This work was funded by F31 HL143873-01 (WBL), R01 HL125371 (RJL and EPS)

All biological samples analyzed in this protocol were obtained from mice, under protocols approved by the University of Colorado Institutional Animal Care and Use Committee.
1. Heparan sulfate isolation
<ol>
	<li>Delipidation of tissue samples 
	NOTE: Delipidation is an optional step for fat-rich tissues.
	<ol>
		<li>Make a 1:1 mixture of methanol and dichloromethane. Prepare approximately 500 &#956;L per sample; larger pieces of tissue may require up to 1 mL.</li>
		<li>Place each tissue sample into a small glass container with a lid for delipidation. 
		NOTE: Sample mass may vary per tissue of interest and experimental needs. 50 mg or less is typically sufficient for adequate GAG yield, but some optimization may be required by the end-user.&#160;</li>
		<li>Add 500 &#956;L of the methanol:dichloromethane solution to each glass container and mix. Ensure all solid tissue samples are completely submerged in solvent. 
		NOTE: Use serological pipettes or plastic conical tubes to mix and handle the methanol/dichloromethane solution; other plastics may dissolve.</li>
		<li>Place samples on a shaker (in secured rack) in a chemical fume hood and agitate gently for 1 h.</li>
		<li>&#160;Agitate the samples to mix and centrifuge at 17,000 x g for 5 min at 4 &#176;C.</li>
		<li>Carefully pipette out the organic solvent fraction (supernatant). This is the lipid fraction - it does not contain GAGs and can be discarded. 
		NOTE: Try not to disturb the tissue as small pieces could be lost. It is preferable to leave some of the organic layer in the sample container, rather than risking the sample loss.</li>
		<li>Leave the lid of the glass container open and let it evaporate overnight in the hood. Change the tube for the next step as those&#160;tubes will not survive further processing.</li>
		<li>Once the solvent mixture has fully evaporated, proceed to mechanical disintegration (if residual sample is &#62;50 mg) or Actinase E digestion (&#60; 50 mg).&#160;</li>
	</ol>
	</li>
	<li>Mechanical disintegration of solid tissue (optional; for larger tissue samples) 
	NOTE: Most smaller pieces of solid tissue (approximately 50 mg or less) should dissolve completely during the digestion step. However, larger samples will require mechanical disintegration.
	<ol>
		<li>Flash-freeze samples of interest by placing them in an appropriately sized polypropylene tube and placing the closed tube into liquid nitrogen. Allow the sample to freeze until completely solid.</li>
		<li>Using a clean mortar and pestle, grind frozen samples into a powder-like consistency.</li>
		<li>Proceed directly to Actinase E digestion (Step 1.4).</li>
	</ol>
	</li>
	<li>Sample desalting and concentration 
	NOTE: This is an optional step and only required for dilute liquid tissue samples (e.g., broncho-alveolar lavage fluid (BALF) or plasma).
	<ol>
		<li>Pool liquid samples into appropriate experimental groups (e.g., by biological replicate, or experimental group)</li>
		<li>Place total sample volume into a 500 &#956;L centrifugal filter column with a molecular weight cut-off (MWCO) of 3,000 Da.</li>
		<li>Spin for 30 min at 14,000 x g at room temperature. Repeat as needed if the desired sample volume exceeds capacity of the centrifugal filter column.</li>
		<li>Wash each column 3x with 400 &#956;L of deionized, filtered water. Discard the flow through.</li>
		<li>Invert the filter and spin for 1 min at 2,000 x g in fresh, appropriately labeled collection tubes. Freeze at -80 &#176;C or proceed to next step.</li>
	</ol>
	</li>
	<li>Actinase E digestion 
	NOTE: This step is required to digest and ultimately remove protein contaminants from your sample.
	<ol>
		<li>Mix samples 1:1 with recombinant Actinase E to a desired concentration of 10 mg/mL. Add appropriate volume of 10x digestion buffer concentrate. Desired final concentration: 0.005 M calcium acetate and 0.01 M sodium acetate, pH 7.5. For example: 190 &#956;L of liquid sample, 190 &#956;L of 20 mg/mL Actinase E, 20 &#956;L of 0.05 M calcium acetate, 0.1 M sodium acetate.</li>
		<li>Agitate samples gently to mix, then digest for 48-72 h at 55 &#176;C (up to 7 days for whole tissue).</li>
		<li>Heat to 80 &#176;C for 15-20 min to heat inactivate Actinase E.&#160;</li>
		<li>Freeze the sample at -80 &#176;C or continue to step 1.5.</li>
	</ol>
	</li>
	<li>Sample desalting and concentration 
	NOTE:&#160;If a low mass of the GAG of interest is anticipated in the biological sample, or if conservation of consumable reagents (i.e., centrifugal filter columns) is desirable, samples can be pooled to produce a single concentrate per experimental&#160;group (e.g., by biological replicate, or experimental group).
	<ol>
		<li>Place total sample volume into a 500 &#956;L centrifugal filter column with a MWCO of 3,000 Da.</li>
		<li>Spin for 30 min at 14,000 x g at room temperature. Repeat as needed, if the desired sample volume exceeds capacity of the centrifugal filter column.</li>
		<li>Wash each column 3x with 400 &#956;L deionized, filtered water. Discard the flow through.</li>
		<li>Invert filter and spin for 1 min at 2,000 x g in fresh, appropriately labeled collection tubes (generally included with the centrifugal filter columns). Freeze at -80 &#176;C or proceed to next step.</li>
	</ol>
	</li>
	<li>Desiccation
	<ol>
		<li>Either place the dissolved samples from step 1.5.5 in a rotational vacuum concentrator overnight or lyophilize the samples as detailed below.</li>
		<li>Freeze samples thoroughly either overnight in -80 &#176;C or by dipping in liquid nitrogen.</li>
		<li>Pierce sample lids with an 18 G needle and place in the lyophilizer chamber. Add paper towels for packing as needed.</li>
		<li>Fix lyophilizer chamber to lyophilizer and freeze dry overnight (at least -40 &#176;C, 0.135 Torr)</li>
	</ol>
	</li>
	<li>Cation exchange column
	<ol>
		<li>Resuspend desiccated samples in up to 400 &#956;L of 8 M urea, 2% CHAPS solution (or, if pooling samples, resuspend to a max of (400/n) &#956;L, where n = the number of samples in desired pool. Use as little of the detergent solution as possible.</li>
		<li>Equilibrate the cation exchange (IEX) column with 400 &#956;L of 8 M urea, 2% CHAPS solution. Spin for 5 min at 2,000 x g at room temperature.</li>
		<li>Load 400 &#956;L of sample/pooled samples into the IEX column. Spin for 5 min at 2,000 x g.</li>
		<li>Wash 3x with 400 &#956;L of 8 M urea, 2% CHAPS solution. Spin for 5 min at 2,000 x g everytime.</li>
		<li>Elute 3x with 400 &#956;L of 0.2 M NaCl. Spin for 5 min at 2,000 x g each. This is the low-affinity fraction - this can be retained for quality control purposes if desired.</li>
		<li>Elute 3x with 400 &#956;L of 2.7 M (16%) NaCl. Spin for 5 min at 2,000 x g each. This fraction will contain the isolated glycosaminoglycans of interest - keep all of it!</li>
		<li>To desalt each eluted fraction, add methanol up to 80 vol% and incubate at 4 &#176;C overnight. Spin each sample for 5 min at 2,000 x g. Recover the solid residue as this is dried de-salted glycosaminoglycan. 
		NOTE: Alternatively, skip this step and proceed to step 1.8 to de-salt the eluate without methanol.</li>
	</ol>
	</li>
	<li>Sample desalting and concentration
	<ol>
		<li>If necessary, pool eluted fractions (from 1.7.6) into appropriate experimental groups (e.g., by biological replicate, or experimental group).</li>
		<li>Place total sample volume into a 500 &#956;L centrifugal filter column with a MWCO of 3,000 Da.</li>
		<li>Spin for 30 min at 14,000 x g at room temperature. Repeat as needed, if the desired sample volume exceeds capacity of the centrifugal filter column.</li>
		<li>Wash each column 3x with 400 &#956;L deionized, filtered water. Discard flow through. Proceed directly to step 1.9.</li>
	</ol>
	</li>
	<li>Chondroitin digestion 
	NOTE: The purpose of this step is to remove GAGs not of interest to the end user. In this case, chondroitinase is used to remove chondroitin. In the tissues used for generating Representative Results (broncho-alveolar lavage fluid and whole lung), digesting chondroitin sulfate leaves heparan sulfate as the primary residual GAG. End users may need to add additional digestion steps depending on their experimental aims.&#160;
	<ol>
		<li>Load 350 &#181;L of digestion buffer (50 mM ammonium acetate with 2 mM calcium chloride adjusted to pH 7.0) to the centrifugal filter column without touching the membrane.</li>
		<li>Add 5 &#181;L of recombinant chondroitinase ABC.</li>
		<li>Place the samples tube into a 37 &#176;C oven and incubate for 1 h.</li>
		<li>Turn the column over and place into appropriately labeled collection tubes. Spin for 1 min at 2000 x g.</li>
		<li>Heat samples to 80 &#176;C for 15-20 min to inactivate chondroitinase ABC.</li>
	</ol>
	</li>
	<li>Sample desalting and concentration
	<ol>
		<li>If necessary, pool chondroitin digested samples from step 1.9.5 into appropriate experimental groups (e.g., by biological replicate, or experimental group)</li>
		<li>Place the total sample volume into a 500 &#956;L centrifugal filter column with a MWCO of 3,000 Da.</li>
		<li>Spin for 30 min at 14,000 x g at room temperature. Repeat as needed if desired sample volume exceeds capacity of the centrifugal filter column.</li>
		<li>Wash each column 3x with 400 &#956;L deionized, filtered water. Discard the flow through.</li>
		<li>Invert filter and spin for 1 min at 2,000 x g in fresh, appropriately labeled collection tubes. Freeze at -80 &#176;C or proceed to next step.</li>
	</ol>
	</li>
	<li>Desiccation 
	NOTE: In this step, desiccation is necessary so that samples can be resuspended in the smallest possible&#160;quantity of water and running buffer.
	<ol>
		<li>Either place the chondroitin digested samples from step 1.10.5 directly in a rotational vacuum concentrator overnight or lyophilize as follows:</li>
		<li>Freeze samples thoroughly either overnight in -80 &#176;C or by dipping in liquid nitrogen.</li>
		<li>Pierce sample lids with an 18 G needle and place in the lyophilizer chamber. Add paper towels for packing as needed.</li>
		<li>Fix lyophilizer chamber to lyophilizer and freeze dry overnight (at least 40 &#176;C, 0.135 Torr)</li>
	</ol>
	</li>
</ol>
2. Polyacrylamide gel electrophoresis of isolated and purified glycosaminoglycans
<ol>
	<li>Prepare solutions necessary for polyacrylamide gel electrophoresis (PAGE) in advance (Table 1). 
	NOTE: Select the percent acrylamide of resolving gel solution depending on the size of the glycosaminoglycans expected to be in the sample. 15% is recommended for resolving larger fragments (greater than 30 disaccharide subunits in length); 22% for smaller fragments (&#60;20 disaccharide subunits in length).</li>
	<li>Place empty cassette into the PAGE tank. Cast the resolving gel as follows: In 15 mL tube, mix 10 mL of resolving gel solution, 60 &#956;L of 10% ammonium persulfate (must be freshly prepared), and 10 &#956;L of TEMED (add TEMED last). Invert the tube gently 2-3x. Use pipette to quickly add the above 10 mL solution to cassette. Overlay with 2 mL of deionized, filtered water and allow the resolving gel to polymerize for 30 min. 
	NOTE: The following PAGE protocol has been optimized for a vertical PAGE system using 13.3 x 8.7cm (width x length) 1.0mm thick casting cassettes with a total volume of approximately 12mL. Other cassette systems can be used but may require optimization by the end-user.</li>
	<li>After the resolving gel has fully polymerized, discard the overlaid water and cast the stacking gel as follows: in a 15 mL tube, mix 3 mL of the stacking gel solution, 90 &#956;L of 10% ammonium persulfate (must be freshly prepared), 3 &#956;L of TEMED (add TEMED last).</li>
	<li>Invert the tube gently 2-3x. Use a pipette to quickly add the stacking gel solution over the solidified resolving gel; fill cassette to brim. Fully insert comb included with the set up. Allow the stacking gel to polymerize for 30 min.</li>
	<li>Once the gel has polymerized, ensure the tape strip is removed from the bottom of the cassette, and place the cassette back into the PAGE tank assembly.</li>
	<li>Fill the upper and lower chambers with upper and lower chamber buffer, respectively.</li>
	<li>Dissolve the dried samples from step 1.11.4 in the minimum necessary volume of deionized, filtered water (at most, 50% of the volume of the wells in the PAGE gel). Mix 1:1 with sample loading buffer. Load the samples and the HS oligosaccharide &#34;ladders&#34; (see Table 1) into the gel.</li>
	<li>Pre-run the gel for 5 min at 100 V. Then run the gel at 200 V for 20-25 min (for a 15% polyacrylamide resolving gel), 40-50 min (for an 18% polyacrylamide resolving gel), 90-100 min (for 22% polyacrylamide resolving gel). 
	NOTE: Some optimization of the 200 V run time may be necessary. Phenol red migrates ahead of heparin oligosaccharides that are 2 polymer subunits in length (i.e., degree of polymerization 2, or dp2); bromophenol blue migrates ahead of dp10-dp14. Best results are obtained when the voltage is applied such that the phenol red band migrates almost, but not quite, to the bottom of the gel. Adjust run time accordingly.</li>
</ol>
3. Silver staining protocol
<ol>
	<li>Prepare all solutions necessary for silver staining in advance (Table 2). 
	NOTE: Do not directly touch the PAGE gel until it has been stained, developed, and placed in stop solution. Instead, manipulate the gel using clean plastic or glass tools. Directly handling the gel will result in finger-print distortions and other visible artifacts on the gel after staining. &#160;</li>
	<li>Once the run is completed, disassemble cassette and extract gel in a clean, medium-large container filled with deionized, filtered water. 
	NOTE: To avoid directly handling the gel, use pipette tip or other plastic object to gently peel the gel away from the cassette while submerged in water. Gel may be fragile - handle carefully.
	<ol>
		<li>Discard the water. Stain the gel in Alcian blue staining solution for 5 min.</li>
		<li>Discard Alcian blue stain. Quickly rinse/wash 2-3x with deionized, filtered water until most of the Alcian blue staining solution has been removed.</li>
		<li>Allow to de-stain in deionized, filtered water overnight on rocker. Ensure that there is ample volume of deionized, filtered water to ensure any residual stain is fully washed off the gel overnight.</li>
		<li>Wash gel in 50% methanol (40 min total, change solution 2-3x).</li>
		<li>Wash gel in deionized, filtered water for 30 min. Discard water and repeat 3 more time for a total of 2 h, replacing the water each time.</li>
		<li>In a fresh, clean container, stain the gel for 30 min in silver nitrate staining solution.</li>
		<li>Quickly rinse/wash 2-3x in deionized, filtered water to fully remove the silver staining solution.</li>
		<li>Wash for 30 min in deionized, filtered water. Discard water and repeat 2x for a total of 90 min, replacing the water bath each time.</li>
		<li>Discard water and add developing solution.</li>
		<li>Once developing solution is added, carefully observe the gel and watch for the appearance of bands. Depending on the quality of the stain and the mass of the sample loaded, development can take anywhere from a few seconds to several minutes.</li>
		<li>As soon as the desired bands are visible, immediately discard developing solution and wash briefly with stop solution.</li>
		<li>Discard the stop solution wash and replace with fresh stop solution. Allow to soak for 1 h on a rocker or shaker.</li>
		<li>Wash in deionized, filtered water overnight (however, the gel can be imaged immediately after stop solution wash).</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

GAGs play a central role in many diverse biological processes. One of the principal functions of sulfated GAGs (such as HS and CS) is to interact with and bind to ligands, which can alter downstream signaling functions. An important determinant of GAG binding affinity to cognate ligands is the length of the GAG polymer chain 8,9,14. For this reason, it is important for researchers to be able to define with reasonable precision t...

Glycosaminoglycans (GAGs) are&#160;a diverse family of linear polysaccharides that are a ubiquitous element in living organisms and contribute to many basic physiological processes1. GAGs such as heparan sulfate (HS) and chondroitin sulfate (CS) may be sulfated at distinct positions along the polysaccharide chain, imparting geographic domains of negative charge. These GAGs, when tethered to cell-surface proteins known as proteoglycans, project into the extracellular space and bind to cognate ligands, allowing for the regulation of both cis- (ligand attached to the same cell) and trans- (ligand attached to neighboring cell) signaling processes2. Furthermore, GAGs also perform critical roles as structural elements in tissues such as the glomerular basement membrane3, the vascular endothelial glycocalyx4 and pulmonary epithelial glycocalyx5, and in connective tissues such cartilage6.
The length of GAG polysaccharide chains varies substantially according to its biological context and can be dynamically lengthened, cleaved, and modified by a highly complex enzymatic regulatory system7. Importantly, the length of GAG polymer chains contributes substantially to their binding affinity for ligands and, subsequently, to their biological function8,9. For this reason, determination of the function of an endogenous GAG requires appreciation of its size. Unfortunately, unlike proteins and nucleic acids, very few readily available techniques exist to detect and measure GAGs, which has historically resulted in relatively limited investigation into the biological roles of this diverse polysaccharide family.
This article describes how to isolate and purify GAGs from most biological tissues, and, also, describes how to use polyacrylamide gel electrophoresis (PAGE) to evaluate the length of the isolated polymers with a fair degree of specificity. In contrast to other, highly complex (and often mass spectrometry-based) glycomic approaches, this method can be employed using standard laboratory equipment and techniques. This practical approach may, therefore, expand investigators&#39; ability to determine the biological role of native GAGs and their interaction with contextually important ligands.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Accuspin Micro17 benchtop microcentrifuge</td>
			<td>thermoFisher Scientific</td>
			<td>13-100-675</td>
			<td>Any benchtop microcentrifuge/rotor combination capable of 14000 xG is appropriate</td>
		</tr>
		<tr>
			<td>Acrylamide (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>BP170-100</td>
			<td>Electrophoresis grade</td>
		</tr>
		<tr>
			<td>Actinase E</td>
			<td>Sigma Aldrich</td>
			<td>P5147</td>
			<td>Protease mix from S. griseus</td>
		</tr>
		<tr>
			<td>Alcian Blue 8GX (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>AC400460100</td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium acetate (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>A639-500</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>Ammonium hydroxide (liquid)</td>
			<td>thermoFisher Scientific</td>
			<td>A669S-500</td>
			<td>certified ACS</td>
		</tr>
		<tr>
			<td>Ammonium persulfate (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>BP179-25</td>
			<td>electrophoresis grade</td>
		</tr>
		<tr>
			<td>Barnstead GenPure Pro Water Purification System</td>
			<td>ThermoFisher Scientific</td>
			<td>10-451-217PKG</td>
			<td>Any water deionizing/ purification system is an acceptable substitute</td>
		</tr>
		<tr>
			<td>Boric acid (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>A73-500</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>Bromphenol blue (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>B392-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium acetate (solid)</td>
			<td>ThermoFisher Scientific</td>
			<td>18-609-432</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>Calcium chloride (solid)</td>
			<td>ThermoFisher Scientific</td>
			<td>AC349610250</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>CHAPS detergent (3-((3-cholamidopropyl) dimethylammonio)-1-propanesulfonate)</td>
			<td>ThermoFisher Scientific</td>
			<td>28299</td>
			<td></td>
		</tr>
		<tr>
			<td>Chondroitinase ABC</td>
			<td>Sigma Aldrich</td>
			<td>C3667</td>
			<td></td>
		</tr>
		<tr>
			<td>Criterion empty cassette for PAGE (1.0mm thick, 12+2 wells)</td>
			<td>Bio-Rad</td>
			<td>3459901</td>
			<td>Any 1.0mm thick PAGE casting cassette system will suffice</td>
		</tr>
		<tr>
			<td>Criterion PAGE Cell system (cell and power supply)</td>
			<td>Bio-Rad</td>
			<td>1656019</td>
			<td>any comparable vertical gel PAGE system will work)</td>
		</tr>
		<tr>
			<td>Dichloromethane (liquid)</td>
			<td>thermoFisher Scientific</td>
			<td>AC610931000</td>
			<td>certified ACS</td>
		</tr>
		<tr>
			<td>EDTA disodium salt (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>02-002-786</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>Glacial acetic acid (liquid)</td>
			<td>thermoFisher Scientific</td>
			<td>A35-500</td>
			<td>Certified ACS</td>
		</tr>
		<tr>
			<td>Glycine (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>G48-500</td>
			<td>Electrophoresis grade</td>
		</tr>
		<tr>
			<td>Heparanase I/III</td>
			<td>Sigma Aldrich</td>
			<td>H3917</td>
			<td>From Flavobacterium heparinum</td>
		</tr>
		<tr>
			<td>Heparin derived decasaccharide (dp10)</td>
			<td>galen scientific</td>
			<td>HO10</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin derived hexasaccharde (dp6)</td>
			<td>Galen scientific</td>
			<td>HO06</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin derived oligosaccharide (dp20)</td>
			<td>galen scientific</td>
			<td>HO20</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloric acid (liquid)</td>
			<td>thermoFisher Scientific</td>
			<td>A466-250</td>
			<td></td>
		</tr>
		<tr>
			<td>Lyophilizer</td>
			<td>Labconco</td>
			<td>7752020</td>
			<td>Any lyophilizer that can achieve -40C and 0.135 Torr will work; can also be replaced with rotational vacuum concentrator</td>
		</tr>
		<tr>
			<td>Methanol (liquid)</td>
			<td>thermoFisher Scientific</td>
			<td>A412-500</td>
			<td>Certified ACS</td>
		</tr>
		<tr>
			<td>Molecular Imager Gel Doc XR System</td>
			<td>Bio-Rad</td>
			<td>170-8170</td>
			<td>Any comparable gel imaging system is an acceptable substitute</td>
		</tr>
		<tr>
			<td>N,N&#39;-methylene-bis-acrylamide (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>BP171-25</td>
			<td>Electrophoresis grade</td>
		</tr>
		<tr>
			<td>Phenol red (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>P74-10</td>
			<td>Free acid</td>
		</tr>
		<tr>
			<td>Q Mini H Ion Exchange Column</td>
			<td>Vivapure</td>
			<td>VS-IX01QH24</td>
			<td>Ion exchange column must have minimum loading volume of 0.4mL, working pH of 2-12, and selectivity for ionic groups with pKa of 11</td>
		</tr>
		<tr>
			<td>Silver nitrate (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>S181-25</td>
			<td>certified ACS</td>
		</tr>
		<tr>
			<td>Sodium Acetate (solid)</td>
			<td>ThermoFisher Scientific</td>
			<td>S210-500</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>Sodium chloride (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>S271-500</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>Sodium hydroxide (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>S392-212</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>BP220-1</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>TEMED (N,N,N&#39;,N&#39;-tetramethylenediamine)</td>
			<td>thermoFisher Scientific</td>
			<td>BP150-20</td>
			<td>Electrophoresis grade</td>
		</tr>
		<tr>
			<td>Tris base (solid)</td>
			<td>thermoFisher Scientific</td>
			<td>BP152-500</td>
			<td>Molecular biology grade</td>
		</tr>
		<tr>
			<td>Ultra Centrifugal filters, 0.5mL, 3000 Da molecular weight cutoff</td>
			<td>Amicon</td>
			<td>UFC500324</td>
			<td>Larger volume filter units may be used, depending on sample size.&#160;</td>
		</tr>
		<tr>
			<td>Urea (solid)</td>
			<td>ThermoFisher Scientific</td>
			<td>29700</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacufuge Plus</td>
			<td>Eppendorf</td>
			<td>22820001</td>
			<td>Any rotational vacuum concentrator will work; can be replaced with lyophilizer</td>
		</tr>
		<tr>
			<td>Vacuum filter unit, single use, 0.22uM pore PES, 500mL volume</td>
			<td>thermoFisher Scientific</td>
			<td>569-0020</td>
			<td>Alternative volumes and filter materials acceptable</td>
		</tr>
	</tbody>
</table>

detection of glycosaminoglycans by polyacrylamide gel electrophoresis and silver staining

Sulfated glycosaminoglycans (GAGs) such as heparan sulfate (HS) and chondroitin sulfate (CS) are ubiquitous in living organisms and play a critical role in a variety of basic biological structures and processes. As polymers, GAGs exist as a polydisperse mixture containing polysaccharide chains that can range from 4000 Da to well over 40,000 Da. Within these chains exists domains of sulfation, conferring a pattern of negative charge that facilitates interaction with positively charged residues of cognate protein ligands. Sulfated domains of GAGs must be of sufficient length to allow for these electrostatic interactions. To understand the function of GAGs in biological tissues, the investigator must be able to isolate, purify, and measure the size of GAGs. This report describes a practical and versatile polyacrylamide gel electrophoresis-based technique that can be leveraged to resolve relatively small differences in size between GAGs isolated from a variety of biological tissue types.

Alcian blue is used to stain sulfated GAGs 10; this signal is amplified by use of a subsequent silver stain 11. Figure 1 provides a visual demonstration of the silver staining development process. As demonstrated, the Alcian blue signal representing GAGs separated by electrophoresis is amplified as the developing agent penetrates the polyacrylamide gel. Typically, the developing process will reduce silver and Alcian blue-stained GAGs in a densi...

Watch this Scientific Journal Video about Detection of Glycosaminoglycans by Polyacrylamide Gel Electrophoresis and Silver Staining at JoVE.com

Detection of Glycosaminoglycans by Polyacrylamide Gel Electrophoresis and Silver Staining

This report describes techniques to isolate and purify sulfated glycosaminoglycans (GAGs) from biological samples and a polyacrylamide gel electrophoresis approach to approximate their size. GAGs contribute to tissue structure and influence signaling processes via electrostatic interaction with proteins. GAG polymer length contributes to their binding affinity for cognate ligands.

Sulfated glycosaminoglycans (GAGs) such as heparan sulfate (HS) and chondroitin sulfate (CS) are ubiquitous in living organisms and play a critical role ...

detection-glycosaminoglycans-polyacrylamide-gel-electrophoresis

Research

JoVE Journal

Biology

4.5K Views.  University of Colorado Anschutz Medical Campus. This report describes techniques to isolate and purify sulfated glycosaminoglycans (GAGs) from biological samples and a polyacrylamide gel electrophoresis approach to approximate their size. GAGs contribute to tissue structure and influence signaling processes via electrostatic interaction with proteins. GAG polymer length contributes to their binding affinity for cognate ligands.

Video: Detection of Glycosaminoglycans by Polyacrylamide Gel Electrophoresis and Silver Staining

Denaturing Gradient Gel Electrophoresis (DGGE)

Denaturing Gradient Gel Electrophoresis DGGE

Okay.Hi, my name is Celeste Peterson and I'm a fellow at Harvard University studying microbial bio film communities. So I am profiling the bacterial communities by looking at their 60 N sequence and how this sequence changes over time in space. And to do that, I'm using DDGE.This is not a normal acry of my well, it needs to be kept at 60 degrees when it's run and the buffer needs to be pumped continuously. So we're gonna warm up the buffer for about an hour until it's at 60 degrees, and then we can run the towel. Now I'm gonna wash this plate with dilute uc solution, peeling around for impurities, andone, light that up.Anything like that can cause regularities. It is important to wipe off all the water. So when I assemble the gel, I'll put that chip on the outside of the gel.So the gel will be in between these two pieces of glass and the chip will be on the outside and it won't affect the gel in between. So now I'm gonna assemble the gel in these clamps and then I squeeze them together and tighten by turning to the right. As with any gel, it's important to make sure that the bottom is flush.To prevent making, we now turn the gel around and put it in here to get sealed with the gasket. So it's important when you do this that the gel, that the plates are actually lined up with the gasket here. So these knobs on the side then go in and you press down to type the seal.So this is a 30 mil syringe. And this part and this part come with the bio red kit and it just on like this. And that's just to get rid of the excess water in the tubes.So put the tubing onto the gel to load the gel. First you put an adapter into the tubing and then a needle look with that small adapter and put it right in the center of the gel just so it's fairly below the bottom glass. Then this wire adapter goes into the long tubing and it's going to connect to equal syringes containing the high and the low density solution.So these are pre-prepared solutions. The blue one, I've added a dye here that just indicates that this is the most denaturing solution and this is the least fitting train solution, the low density one. And they have everything in them.And now I'm gonna to make, gonna be Actually I twice as much the week. And once you've got the tempt, you have about five to seven minutes to fill the gel before the colon rises. And now we fill the syringe juice with the solution.So I'm gonna fill it with about 15 the solution. And then to get rid of any air bubbles, I'll turn it upside down and squeeze gently until that top air bubble is gone. Now this little screw here is gonna go into this groove.This adapter is gonna line up with this clamp here, and it's gonna get C clamped in very tightly. And then this tubing is gonna attach to the syringe. And the same thing is done on the other side with the high density matrix where it's time to deliver.And so I'm gonna be turning this wheel in a steady clockwise motion. And first the blue solution is gonna go out first because that's the the most in nature, and it's gonna form the bottom part of the gel. And then after a little while, the clear solution will start filling in the gel and the Gradient will be formed.One moment is the gently on an angle to And as soon as you finish pouring the gel to wash out your tubes immediately because they will polymerize Going, It's the group there. And then press down. So this little, And on the other side, I'll do the same thing.Okay.Hi, my name is Celeste Peterson and I'm a fellow at Harvard University studying microbial bio communities. So I am profiling the bacterial communities by looking at their 60 F sequence and how the sequence changes over time in space. And to do that, I'm using DDGE.So DGE is Dena gel electrophoresis. And basically the idea is that there's a gradient and the DNA, as it runs down the gradient gets de natured. And so different 16 species will stop at different points along the gradient.And so it ends up being a fingerprint of the different sexiness species in the community. So the goal of my experiment is to see who is in the community and how that's changing over time and space. And I can run the sexiness DDD gel and then I can cut out the bands and identify the species that are represented in each community.So one of the biggest issues when, when doing A DDG gel is to figure out the gradient that's the best. And so that is just something that needs to be played around with and figuring out for a particular community which type of gradient. So the one I'm using today is 35 to 55%urea.One of the big problems that can happen with this technique is that smears come up in in the leans. And I found for that, for whatever reason, one of the best things to do is to run the gels for a really long time and then the smears disappear. Well, today I'm just using it for general bacterial communities, but this can also be used for a specific family of bacteria to see how those change over time, and those tend to be very clean gels.You can also look at mutations in single genes with DDGE.

Screening for Amyloid Aggregation by Semi-Denaturing Detergent-Agarose Gel Electrophoresis

Amyloid aggregation is associated with numerous protein misfolding pathologies and underlies the infectious properties of prions, which are conformationally self-templating proteins that are thought to have beneficial roles in lower organisms.  Amyloids have been notoriously difficult to study due to their insolubility and structural heterogeneity.  However, resolution of amyloid polymers based on size and detergent insolubility has been made possible by Semi-Denaturing Detergent-Agarose Gel Electrophoresis (SDD-AGE).  This technique is finding widespread use for the detection and characterization of amyloid conformational variants.  Here, we demonstrate an adaptation of this technique that facilitates its use in large-scale applications, such as screens for novel prions and other amyloidogenic proteins.  The new SDD-AGE method uses capillary transfer for greater reliability and ease of use, and allows any sized gel to be accomodated.  Thus, a large number of samples, prepared from cells or purified proteins, can be processed simultaneously for the presence of SDS-insoluble conformers of tagged proteins.

SDD-AGE is a useful technique for the detection and characterization of amyloid-like polymers in cells. Here we demonstrate an adaptation that makes this technique amenable to large-scale applications.

Amyloid aggregation is associated with numerous protein misfolding pathologies and underlies the infectious properties of prions, which are ...

Fast and Sensitive Colloidal Coomassie G-250 Staining for Proteins in Polyacrylamide Gels

Coomassie Brilliant Blue (CBB) is a dye commonly used for the visualization of proteins separated by SDS-PAGE, offering a simple staining procedure and high quantitation. Furthermore, it is completely compatible with mass spectrometric protein identification. But despite these advantages, CBB is regarded to be less sensitive than silver or fluorescence stainings and therefore rarely used for the detection of proteins in analytical gel-based proteomic approaches.
Several improvements of the original Coomassie protocol1 have been made to increase the sensitivity of CBB. Two major modifications were introduced to enhance the detection of low-abundant proteins by converting the dye molecules into colloidal particles: In 1988, Neuhoff and colleagues applied 20% methanol and higher concentrations of ammonium sulfate into the CBB G-250 based staining solution2, and in 2004 Candiano et al. established Blue Silver using CBB G-250 with phosphoric acid in the presence of ammonium sulfate and methanol3. Nevertheless, all these modifications just allow a detection of approximately 10 ng protein. A widely fameless protocol for colloidal Coomassie staining was published by Kang et al. in 2002 where they modified Neuhoff's colloidal CBB staining protocol regarding the complexing substances. Instead of ammonium sulfate they used aluminum sulfate and methanol was replaced by the less toxic ethanol4. The novel aluminum-based staining in Kang's study showed superior sensitivity that detects as low as 1 ng/band (phosphorylase b) with little sensitivity variation depending on proteins.
Here, we demonstrate application of Kang's protocol for fast and sensitive colloidal Coomassie staining of proteins in analytical purposes. We will illustrate the quick and easy protocol using two-dimensional gels routinely performed in our working group.

This video shall popularize a colloidal Coomassie G-250 staining protocol according to Kang et al. for the detection of average 4 ng protein in gels. The staining is completed within 2 hours and without any effort. We routinely use Kang's protocol for analytical purposes in gel-based proteomics.

Coomassie Brilliant Blue (CBB) is a dye commonly used for the visualization of proteins separated by SDS-PAGE, offering a simple staining procedure and ...

Denaturing Urea Polyacrylamide Gel Electrophoresis (Urea PAGE)

Urea PAGE or denaturing urea polyacrylamide gel electrophoresis employs 6-8 M urea, which denatures secondary DNA or RNA structures and is used for their separation in a polyacrylamide gel matrix based on the molecular weight. Fragments between 2 to 500 bases, with length differences as small as a single nucleotide, can be separated using this method1. The migration of the sample is dependent on the chosen acrylamide concentration. A higher percentage of polyacrylamide resolves lower molecular weight fragments. The combination of urea and temperatures of 45-55 &deg;C during the gel run allows for the separation of unstructured DNA or RNA molecules.
In general this method is required to analyze or purify single stranded DNA or RNA fragments, such as synthesized or labeled oligonucleotides or products from enzymatic cleavage reactions.
In this video article we show how to prepare and run the denaturing urea polyacrylamide gels. Technical tips are included, in addition to the original protocol 1,2.

Denaturing Urea Polyacrylamide Gel Electrophoresis Urea PAGE

Denaturing urea polyacrylamide gel electrophoresis is used to separate single-stranded DNA or RNA up to a limit of 500 nucleotides. Urea in combination with heat denatures samples and unstructured single strands migrate within the gel matrix according to their molecular weight.

Urea PAGE or denaturing urea polyacrylamide gel electrophoresis employs 6-8 M urea, which denatures secondary DNA or RNA structures and is used for their ...

Blue Native Polyacrylamide Gel Electrophoresis (BN-PAGE) for Analysis of Multiprotein Complexes from Cellular Lysates

Multiprotein complexes (MPCs) play a crucial role in cell signalling, since most proteins can be found in functional or regulatory complexes with other proteins (Sali, Glaeser et al. 2003). Thus, the study of protein-protein interaction networks requires the detailed characterization of MPCs to gain an integrative understanding of protein function and regulation. For identification and analysis, MPCs must be separated under native conditions. In this video, we describe the analysis of MPCs by blue native polyacrylamide gel electrophoresis (BN-PAGE). BN-PAGE is a technique that allows separation of MPCs in a native conformation with a higher resolution than offered by gel filtration or sucrose density ultracentrifugation, and is therefore useful to determine MPC size, composition, and relative abundance (Sch&auml;gger and von Jagow 1991); (Sch&auml;gger, Cramer et al. 1994). By this method, proteins are separated according to their hydrodynamic size and shape in a polyacrylamide matrix. Here, we demonstrate the analysis of MPCs of total cellular lysates, pointing out that lysate dialysis is the crucial step to make BN-PAGE applicable to these biological samples. Using a combination of first dimension BN- and second dimension SDS-PAGE, we show that MPCs separated by BN-PAGE can be further subdivided into their individual constituents by SDS-PAGE. Visualization of the MPC components upon gel separation is performed by standard immunoblotting. As an example for MPC analysis by BN-PAGE, we chose the well-characterized eukaryotic 19S, 20S, and 26S proteasomes.

Blue Native Polyacrylamide Gel Electrophoresis BN-PAGE for Analysis of Multiprotein Complexes from Cellular Lysates

In this video, we describe the characterization of multiprotein complexes (MPCs) by blue native polyacrylamide gel electrophoresis (BN-PAGE). In a first dimension, dialyzed cellular lysates are separated by BN-PAGE to identify individual MPCs. In a second dimension SDS-PAGE, MPCs of interest are further subdivided to analyze their constituents by immunoblotting.

Multiprotein complexes (MPCs) play a crucial role in cell signalling, since most proteins can be found in functional or regulatory complexes with other ...

Detection of Functional Matrix Metalloproteinases by Zymography

Matrix metalloproteinases (MMPs) are zinc-containing endopeptidases. They degrade proteins by cleavage of peptide bonds. More than twenty MMPs have been identified and are separated into six groups based on their structure and substrate specificity (collagenases, gelatinases, membrane type [MT-MMP], stromelysins, matrilysins, and others). MMPs play a critical role in cell invasion, cartilage degradation, tissue remodeling, wound healing, and embryogenesis. They therefore participate in both normal processes and in the pathogenesis of many diseases, such as rheumatoid arthritis, cancer, or chronic obstructive pulmonary disease1-6. Here, we will focus on MMP-2 (gelatinase A, type IV collagenase), a widely expressed MMP. We will demonstrate how to detect MMP-2 in cell culture supernatants by zymography, a commonly used, simple, and yet very sensitive technique first described in 1980 by C. Heussen and E.B. Dowdle7-10. This technique is semi-quantitative, it can therefore be used to determine MMP levels in test samples when known concentrations of recombinant MMP are loaded on the same gel11.
Solutions containing MMPs (e.g. cell culture supernatants, urine, or serum) are loaded onto a polyacrylamide gel containing sodium dodecyl sulfate (SDS; to linearize the proteins) and gelatin (substrate for MMP-2). The sample buffer is designed to increase sample viscosity (to facilitate gel loading), provide a tracking dye (bromophenol blue; to monitor sample migration), provide denaturing molecules (to linearize proteins), and control the pH of the sample. Proteins are then allowed to migrate under an electric current in a running buffer designed to provide a constant migration rate. The distance of migration is inversely correlated with the molecular weight of the protein (small proteins move faster through the gel than large proteins do and therefore migrate further down the gel). After migration, the gel is placed in a renaturing buffer to allow proteins to regain their tertiary structure, necessary for enzymatic activity. The gel is then placed in a developing buffer designed to allow the protease to digest its substrate. The developing buffer also contains p-aminophenylmercuric acetate (APMA) to activate the non-proteolytic pro-MMPs into active MMPs. The next step consists of staining the substrate (gelatin in our example). After washing the excess dye off the gel, areas of protease digestion appear as clear bands. The clearer the band, the more concentrated the protease it contains. Band staining intensity can then be determined by densitometry, using a software such as ImageJ, allowing for sample comparison.

This protocol describes an activity-based assay for detecting matrix metalloproteinases in culture supernatants or body fluids.

Matrix metalloproteinases (MMPs) are zinc-containing endopeptidases. They degrade proteins by cleavage of peptide bonds. More than twenty MMPs have been ...

Pouring and Running a Protein Gel by reusing Commercial Cassettes

The evaluation of proteins using sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) analysis is a common technique used by biochemistry and molecular biology researchers1-4. For laboratories that perform daily analyses of proteins, the cost of commercially available polyacrylamide gels (&tilde;$10/gel) can be considerable over time. To mitigate this cost, some researchers prepare their own polyacrylamide gels. Traditional methods of pouring these gels typically utilize specialized equipment and glass gel plates that can be expensive and preclude pouring many gels and storing them for future use. Furthermore, handling of glass plates during cleaning or gel pouring can result in accidental breakage creating a safety hazard, which may preclude their use in undergraduate laboratory classes. Our protocol demonstrates how to pour multiple protein gels simultaneously by recycling Invitrogen Nupage Novex minigel cassettes, and inexpensive materials purchased at a home improvement store. This economical and streamlined method includes a way to store the gels at 4&deg;C for a few weeks. By re-using the plastic gel cassettes from commercially available gels, labs that run frequent protein gels can save significant costs and help the environment. In addition, plastic gel cassettes are extremely resistant to breakage, which makes them ideal for undergraduate laboratory classrooms.

Our protocol demonstrates how to pour multiple protein gels at a time by recycling Invitrogen Nupage Novex minigel cassettes, and inexpensive materials purchased at a home improvement store. This economical and streamlined method includes a way to store the gels at 4&deg;C for a few weeks. By re-using the plastic gel cassettes from commercially available gels, labs that run frequent protein gels can save significant costs and help the environment.

The evaluation of proteins using sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) analysis is a common technique used by biochemistry ...

Agarose Gel Electrophoresis for the Separation of DNA Fragments

Agarose gel electrophoresis is the most effective way of separating DNA fragments of varying sizes ranging from 100 bp to 25 kb1. Agarose is isolated from the seaweed genera Gelidium and Gracilaria, and consists of repeated agarobiose (L- and D-galactose) subunits2. During gelation, agarose polymers associate non-covalently and form a network of bundles whose pore sizes determine a gel&#39;s molecular sieving properties. The use of agarose gel electrophoresis revolutionized the separation of DNA. Prior to the adoption of agarose gels, DNA was primarily separated using sucrose density gradient centrifugation, which only provided an approximation of size. To separate DNA using agarose gel electrophoresis, the DNA is loaded into pre-cast wells in the gel and a current applied. The phosphate backbone of the DNA (and RNA) molecule is negatively charged, therefore when placed in an electric field, DNA fragments will migrate to the positively charged anode. Because DNA has a uniform mass/charge ratio, DNA molecules are separated by size within an agarose gel in a pattern such that the distance traveled is inversely proportional to the log of its molecular weight3. The leading model for DNA movement through an agarose gel is &#34;biased reptation&#34;, whereby the leading edge moves forward and pulls the rest of the molecule along4. The rate of migration of a DNA molecule through a gel is determined by the following: 1) size of DNA molecule; 2) agarose concentration; 3) DNA conformation5; 4) voltage applied, 5) presence of ethidium bromide, 6) type of agarose and 7) electrophoresis buffer. After separation, the DNA molecules can be visualized under uv light after staining with an appropriate dye. By following this protocol, students should be able to: 
 
1. Understand the mechanism by which DNA fragments are separated within a gel matrix 
2. Understand how conformation of the DNA molecule will determine its mobility through a gel matrix 
3. Identify an agarose solution of appropriate concentration for their needs 
4. Prepare an agarose gel for electrophoresis of DNA samples 
5. Set up the gel electrophoresis apparatus and power supply 
6. Select an appropriate voltage for the separation of DNA fragments 
7. Understand the mechanism by which ethidium bromide allows for the visualization of DNA bands 
8. Determine the sizes of separated DNA fragments 
&#160;

A basic protocol for the separation of DNA fragments using agarose gel electrophoresis is described.

Agarose gel electrophoresis is the most effective way of separating DNA fragments of varying sizes ranging from 100 bp to 25 kb1. Agarose is isolated from ...

Detection of Viral RNA by Fluorescence in situ Hybridization (FISH)

Viruses that infect cells elicit specific changes to normal cell functions which serve to divert energy and resources for viral replication. Many aspects of host cell function are commandeered by viruses, usually by the expression of viral gene products that recruit host cell proteins and machineries. Moreover, viruses engineer specific membrane organelles or tag on to mobile vesicles and motor proteins to target regions of the cell (during de novo infection, viruses co-opt molecular motor proteins to target the nucleus; later, during virus assembly, they will hijack cellular machineries that will help in the assembly of viruses). Less is understood on how viruses, in particular those with RNA genomes, coordinate the intracellular trafficking of both protein and RNA components and how they achieve assembly of infectious particles at specific loci in the cell. The study of RNA localization began in earlier work. Developing lower eukaryotic embryos and neuronal cells provided important biological information, and also underscored the importance of RNA localization in the programming of gene expression cascades. The study in other organisms and cell systems has yielded similar important information. Viruses are obligate parasites and must utilise their host cells to replicate. Thus, it is critical to understand how RNA viruses direct their RNA genomes from the nucleus, through the nuclear pore, through the cytoplasm and on to one of its final destinations, into progeny virus particles 1.
FISH serves as a useful tool to identify changes in steady-state localization of viral RNA. When combined with immunofluorescence (IF) analysis 22, FISH/IF co-analyses will provide information on the co-localization of proteins with the viral RNA3. This analysis therefore provides a good starting point to test for RNA-protein interactions by other biochemical or biophysical tests 4,5, since co-localization by itself is not enough evidence to be certain of an interaction. In studying viral RNA localization using a method like this, abundant information has been gained on both viral and cellular RNA trafficking events 6. For instance, HIV-1 produces RNA in the nucleus of infected cells but the RNA is only translated in the cytoplasm. When one key viral protein is missing (Rev) 7, FISH of the viral RNA has revealed that the block to viral replication is due to the retention of the HIV-1 genomic RNA in the nucleus 8. 
Here, we present the method for visual analysis of viral genomic RNA in situ. The method makes use of a labelled RNA probe. This probe is designed to be complementary to the viral genomic RNA. During the in vitro synthesis of the antisense RNA probe, the ribonucleotide that is modified with digoxigenin (DIG) is included in an in vitro transcription reaction. Once the probe has hybridized to the target mRNA in cells, subsequent antibody labelling steps (Figure 1) will reveal the localization of the mRNA as well as proteins of interest when performing FISH/IF.

Detection of Viral RNA by Fluorescence in situ Hybridization FISH

A fluorescence in situ hybridization (FISH) method was developed to visually detect viral genomic RNA using fluorescence microscopy. A probe is made with specificity to the viral RNA that can then be identified using a combination of hybridization and immunofluorescence techniques. This technique offers the advantage of identifying the localization of the viral RNA or DNA at steady-state, providing information on the control of intracellular virus trafficking events.

Viruses  that infect cells elicit specific changes to normal cell functions which serve  to divert energy and resources for viral replication. Many ...

Preparation of DNA-crosslinked Polyacrylamide Hydrogels

Mechanobiology is an emerging scientific area that addresses the critical role of physical cues in directing cell morphology and function. For example, the effect of tissue elasticity on cell function is a major area of mechanobiology research because tissue stiffness modulates with disease, development, and injury. Static tissue-mimicking materials, or materials that cannot alter stiffness once cells are plated, are predominately used to investigate the effects of tissue stiffness on cell functions. While information gathered from static studies is valuable, these studies are not indicative of the dynamic nature of the cellular microenvironment in vivo. To better address the effects of dynamic stiffness on cell function, we developed a DNA-crosslinked polyacrylamide hydrogel system (DNA gels). Unlike other dynamic substrates, DNA gels have the ability to decrease or increase in stiffness after fabrication without stimuli. DNA gels consist of DNA crosslinks that are polymerized into a polyacrylamide backbone. Adding and removing crosslinks via delivery of single-stranded DNA allows temporal, spatial, and reversible control of gel elasticity. We have shown in previous reports that dynamic modulation of DNA gel elasticity influences fibroblast and neuron behavior. In this report and video, we provide a schematic that describes the DNA gel crosslinking mechanisms and step-by-step instructions on the preparation DNA gels.

Our laboratory has developed DNA-crosslinked polyacrylamide hydrogels, a dynamic hydrogel system, to better understand the effects of modulating tissue stiffness on cell function. Here, we provide schematics, descriptions, and protocols to prepare these hydrogels.

Mechanobiology is an emerging scientific area that addresses the critical role of physical cues in directing cell morphology and function. For example, ...