds a single fluorescent molecule to the reducing end of each oligosaccharide, so the number of oligosaccharides determines the sensitivity of the experiment. As a guideline, for glucomannan in Arabidopsis stem digested with GH5/GH26 (as described), 500 &#181;g is recommended11, whereas for xylan in Arabidopsis stem digested with GH11, 100 &#181;g is recommended as in Mortimer&#39;s study3.
<ol>
	<li>Weigh AIR (10 - 15 mg) into a 15-mL centrifuge tube, and add H2O to a final concentration of 2 mg/mL. Vortex to achieve an even suspension. If this is problematic, then use a glass homogenizer to assist this process.</li>
	<li>Aliquot 500 &#181;g into microfuge tubes. Dry aliquots using a vacuum centrifuge (see the Table of Materials) overnight at 30 &#176;C.</li>
	<li>Pre-treat AIR with 1 M NaOH (20 &#181;L) for 1 h at room temperature; this de-acetylates the polysaccharides and swells the cellulose microfibrils, disrupting the biomass structure and allowing GH access. 
	NOTE: This step can be skipped for purified polysaccharides. CAUTION &#8211; strong acid/base.
	<ol>
		<li>Include an aliquot for a no-enzyme AIR control (treated the same as all samples, except step 4.1. is excluded).</li>
	</ol>
	</li>
	<li>Add 200 &#181;L of H2O and 20 &#181;L 1 M HCl to neutralize. Test that the pH is ~6-7 by removing 1 &#181;L and spotting it onto paper pH indicator strips.</li>
	<li>Add 50 &#181;L 1 M ammonium acetate buffer, pH 6.0 (or whichever pH is appropriate for the GHs used in the study) and H2O to give a total volume of 500 &#181;L. 
	Note: Ammonium acetate is used since it sublimes upon drying, and so it does not add additional salt to the sample. An excess of salt can lead to poor band-shape and resolution on the PACE gel.</li>
</ol>
4. Hydrolysis of AIR with Glycosyl Hydrolases (GHs)
Note: As mentioned in the Discussion, the purity of the GHs is critical. Only use heterologously expressed, affinity purified enzymes. Upon receipt of each lot from the manufacturer, hydrolyze defined aliquots of substrate (e.g., 500 &#181;g AIR) with increasing quantities of GH overnight (e.g., 0.5, 1, 2, 5, 10, 15, 20 &#181;L) (3 units/&#181;L). When there is an excess of GH, the PACE fingerprint will look identical. An excess of enzyme is required to deliver a reproducible result because it must be certain that the hydrolysis reaction is approaching the endpoint.
<ol>
	<li>Add a pre-determined amount (see above) of mannanases (GH5 and GH2611) to the AIR aliquots in buffer from step 3.5, as well as a positive control (30 &#181;g of konjac glucomannan), and a no-AIR negative control (enzyme mix in an empty tube). Vortex, and then spin briefly to collect the reaction mixture in the bottom of the tube.</li>
	<li>Incubate overnight at 37 &#176;C (or the appropriate temperature for the GH of choice) with gentle agitation (~100 rpm).</li>
	<li>Stop the reaction by incubating at 95 &#176;C for 20 min. 
	Note: Cap closures can be used to seal flip-top microfuge tubes.
	<ol>
		<li>Centrifuge using a bench top microfuge at maximum speed for 10 min, and retain supernatant.</li>
	</ol>
	</li>
	<li>Resuspend pellet in 250 &#181;L H2O, centrifuge as above, and retain supernatant.</li>
	<li>Combine both supernatants, and dry in a vacuum centrifuge (see the Table of Materials) at 30 &#176;C (~3 h or overnight without heating).</li>
</ol>
5. Preparation of Oligosaccharide Standards
<ol>
	<li>Prepare 1 mM stock solutions in H2O of mannose (Man), mannobiose (Man2), mannotriose (Man3), mannotetraose (Man4), mannopentaose (Man5) and mannohexaose (Man6), all &#946;,1-4 linked. Aliquot and store at -20 &#176;C until required.</li>
	<li>Prepare 3 different concentrations of a Man1-6 mixture by combining 1 &#181;L (Standard S1), 2 &#181;L (S2) or 5 &#181;L (S3) of all six.</li>
	<li>Dry in a vacuum centrifuge (see Table of Materials) at 30 &#176;C (~1 h).</li>
</ol>
6. Derivitization of Oligosaccharides
<ol>
	<li>Prepare a stock solution of 0.2 M ANTS (8-Aminonaphthalene-1,3,6-trisulfonic acid) in H2O:acetic acid 17:3. Warm stock to 60 &#176;C to completely dissolve the solid. Store at -20 &#176;C, protected from light, for 2 - 3 months.</li>
	<li>Prepare a 0.2 M stock solution of 2-picoline borane (2-PB) in DMSO. This is extremely hygroscopic, so immediately resuspend all powder upon receipt in DMSO. Aliquot, and store at -20 &#176;C for 1 - 2 years. Thaw aliquots as required (store at 4 &#176;C for 2 weeks, and then discard).</li>
	<li>Prepare the derivatization buffer of H2O:acetic acid:DMSO at 17:3:20.</li>
	<li>To each sample, add 5 &#181;L of ANTS, 5 &#181;L of 2-PB and 10 &#181;L of derivatization buffer. Spin briefly to collect in the bottom of the tube, vortex thoroughly, and then spin briefly again. See Figure 1 for reaction description.</li>
	<li>Incubate samples overnight at 37 &#176;C, protected from light.</li>
	<li>Dry in a vacuum centrifuge (see Table of Materials) at 30 &#176;C (~2 h).</li>
	<li>Resuspend samples and standard in 100 &#181;L 3 M urea. Store at -20 &#176;C, protected from the light until required.</li>
</ol>
7. Preparation of PACE gels
Note: Assembly of the gel casting equipment will depend on the brand. Here, we use a vertical electrophoresis system (see Table of Materials), equipped with 18 cm x 24 cm glass plates and 1.5 mm spacers.
<ol>
	<li>Assemble gel casting equipment per the manufacturer&#39;s instructions.</li>
	<li>Make 10x PACE buffer (1M Tris-Borate, pH 8.2) as follows: add 121.14 g of Tris-Base to ~400 mL of H2O, and mix to dissolve. Adjust the pH to 8.2 by addition of solid boric acid (approximately 60 g), and then make volume up to 1 L.</li>
	<li>Make a 10% (w/v) ammonium persulfate (APS) stock in H2O. Aliquot and store at -20 &#176;C. Thaw aliquots once, store at 4 &#176;C and discard after 2 weeks.</li>
	<li>Make and pour the resolving gel. For the above equipment, 1 gel equate to 50 mL. In a 50 mL centrifuge tube, mix H2O (20.2 mL), 5 mL 10x PACE buffer, 24.6 mL 40% acrylamide/Bis-acrylamide (29:1 acrylamide:Bis). (Caution: toxic).
	<ol>
		<li>Add 200 &#181;L of APS and 20 &#181;L of N,N,N,N&#180;-tetramethyl-ethylenediamine (TEMED). Invert gently to mix (to avoid introducing air bubbles). Pour the gel using a serological or other large volume pipette, to ~4 cm below the top of the glass plates. Pay attention to the possibility of air bubbles getting trapped in the gel. If they do, stop pouring, and tilt/tap the gel to release them.</li>
	</ol>
	</li>
	<li>Overlay the gel with isopropanol (Caution: flammable) or, carefully, with H2O. Allow gel to polymerize (20 - 30 mins), then pour off the top layer. If isopropanol was used, then wash 2x with H2O. Dry any excess liquid using blotting paper.</li>
	<li>Make and pour the stacking gel. Mix 6.8 mL H2O, 1 mL 10x PACE buffer, 2.8 mL acrylamide/Bis-acrylamide (Caution: toxic), 80 &#181;L APS, and 8 &#181;L of TEMED in a 15 mL centrifuge tube. Invert to mix, and overlay on top of polymerized resolving gel. Gently insert combs, avoiding trapping air bubbles under the comb teeth.</li>
	<li>Allow gel to polymerize (20-30 min), wrap in moist tissue and then in plastic wrap, and store at 4 &#176;C until required. Store with the combs in place. 
	Note: PACE gels can be stored for a maximum of 2 weeks (although less than 1 week is ideal), if they are kept moist.</li>
</ol>
8. Running a PACE Gel
<ol>
	<li>Use a permanent marker to label the well positions on the glass, which will assist in keeping track of the loading order, and in identifying where the wells are once the comb is removed.</li>
	<li>Remove the comb. Fill the wells with 1x PACE buffer.</li>
	<li>Use a 10 &#181;L microsyringe to load 2 &#181;L of standards and samples into wells. Avoid using the outermost lanes, which tend to run samples poorly.</li>
	<li>Assemble the upper chamber of the gel-running apparatus, and place the gel in a cooled (~10 &#176;C) running tank (10 &#176;C) containing 1x PACE buffer. Fill the upper chamber with 1x PACE buffer.</li>
	<li>Turn on the power, and run the gel at 200 V (constant V) for 30 mins, and then increase voltage to 1000 V for 1 h 40 mins. Protect gels from light (e.g., by wrapping tanks in black garbage bags). 
	Note: Check on the gels ~5 min after turning the voltage on, and then another 5 mins after increasing the voltage to 1,000 V. If the power pack is unable to maintain the voltage then there is likely a buffer leak. Remove the gel from the tank. Reassemble the upper chamber, carefully checking placement of all gaskets. A large chip on the upper edge of the glass plate may prevent the plate forming a good seal with the gasket. This can usually be temporarily resolved by adding a drop of 5 % (w/v) molten agarose to the chipped part of the glass, before adding the upper chamber.</li>
</ol>
9. Visualizing a PACE gel
Note: Use a gel imaging system equipped with long-wave UV bulbs in the transilluminator, and an emission filter for the camera which is suitable for the fluorophore, here 530 nm. Alternatively, a laser scanner may also be used, as described in Goubet2.
<ol>
	<li>Ensure the gel imaging system is dust free by wiping it with moist lint-free tissue.</li>
	<li>Remove the gel from the PACE tank, and view briefly whilst the gel is still in the glass plates (&#60;80 ms) to determine if the dye front is still on the gel.</li>
	<li>Open the gel using a wedge tool, and whilst gel is still on one glass plate, use a pizza cutter to remove both the stacking gel and, if the dye front is still on the gel, the bottom of the gel.</li>
	<li>Put ~5 mL H2O onto the surface of the transilluminator, and then transfer the gel directly onto the transilluminator. Set the filter to UV 605, and turn on the longwave UV transilluminator (see Table of Materials) using the software. 
	Note: The ANTS fluorophore used here has an excitation/emission of 353/520 nm.</li>
	<li>Take several images at various exposure times (e.g., 100 ms to 10 s). Ensure that the UV light is turned off between images by clicking the UV light button (to turn it off) to avoid degrading the fluorescence. Ensure that at least 2 of the images have no saturated bands (the gel imaging software will indicate this).</li>
	<li>Save files as high-resolution.tif images. 
	Note: Images can be analyzed using the software provided with the gel imaging system, or by using freely available image analysis software, such as ImageJ (https://imagej.nih.gov/ij/). See the Results section for a discussion of quantitation.</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+Analysis+of+Cell+Wall+Polysaccharides+Using+PACE.">Structural Analysis of Cell Wall Polysaccharides Using PACE.</a> Methods Mol Biol. 1544, 223-231 (2017).</li><li>Pauly, M., Keegstra, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-wall+carbohydrates+and+their+modification+as+a+resource+for+biofuels.">Cell-wall carbohydrates and their modification as a resource for biofuels.</a> Plant J. 54 (4), 559-568 (2008).</li><li>Lopez-Casado, G., Urbanowicz, B. R., Damasceno, C. M., Rose, J. 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The authors have nothing to disclose.

PACE is a straightforward method for characterizing polysaccharide structure. It can be applied to any polysaccharide for which there are known GHs with characterized activity, see numerous examples in the literature1,6,9,11. Tt has also been applied to the characterization of novel GHs12,13,18 and glycosyltransferases9, by making use of defined polysaccharide substrates and acceptors.
Reproducible, interpretable results are dependent on three key steps. First, the GHs used should be free from contaminating activities. This is best achieved by only using heterologously expressed, affinity purified enzymes. Second, for most experiments, it is important that the enzymatic hydrolysis of the substrates is at completion. This will ensure that the same biomass hydrolyzed with the same GH gives the same fingerprint in every experiment. Finally, there should be an excess of fluorophore in the derivatization reaction. This ensures that the available reducing ends are labelled at close to 100%. This will result in high reproducibility of results, as well as data that is quantitative.
Gel and reagent quality are critical to ensuring reproducible data. Poor quality buffers, especially of incorrect pH, air bubbles in the gel, and samples with an excess of salt can all affect resolution and retention factor of the oligosaccharides. However, inclusion of the recommended controls described in the Protocol and Results section will enable troubleshooting.
PACE is limited by its ability to identify oligosaccharides. For some experiments, a simple fingerprint is all that&#39;s necessary. However, to truly quantitate the amount of glucomannan in a sample of AIR, for example, the identity of all the released oligosaccharides is required, which is a time-consuming process. This is more straightforward for less complex polysaccharides such as xylan3. Since there are few oligosaccharide standards commercially available, it may not always be possible or desirable to identify all the bands. In this case, PACE can provide very complimentary data to mass spectrometry (i.e., MALDI-CID9 or ESI-MS7, and NMR6). For these methods, it is often helpful to do a large-scale preparation, separate by size-exclusion chromatography, and then analyze each fraction by both PACE prior to MS or NMR. Whilst it has been reported that bands can be excised and identified by MALDI-CID, in practice we have found that this has a low rate of success (possibly due to interactions of the labeled oligosaccharides with the acrylamide gel when exposed to UV light).
The other major limitation of PACE is throughput. To have good quality, interpretable gels with the appropriate controls, each gel will only have ~10 experimental samples, and a researcher can expect to run ~4 PACE gels per day. Recently, a version of PACE using capillary electrophoresis (CE) has been developed19, which allows preparation of samples in 96-well plates. It has been used successfully to characterize glycosyltransferase enzyme activities and polysaccharide structures6,19, although it requires access to a CE machine, which can be expensive to purchase and maintain.

Polysaccharide analysis by gel electrophoresis (PACE) is a method for the detailed characterization of polysaccharides1,2,3. Plant cell wall polysaccharides are notoriously difficult to analyze, and most methods require expensive equipment, skilled operators, and large amounts of purified material. Here, we describe a simple method for gaining detailed polysaccharide structural information, including resolution of structural isomers.
Understanding plant cell wall polysaccharide structure is necessary for those researchers exploring plant cell wall biosynthesis or the role of the plant cell wall in plant development. Recently, however, plant cell wall composition has become interesting to a wider group of researchers due to the focus on using plant biomass (essentially the cell wall) as a feedstock to produce biofuels and biochemicals4. As an example, efficient enzymatic saccharification of this material requires a detailed understanding of the polysaccharide structures, so that optimized enzymatic cocktails can be selected5.
PACE has several advantages over alternative methods which make it ideal for the rapid analysis of complex glycan structures. Firstly, it does not require purification of the polysaccharide in question, as is required for solution state nuclear magnetic resonance (NMR)6. Secondly, PACE can resolve structural isomers, unlike mass spectrometry (MS, e.g., matrix-assisted laser desorption/ionization (MALDI) and electrospray ionization (ESI)), which can also be challenging to perform by liquid chromatography (LC)7. Thirdly, PACE is very sensitive, with low-picomole resolution, unlike thin-layer chromatography (TLC) or paper chromatography. Finally, it does not require expensive equipment or specialist knowledge, as is the case for MS, NMR, and LC.
The PACE method relies on the specificity of glycosyl hydrolase (GH) enzymes, which target certain glycosidic linkages in a mixture of polysaccharides. When the GH enzyme acts on the polysaccharide chain, it reveals a reducing end which can then be chemically derivatized, in this case with a fluorescent label. The unhydrolyzed portion of the sample is therefore rendered undetectable by this method. The labeled oligosaccharides are then separated in a large-format acrylamide gel by electrophoresis. This gives excellent resolution of very similar molecules, for example, the trisaccharides Glc-Man-Man and Glc-Man-Glc will have a different Rf.
PACE has been used extensively to characterize different xylan structures across plant species8, to identify glycosyltransferase mutants in Arabidopsis6,9,10,11, to perform glycosyltransferase assays9, and to characterize novel GH activities12,13. We have also recently used it to characterize yeast cell wall mannan (Mahboubi and Mortimer, in preparation). Here we describe a method for characterizing plant cell wall glucomannan structure, based on previous reports11,14.

<table><tbody><tr><td>&lt;strong&gt;Oligosaccaharide Standards&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Mannose</td><td>Sigma-Aldrich</td><td>CAS 3458-28-4</td><td></td></tr><tr><td>Mannobiose</td><td>Megazyme</td><td>CAS: 14417-51-7</td><td></td></tr><tr><td>Mannotriose</td><td>Megazyme</td><td>CAS: 28173-52-6</td><td></td></tr><tr><td>Mannotetraose</td><td>Megazyme</td><td>CAS: 51327-76-5</td><td></td></tr><tr><td>Mannopentaose</td><td>Megazyme</td><td>CAS: 70281-35-5</td><td></td></tr><tr><td>Mannhexaose</td><td>Megazyme</td><td>CAS: 70281-36-6</td><td></td></tr><tr><td>Glucomannan (Konjac; Low Viscosity)</td><td>Megazyme</td><td>P-GLCML</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Other specialty chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>8-Aminonaphthalene-1,3,6-trisulfonic acid</td><td>(Molecular probes) Thermo</td><td>A350</td><td></td></tr><tr><td>2-picoline borane</td><td>TCI</td><td>B3018</td><td></td></tr><tr><td>40 % acrylamide/Bis-acrylamide (29:1 acrylamide:Bis)</td><td>Bio-rad</td><td>1610146</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Specialty Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Gel casting kit</td><td>Hoefer</td><td>SE660 kit</td><td>18x24 cm glass plates, 0.75 mm spacers</td></tr><tr><td>Cooling recirculating water bath&amp;nbsp;</td><td>Hoefer</td><td>RCB20-plus 115v</td><td>Needs to be able to maintain temperature at ~10 C</td></tr><tr><td>G:Box Chemi XRQ Imaging System</td><td>Syngene</td><td>05-GBOX-CHEMI-XRQ</td><td>Order with filters appropriate to fluorphore, and transilluminator should be fitted with long-wave UV light bulbs</td></tr><tr><td>High Voltage Power Pack</td><td>Thermo</td><td>EC1000XL</td><td>1000V&amp;nbsp;</td></tr><tr><td>Vacuum centrifuge(Speedvac)</td><td>Savant</td><td>SPD131</td><td></td></tr><tr><td>Vertical Gel electrophoresis system</td><td>Hoefer</td><td>SE660</td><td></td></tr><tr><td>&lt;strong&gt;Glycosyl hydrolases&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>These can be obtained from companies e.g. Megazyme (https://www.megazyme.com/) or Prozomix (http://www.prozomix.com/), or can be expressed in house by requesting the plasmid from the relevant research group.</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Lab supplies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>15 ml Centrifuge tube ( Falcon Centrifuge Tubes, Polypropylene, Sterile, Corning)</td><td>VWR</td><td>21008-918</td><td></td></tr><tr><td>50 ml Centrifuge tube ( Falcon Centrifuge Tubes, Polypropylene, Sterile, Corning)</td><td>VWR</td><td>21008-951</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Gel imaging software ( Genesys)</td><td>Syngene</td><td></td><td></td></tr></tbody></table>

structural characterization of mannan cell wall polysaccharides in plants using pace

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

Here, we show an example PACE gel run per the protocol, along with descriptions to assist with data interpretation and troubleshooting, and this is followed by a general guide to successful PACE gel interpretation13,16. A representative gel of a standard PACE assay of cell wall mannan content is shown in Figure 2, and is described lane by lane.
Standards 
Lanes 1, 2 and 3 show a ladder of commercially purchased oligosaccharides (Man1- Man6) at 5 (S1), 10 (S2) and 50 (S3) pmol concentration. When receiving a new lot of a commercial oligosaccharide, it is important to check the purity on a gel. Many oligosaccharides, particularly tetrasacccharides and longer, can have significant contamination with oligosaccharides of other degrees of polymerization (DPs), as shown here for Man6 (marked with *). While quantitating a gel, it is important to have at least 3 concentrations of the standards, since it is necessary for calculating the standard curve. Calculation of the standard curve can also be helpful for ensuring that the derivatization reaction is essentially at completion. Standards of known concentrations allow comparisons between gels, and are a useful control for separation quality and derivatization quality.
Positive control 
Lane 4 shows a mannanase digestion of konjac glucomannan. Konjac glucomannan is available commercially, and whilst it does not have the same structure as that, for example, found in Arabidopsis, it serves two purposes. Firstly, it is an important control for the enzymatic digestion. The researcher should establish what a commercial substrate digested with their GHs of choice looks like when digested to completion (i.e., a vast excess of GH is added to the reaction). If in future experiments the pattern changes, this can indicate either a loss of activity or contamination of the GH stock. Secondly, it serves as a control for the derivatization reaction in the presence of the enzyme and buffer salts. If the standards look good, but the positive control is poor, then this may indicate that a component of the hydrolysis reaction is inhibitory to the derivatization.
Wild type (WT) Arabidopsis stem AIR + mannanase cocktail (GH5 + GH26) 
Lane 5 shows the PACE fingerprint of a WT Arabidopsis stem (compare with references11,14).
csla9 Arabidopsis stem AIR + mannanase cocktail (GH5 + GH26) 
Lane 6 shows shows the PACE fingerprint of the stem from an Arabidopsis plant lacking the major stem mannan synthase11. Compare to the WT and negative control fingerprints. Whilst the majority of the mannan is absent in this plant, a small quantity remains, as evidenced by the reduced band intensities for all mannanase-derived oligosaccharides This is synthesized by additional mannan synthases (CSLA2, 3)11.
Negative control - enzyme only 
Lane 8 shows bands on the gel that are not specific that derive from contaminants in the mannanase cocktail, and should be excluded from the analysis (marked with *).
Negative control - WT AIR only 
Lane 9 shows bands on the gel that are not specific and should be excluded from the analysis (marked with *).
Negative control - csla9 AIR only 
Lane 10 shows bands on the gel that are not specific and should be excluded from the analysis (marked with *).
Pine wood AIR + mannanase cocktail (GH5 + GH26) 
Lane 7 shows the PACE fingerprint of pine wood, which contains galactoglucomannan. This clearly has a different pattern of released oligosaccharides to Arabidopsis, due to its different structure.
Negative control - Pine AIR only 
Lane 11 shows bands on the gel that are not specific and should be excluded from the analysis (marked with *).
Interpreting a PACE gel is relatively straightforward, but requires awareness of the following points. For robust data, it is recommended to carry out PACE on at least 3 independently grown biological replicates, and it is recommended to perform at least 2 technical replicates on each biological replicate. The controls described are critical for interpretation, as non-specific bands need to be excluded. We also recommend loading samples in a different order on replicate gels, to exclude effects which result from uneven illumination by the transilluminator. Accurate interpretation requires analysis of bands which are not saturated. Since some oligosaccharide fingerprints may contain both very high and low abundance polysaccharides, we recommend analyzing multiple exposures of the same gel. The standards can be used to normalize between different images.
For some types of experiments, it may be desirable to quantify the amount of polysaccharide in the AIR preparation. Whilst it is possible to quantify from a PACE gel, it requires knowledge of the exact molecular identity of each oligosaccharide released by the GHs and where it is located on the PACE gel. This is currently not fully known for mannan, but it has been determined for other polysaccharides e.g. xylan3. The standards do provide a way of normalizing between gels, even when quantitation is not being performed, and so will assist in any qualitative or semi-quantitative analysis.
Identifying the structure of individual oligosaccharides can be achieved using cocktails of GHs. Sequential addition of further GHs can reveal details about linkages and substitutions of the released oligosaccharides (e.g., addition of &#946;-1,4-glucosidase that acts on the non-reducing end will reveal which oligosaccharides in the mixture contain that feature). Available enzymes include galactosidases, glucuronidases and glucosidases (see the CAZy database16 for additional information); see Hogg, D. et al.17 as an example.
The protocol above can be used to characterize the activity of unknown GHs. In this case, a defined biomass, such as Arabidopsis stem or konjac glucomannan, should be used to screen the unknown GHs13.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56424/56424fig1.jpg" /> 
Figure 1: This shows the scheme for the fluorescent derivatization of oligosaccharides with ANTS. Modified from Goubet2. <a href="https://cloudflare.jove.com/files/ftp_upload/56424/56424fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56424/56424fig2.jpg" /> 
Figure 2: Representative PACE gel showing the mannan fingerprint from Arabidopsis, pine and an Arabidopsis mutant (csla9) which has impaired mannan biosynthesis. AlR was hydrolyzed with a mannanase cocktail, and the released oligosaccharides were derivatized with a fluorophore. The oligosaccharides were separated by gel electrophoresis on the basis of size, shape, and charge. A ladder of manno-oligosaccharides (Man1-6) is shown to assist in identifying relative mobilities of the released oligosaccharides, and a hydrolysis of purified mannan (derived from konjac) is shown as a positive control. * = non-specific bands.&#160;<a href="https://cloudflare.jove.com/files/ftp_upload/56424/56424fig2large.jpg" style="font-size: 14px;" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Structural Characterization of Mannan Cell Wall Polysaccharides in Plants Using PACE at JoVE.com

Structural Characterization of Mannan Cell Wall Polysaccharides in Plants Using PACE

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

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

structural-characterization-mannan-cell-wall-polysaccharides-plants

Research

JoVE Journal

Biochemistry

9.6K Views.  Joint BioEnergy Institute. Een protocol voor de structurele analyse van polysacchariden door elektroforese van het gel (PACE), met behulp van mannan als voorbeeld is beschreven.

Video: Structural Characterization of Mannan Cell Wall Polysaccharides in Plants Using PACE

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 ...

Sequencing of Plant Wall Heteroxylans Using Enzymic, Chemical (Methylation) and Physical (Mass Spectrometry, Nuclear Magnetic Resonance) Techniques

This protocol describes the specific techniques used for the characterization of reducing end (RE) and internal region glycosyl sequence(s) of heteroxylans. De-starched wheat endosperm cell walls were isolated as an alcohol-insoluble residue (AIR)1 and sequentially extracted with water (W-sol Fr) and 1 M KOH containing 1% NaBH4 (KOH-sol Fr) as described by Ratnayake et al. (2014)2. Two different approaches (see summary in Figure 1) are adopted. In the first, intact W-sol AXs are treated with 2AB to tag the original RE backbone chain sugar residue and then treated with an endoxylanase to generate a mixture of 2AB-labelled RE and internal region reducing oligosaccharides, respectively. In a second approach, the KOH-sol Fr is hydrolyzed with endoxylanase to first generate a mixture of oligosaccharides which are subsequently labelled with 2AB. The enzymically released ((un)tagged) oligosaccharides from both W- and KOH-sol Frs are then methylated and the detailed structural analysis of both the native and methylated oligosaccharides is performed using a combination of MALDI-TOF-MS, RP-HPLC-ESI-QTOF-MS and ESI-MSn. Endoxylanase digested KOH-sol AXs are also characterized by nuclear magnetic resonance (NMR) that also provides information on the anomeric configuration. These techniques can be applied to other classes of polysaccharides using the appropriate endo-hydrolases.

Sequencing of Plant Wall Heteroxylans Using Enzymic, Chemical Methylation and Physical Mass Spectrometry, Nuclear Magnetic Resonance Techniques

This protocol describes the specific techniques used for the structural characterization of reducing end (RE) and internal region glycosyl sequence(s) of heteroxylans by tagging the RE with 2 aminobenzamide prior to enzymatic (endoxylanase) hydrolysis and then analysis of the resultant oligosaccharides using mass spectrometry (MS) and nuclear magnetic resonance (NMR).

This protocol describes the specific techniques used for the characterization of reducing end (RE) and internal region glycosyl sequence(s) of ...

Chemistry

Preparation of Fungal and Plant Materials for Structural Elucidation Using Dynamic Nuclear Polarization Solid-State NMR

This protocol shows how uniformly 13C, 15N-labeled fungal materials can be produced and how these soft materials should be proceeded for solid-state NMR and sensitivity-enhanced DNP experiments. The sample processing procedure of plant biomass is also detailed. This method allows the measurement of a series of 1D and 2D 13C-13C/15N correlations spectra, which enables high-resolution structural elucidation of complex biomaterials in their native state, with minimal perturbation. The isotope-labeling can be examined by quantifying the intensity in 1D spectra and the polarization transfer efficiency in 2D correlation spectra. The success of dynamic nuclear polarization (DNP) sample preparation can be evaluated by the sensitivity enhancement factor. Further experiments examining the structural aspects of the polysaccharides and proteins will lead to a model of the three-dimensional architecture. These methods can be modified and adapted to investigate a wide range of carbohydrate-rich materials, including the natural cell walls of plants, fungi, algae and bacteria, as well as synthesized or designed carbohydrate polymers and their complex with other molecules.

A protocol for preparing 13C,15N-labeled fungal and plant samples for multidimensional solid-state NMR spectroscopy and dynamic nuclear polarization (DNP) investigations is presented.

This protocol shows how uniformly 13C, 15N-labeled fungal materials can be produced and how these soft materials should be proceeded for solid-state NMR ...

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

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

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

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

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

Biology

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

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st century. This has rekindled interest in the use of plant products as industrial raw materials for the production of liquid fuels for transportation2 and other products such as biocomposite materials6. Plant biomass remains one of the greatest untapped reserves on the planet4. It is mostly comprised of cell walls that are composed of energy rich polymers including cellulose, various hemicelluloses, and the polyphenol lignin5 and thus sometimes termed lignocellulosics. However, plant cell walls have evolved to be recalcitrant to degradation as walls contribute extensively to the strength and structural integrity of the entire plant. Despite its necessary rigidity, the cell wall is a highly dynamic entity that is metabolically active and plays crucial roles in numerous cell activities such as plant growth and differentiation5. Due to the various functions of walls, there is an immense structural diversity within the walls of different plant species and cell types within a single plant4. Hence, depending of what crop species, crop variety, or plant tissue is used for a biorefinery, the processing steps for depolymerisation by chemical/enzymatic processes and subsequent fermentation of the various sugars to liquid biofuels need to be adjusted and optimized. This fact underpins the need for a thorough characterization of plant biomass feedstocks. Here we describe a comprehensive analytical methodology that enables the determination of the composition of lignocellulosics and is amenable to a medium to high-throughput analysis (Figure 1). The method starts of with preparing destarched cell wall material. The resulting lignocellulosics are then split up to determine its monosaccharide composition of the hemicelluloses and other matrix polysaccharides1, and its content of crystalline cellulose7. The protocol for analyzing the lignin components in lignocellulosic biomass is discussed in Part I3.

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

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

Fluorescent Immunolocalization of Arabinogalactan Proteins and Pectins in the Cell Wall of Plant Tissues

Plant development involves constant adjustments of the cell wall composition and structure in response to both internal and external stimuli. Cell walls are composed of cellulose and non-cellulosic polysaccharides together with proteins, phenolic compounds and water. 90% of the cell wall is composed of&#160;polysaccharides (e.g., pectins) and arabinogalactan proteins (AGPs). The fluorescent immunolocalization of specific glycan epitopes in plant histological sections remains a key tool to uncover remodeling of wall polysaccharide networks, structure and components.
Here, we report an optimized fluorescent immunolocalization procedure to detect glycan epitopes from AGPs and pectins in plant tissues. Paraformaldehyde/glutaraldehyde fixation was used along with LR-White embedding of the plant samples, allowing for a better preservation of the tissue structure and composition. Thin sections of the embedded samples obtained with an ultra-microtome were used for immunolocalization with specific antibodies. This technique offers great resolution, high specificity, and the chance to detect multiple glycan epitopes in the same sample. This technique allows subcellular localization of glycans and detects their level of accumulation in the cell wall. It also permits the determination of spatio-temporal patterns of AGP and pectin distribution during developmental processes. The use of this tool may ultimately guide research directions and link glycans to specific functions in plants. Furthermore, the information obtained can complement biochemical and gene expression studies.

This protocol describes in detail how the plant material for immunolocalization of Arabinogalactan proteins and pectins is fixed, embedded in a hydrophilic acrylic resin, sectioned and mounted on glass slides. We show cell wall related epitopes will be detected with specific antibodies.

Plant development involves constant adjustments of the cell wall composition and structure in response to both internal and external stimuli. Cell walls ...

Developmental Biology

OLIgo Mass Profiling (OLIMP) of Extracellular Polysaccharides

The direct contact of cells to the environment is mediated in many organisms by an extracellular matrix. One common aspect of extracellular matrices is that they contain complex sugar moieties in form of glycoproteins, proteoglycans, and/or polysaccharides. Examples include the extracellular matrix of humans and animal cells consisting mainly of fibrillar proteins and proteoglycans or the polysaccharide based cell walls of plants and fungi, and the proteoglycan/glycolipid based cell walls of bacteria. All these glycostructures play vital roles in cell-to-cell and cell-to-environment communication and signalling.
An extraordinary complex example of an extracellular matrix is present in the walls of higher plant cells. Their wall is made almost entirely of sugars, up to 75% dry weight, and consists of the most abundant biopolymers present on this planet. Therefore, research is conducted how to utilize these materials best as a carbon-neutral renewable resource to replace petrochemicals derived from fossil fuel. The main challenge for fuel conversion remains the recalcitrance of walls to enzymatic or chemical degradation due to the unique glycostructures present in this unique biocomposite.
Here, we present a method for the rapid and sensitive analysis of plant cell wall glycostructures. This method OLIgo Mass Profiling (OLIMP) is based the enzymatic release of oligosaccharides from wall materials facilitating specific glycosylhydrolases and subsequent analysis of the solubilized oligosaccharide mixtures using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF/MS)1 (Figure 1). OLIMP requires walls of only 5000 cells for a complete analysis, can be performed on the tissue itself2, and is amenable to high-throughput analyses3. While the absolute amount of the solubilized oligosaccharides cannot be determined by OLIMP the relative abundance of the various oligosaccharide ions can be delineated from the mass spectra giving insights about the substitution-pattern of the native polysaccharide present in the wall.
OLIMP can be used to analyze a wide variety of wall polymers, limited only by the availability of specific enzymes4. For example, for the analysis of polymers present in the plant cell wall enzymes are available to analyse the hemicelluloses xyloglucan using a xyloglucanase5, 11, 12, 13, xylan using an endo-&beta;-(1-4)-xylanase 6,7, or for pectic polysaccharides using a combination of a polygalacturonase and a methylesterase 8. Furthermore, using the same principles of OLIMP glycosylhydrolase and even glycosyltransferase activities can be monitored and determined 9.

OLIgo Mass Profiling OLIMP of Extracellular Polysaccharides

A rapid way is described to gain insights into the structure of polysaccharides in an extracellular matrix. The method takes advantage of the specificity of glycosylhydrolases and the sensitivity of mass spectrometry allowing minute amounts of materials to be analyzed. This technique is adaptable to be used directly on tissue itself.

The direct contact of cells to the environment is mediated in many organisms by an extracellular matrix. One common aspect of extracellular matrices is ...

Glycan Profiling of Plant Cell Wall Polymers using Microarrays 

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the raw materials for human societies (e.g. wood, paper, textile and biofuel industries)1,2. However, understanding the biosynthesis and function of these components remains challenging.
Cell wall glycans are chemically and conformationally diverse due to the complexity of their building blocks, the glycosyl residues. These form linkages at multiple positions and differ in ring structure, isomeric or anomeric configuration, and in addition, are substituted with an array of non-sugar residues. Glycan composition varies in different cell and/or tissue types or even sub-domains of a single cell wall3. Furthermore, their composition is also modified during development1, or in response to environmental cues4.
In excess of 2,000 genes have Plant cell walls are complex matrixes of heterogeneous glycans been predicted to be involved in cell wall glycan biosynthesis and modification in Arabidopsis5. However, relatively few of the biosynthetic genes have been functionally characterized 4,5. Reverse genetics approaches are difficult because the genes are often differentially expressed, often at low levels, between cell types6. Also, mutant studies are often hindered by gene redundancy or compensatory mechanisms to ensure appropriate cell wall function is maintained7. Thus novel approaches are needed to rapidly characterise the diverse range of glycan structures and to facilitate functional genomics approaches to understanding cell wall biosynthesis and modification.
Monoclonal antibodies (mAbs)8,9 have emerged as an important tool for determining glycan structure and distribution in plants. These recognise distinct epitopes present within major classes of plant cell wall glycans, including pectins, xyloglucans, xylans, mannans, glucans and arabinogalactans. Recently their use has been extended to large-scale screening experiments to determine the relative abundance of glycans in a broad range of plant and tissue types simultaneously9,10,11.
Here we present a microarray-based glycan screening method called Comprehensive Microarray Polymer Profiling (CoMPP) (Figures 1 &amp; 2)10,11 that enables multiple samples (100 sec) to be screened using a miniaturised microarray platform with reduced reagent and sample volumes. The spot signals on the microarray can be formally quantified to give semi-quantitative data about glycan epitope occurrence. This approach is well suited to tracking glycan changes in complex biological systems12 and providing a global overview of cell wall composition particularly when prior knowledge of this is unavailable.

Glycan Profiling of Plant Cell Wall Polymers using Microarrays

A technique called Comprehensive Microarray Polymer Profiling (CoMPP) for the characterisation of plant cell wall glycans is described. This method combines the specificity of monoclonal antibodies directed to defined glycan-epitopes with a miniature microarray analytical platform allowing screening of glycan occurrence in a broad range of biological contexts.

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the ...

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix containing polysaccharides that include cellulose microfibrils that form both crystalline structures and cellulose chains of amorphous organization. The orientation of the cellulose fibers and their concentrations dictate the mechanical properties of the cell. Several methods are used to determine the levels of crystalline cellulose, each bringing both advantages and limitations. Some can distinguish the proportion of crystalline regions within the total cellulose. However, they are limited to whole-organ analyses that are deficient in spatiotemporal information. Others relying on live imaging, are limited by the use of imprecise dyes. Here, we report a sensitive polarized light-based system for specific quantification of relative light retardance, representing crystalline cellulose accumulation in cross sections of Arabidopsis thaliana roots. In this method, the cellular resolution and anatomical data are maintained, enabling direct comparisons between the different tissues composing the growing root. This approach opens a new analytical dimension, shedding light on the link between cell wall composition, cellular behavior and whole-organ growth.

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Crystalline cellulose is an important constituent of the plant cell wall. However, its quantification at a cellular resolution is technically challenging. Here, we report the use of polarized light technology and root cross sections to obtain information of cell wall composition at a spatiotemporal resolution.

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix ...

Elucidating Glycan Structure and Occurrence Using Microarray Polymer Profiling

Microarray Polymer Profiling (MAPP) for High-Throughput Glycan Analysis

Microarray polymer profiling (MAPP) is a robust and reproducible approach to systematically determine the composition and relative abundance of glycans and glycoconjugates within a variety of biological samples, including plant and algal tissues, food materials, and human, animal, and microbial samples. Microarray technology underpins the efficacy of this method by providing a miniaturized, high-throughput screening platform, allowing thousands of interactions between glycans and highly specific glycan-directed molecular probes to be characterized concomitantly, using only small amounts of analytes. Constituent glycans are chemically and enzymatically fractionated, before being sequentially extracted from the sample and directly immobilized onto nitrocellulose membranes. The glycan composition is determined by the attachment of specific glycan-recognizing molecular probes to the extorted and printed molecules. MAPP is complementary to conventional glycan analysis techniques, such as monosaccharide and linkage analysis and mass spectrometry. However, glycan-recognizing molecular probes provide insight into the structural configurations of glycans, which can aid in elucidating biological interactions and functional roles.

Author Spotlight: MAPP Protocol &#8211; Advancing Glycan Analysis 

Microarray polymer profiling (MAPP) is a high-throughput technique for compositional analysis of glycans in biological samples.

Microarray polymer profiling (MAPP) is a robust and reproducible approach to systematically determine the composition and relative abundance of glycans ...

Structural Characterization of Mannan Cell Wall Polysaccharides in Plants Using PACE

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