This work was supported by an NSF Grant (2152065) and an NSF CAREER Award (1846857). The authors would like to thank all current and past members of the Wachs Lab for contributing to this protocol. Diagrams in Figure 1 were made in Biorender.

DRG harvest was performed in compliance with the Institutional Animal Care and Use Committee (IACUC) at the University of Nebraska-Lincoln. Female Sprague Dawley rats aged 12 weeks (~250 g) were used for the study. The details of the animals, reagents, and equipment used in the study are listed in the Table of Materials.
1. Multi-compartment device fabrication and assembly
<ol>
	<li>Computer-aided design and 3D printing of the multi-compartment device 
	NOTE: The MC device consists of three compartments for the isolation of DRG cell bodies and neurites (Figure 2A). The MC device is not commercially available; however, the STL file is provided herein (Supplementary Coding File 1) to enable 3D printing. Printing the MC takes one (1) day to complete.</li>
	<li>Upload the STL file of the MC to a Resin 3D printer using compatible software. 
	NOTE: The material ideal for printing is a high-temperature resin. A key advantage of using high-temperature resin is it is biocompatible and can be autoclaved and reused. A Digital light processing (DLP) 3D printer or other 3D printers could also be used to print the device.</li>
	<li>Transfer the printed devices into a washing solution for 30 min in isopropanol (&#62;96%) to remove residual resin from the surface of the printed MC device.</li>
	<li>Post-cure the devices at 80 &#176;C for 120 min using a commercially available curing agent (see Table of Materials). The curing agent enhances the mechanical properties of the printed MC device by exposing it to temperature and UV.</li>
	<li>Cure the MC devices thermally in an oven at 160 &#176;C for 180 min.</li>
	<li>Sterilize the devices using an autoclave for 45 min on a gravity cycle.</li>
</ol>
2. Hydrogel preparation
<ol>
	<li>Synthesize methacrylated hyaluronic acid (MAHA, 85%-115% methacrylation) and characterize using a protocol described by Seidlits et al.20. 
	NOTE: To support the growth of DRGs, triple hydrogels composed of methacrylated hyaluronic acid, type I collagen, and laminin are commonly used. Findings indicate that DRGs commonly grow in MAHA, ranging from 1.25-2.5 mg/mL and collagen from 2.0-4.5 mg/mL. For laminin, it is found that 0.75 mg/mL is enough to ensure robust growth of DRGs. Herein, methods to create a triple gel with 1.25 mg/mL MAHA, 4.5 mg/mL collagen, and 0.75 mg/mL laminin are detailed. Hydrogel preparation should be done in a laminar flow cabinet to provide an aseptic workspace.</li>
</ol>
3. Photoinitiator solution preparation
NOTE: A photoinitiator is necessary to crosslink the MAHA under UV light. It is commonly used at percentages from 0.3% to 0.6%.
<ol>
	<li>Prepare the photoinitiator solution 3 days prior to the DRG harvest.</li>
	<li>Pre-heat the sonicating water bath to 37 &#176;C before preparing the photoinitiator solution.</li>
	<li>Working with the sterile technique in a flow hood, prepare 23.125 mg/mL sodium bicarbonate (NaHCO3) solution by dissolving NaHCO3 in 5x Dulbecco&#39;s Modified Eagle Medium (DMEM)-HEPES solution.</li>
	<li>In a glass vial, mix 0.6% weight/volume (w/v) photoinitiator in a 20% vol/vol 1x sterile phosphate-buffered saline (PBS) and 80% vol/vol 5X DMEM/HEPES solution.</li>
	<li>To protect from light, wrap the glass vial in aluminum foil. However, ensure to remove the aluminum foil before transferring the vial to the sonicating water bath heated to 37 &#176;C with sonication between 30-40 min. 
	NOTE: Photoinitiator solution must be protected from light because it can break down into radicals and start the polymerization process when exposed to light21.</li>
	<li>Sterile filter the solution with a 0.22 &#181;m syringe filter into a new sterile glass vial.</li>
</ol>
4. Methacrylated hyaluronic acid dissolution
<ol>
	<li>Dissolve the methacrylated hyaluronic acid (prepared in step 2) on the same day the photoinitiator solution is prepared (step 3).</li>
	<li>Place the frozen lyophilized MAHA in a desiccator for 15 min to come to room temperature.</li>
	<li>Working in a laminar flow hood and using the sterile weighing technique, weigh the lyophilized MAHA in a glass vial.</li>
	<li>Add the necessary volume of filtered photoinitiator solution to MAHA to come to the desired MAHA concentration. The common final gel concentration is 1.25 mg/mL MAHA and 4.5 mg/mL collagen.</li>
	<li>Seal the cap with laboratory sealing film, wrap the glass vial with aluminum foil, and place it on an orbital shaker plate at room temperature until fully dissolved (usually 3 days). Occasionally, vortex the solution to break apart large clumps of MAHA to ensure uniform dissolution of MAHA.</li>
</ol>
5. Device assembly
<ol>
	<li>Place the MC device into a 48-well plate a day before the DRG harvest.</li>
	<li>Apply a thin line of silicone gel in the groove beneath the MC device (Figure 2B) using a 3 mL syringe. This will help the device to securely attach to the well plate.</li>
	<li>With the help of #3 forceps, gently place the MC device into the well of a 48-well plate, as shown in Figure 2C. 
	NOTE: The size of the device can be modified to suit the intended application, and a plate of a different size can be used if desired. The devices can be reused. To reuse, the hydrogel must be gently washed out from all tunnels, and the loading groove beneath the device must be used with 70% ethanol with gentle agitation. After this is complete, devices can be re-autoclaved to sterilize prior to use.</li>
</ol>
6. Animal preparation
<ol>
	<li>Autoclave all the necessary dissection tools prior to the start of harvest.</li>
	<li>Obtain an adult male or female Sprague Dawley rat aged 12-20 weeks. The sex of the rat does not impact DRG growth.</li>
	<li>Euthanize the rat according to the approved IACUC protocol using CO2 overdose as the primary method of euthanasia and bilateral pneumothorax puncture as a secondary method of euthanasia according to American Veterinary Medical Association Guidelines22.</li>
	<li>Place the rat on a sterile underpad on a dissecting table with the ventral side down. Spray the fur down with 70% ethanol.</li>
</ol>
7. Dorsal root ganglia (DRG) harvest
<ol>
	<li>Prepare trimming media containing 86% Neurobasal A media, 10% fetal bovine serum (FBS), 1% penicillin-streptomycin (PS), 1% Glutamine, and 2% B27 plus 50x and transfer into a 24-well plate. This media and well-plate will be used for temporary storage of harvested DRGs prior to embedding them in the hydrogel.</li>
	<li>Wear a lab coat, surgical mask, bonnet, safety glasses, and gloves.</li>
	<li>Using large blunt-nose scissors, cut through the skin at the base of the cervical spine and then down the length of the spine.</li>
	<li>To further drain blood away from the spinal canal, cut underneath the shoulder blades and down through the muscle to sever major blood vessels.</li>
	<li>Use large, sharp-nose scissors to cut the muscle away from the spine. Clear as much muscle as possible to visualize the spine.</li>
	<li>Remove the dorsal and lateral segments of the vertebrae. Use a straight Rongeur and sharp-nosed scissors to remove the spinous and transverse processes of the vertebrae, starting at the cervical end and moving toward the lumbar end.</li>
	<li>Remove the spinal cord. Use small sharp-nose scissors to carefully cut through the nerve roots connected to the DRGs along both sides of the spinal cord. Cut the cervical end of the spinal cord and carefully lift out of the spinal canal gradually, cutting through any remaining nerve roots connected to the DRGs. 
	NOTE: Fine motor skills are essential to ensure the DRG body is not severed in the process.</li>
	<li>To expose DRGs, use the curved Rongeur to remove more of the lateral parts of the vertebrae. Pull outward from the midline, alternating sides. Be careful not to cut through the DRG body since it may cause damage to the cell bodies and axons.</li>
	<li>Using the #3 forceps and straight-edge spring scissors, remove DRGs in between each level of the vertebrae. Grip the spinal nerve roots from the DRG, carefully pull them out of the pocket, and cut through the other end of the spinal nerve. Be careful not to grip the DRG body directly.</li>
	<li>Place the DRGs in wells of a 24-well plate (containing trimming media, step 7.1) on ice.</li>
	<li>After collecting all DRGs, clean up the workspace, place the carcass in an autoclave bag, and store/dispose of it appropriately according to IACUC protocol.</li>
</ol>
8. DRG trimming and cutting
<ol>
	<li>Fill a 60 mm Petri dish on a clean bench with trimming media (step 7.1). Set up the dissecting scope and glass bead sterilizer and place an autoclaved toolbox with trimming tools (#3 forceps and straight-edge spring scissors) by the scope.</li>
	<li>Under the dissecting scope, remove excess tissue around the DRG and trim the nerve roots close to the DRG body (typically slightly darker than the surrounding nerves).</li>
	<li>Fill a second 60 mm Petri dish with fresh media. Under the dissecting scope, cut the trimmed DRGs to about 0.5 mm (or between 0.5 mm and 0.8 mm). A ruler is placed under the Petri dish during cutting to have the correct estimated size of cut DRGs.</li>
	<li>Transfer cut DRGs into a fresh 24-well plate with the media on ice. 
	NOTE: DRG trimming, cutting, and embedding should be done in a laminar flow cabinet to provide an aseptic workspace. In the absence of a laminar flow cabinet, aseptic techniques must be observed, with minimum appropriate personal protective equipment (PPE) recommended (lab coat, masks, goggles, and gloves) and all tools sterilized throughout the process to mitigate potential contamination.</li>
</ol>
9. Collagen neutralization
<ol>
	<li>Neutralize the collagen on the same day as the DRG harvest after trimming and cutting the DRGs.</li>
	<li>Working on ice, pipette ~8.16 % ultra-pure water, ~1.84% 1 M sodium hydroxide (NaOH), ~10% 10x PBS, and ~80% of collagen of the total volume of solution into a glass vial.</li>
	<li>Mix slowly using a sterile low-retention pipette tip to pipette collagen (usually added last) into the mixture. Ensure to keep the pipette tip in the solution while mixing to avoid creating bubbles. Check the pH of the neutralized collagen using pH strips to be sure it is ~7. 
	NOTE: Neutralized collagen can be kept on ice for 2-3 h without gelling; therefore, hydrogel fabrication should be carried out expeditiously once collagen is neutralized.</li>
</ol>
10. Hydrogel fabrication
<ol>
	<li>Working on ice, pipette laminin (to a final concentration of 0.75 mg/mL) and 1x PBS into a glass vial.</li>
	<li>Using sterile low retention pipette tips, pipette neutralized collagen and MAHA solution into the mixture to arrive at the desired concentration.</li>
	<li>Mix slowly and check the pH of the resulting hydrogel (should be ~7).</li>
</ol>
11. DRG embedding
<ol>
	<li>Perform multi-compartment embedding.
	<ol>
		<li>Pipette 65.1 &#181;L and 52.8 &#181;L of the pre-mixed hydrogel into the soma and neurite compartments, respectively. 
		NOTE: This assumes the use of MC devices in a 48-well plate. These volumes can be changed based on the size of the MC device used.</li>
		<li>Gently embed the cut DRGs into the soma compartment (Figure 2A), ensuring they do not scratch the bottom of the well plate. Place the DRG in the middle so the neurites can grow into the adjacent compartments through the tunnels.</li>
		<li>Thermal crosslink the hydrogel in the incubator at 37 &#176;C for 30 min. While this occurs, turn on the UV lamp to warm up.</li>
		<li>After thermal crosslinking, UV crosslink the hydrogel for 90 s.</li>
		<li>Add DRG growth media to the hydrogel and DRG and perform a full media change after an hour to eliminate unreacted or remaining traces of photoinitiator that persist in the media.</li>
		<li>Perform a half media change every 3 days and monitor the growth of the DRGs by viewing under the microscope.</li>
		<li>After ~4 weeks, when DRG neurites begin to grow (Figure 3A), capture images with the fluorescent plate imager and quantify the length of neurites using FIJI (ImageJ).</li>
	</ol>
	</li>
</ol>
12. Control DRG embedding
<ol>
	<li>Pipette 250 &#181;L of hydrogel in a 48-well plate (without MC).</li>
	<li>Gently place the cut DRG in the middle of the well and halfway down into the hydrogel, ensuring it does not scratch the bottom of the well plate.</li>
	<li>Thermal crosslink the hydrogel in the incubator at 37 &#176;C for 30 min.</li>
	<li>UV crosslink the hydrogel for 90 s.</li>
	<li>Add DRG growth media to the hydrogel and DRG and perform a full media change after an hour.</li>
	<li>Perform half media change every 3 days and monitor the growth of the DRG by viewing it under the microscope.</li>
	<li>After ~4 weeks, when DRG neurites begin to grow (Figure 3B), capture images with the fluorescent plate imager. 
	NOTE: Any brightfield microscope will work for imaging. An automated plate-based microscope makes this process faster. Control embedding in plain gels without the device is done to compare the growth of DRG neurites in multi-compartment. Once users validate that DRGs grow similarly in MC and control gels, they may not need to repeat control gels each time.</li>
</ol>
13. DRG imaging
<ol>
	<li>Capture brightfield images at 4x magnification using a fluorescent plate imager at a range of focal distances to visualize the entire DRG and MC Device.</li>
	<li>Adjust the brightness and contrast of the image to make it easier to view the neurites.</li>
	<li>Save the image as a .tiff file.</li>
</ol>
14. DRG neurite quantification
<ol>
	<li>Download and install FIJI (ImageJ).</li>
	<li>Open ImageJ. Open the image file and ensure that the file is monochrome.</li>
	<li>Enhance the contrast so that the smallest neurites are visible. Do this by opening the brightness and contrast option using the shortcut Ctrl+Shift+C.</li>
	<li>Adjust the sliders of the controller as needed to visualize the neurites, and click on Apply for the desired brightness and contrast.</li>
	<li>Open the neurite tracer by opening the Plugins menu, selecting Segmentation, and clicking on Simple Neurite Tracer.</li>
	<li>Start a new trace by clicking on one end of the neurite right up against the DRG body.</li>
	<li>Click another point along the neurite to add a trace until the neurite is traced end to end. Click on Finish Path after tracing the neurite.</li>
	<li>Convert the traced length to real units and save the results by copying and pasting them into Excel.</li>
	<li>Use the straight-line tool in FIJI to draw a line of the same length as the scale bar (not letting go of the end of the line) and look at the length value visible under the toolbar. The length value is used for the conversion.</li>
	<li>Measure the length of six long neurites extending from both sides of the DRG (Figure 3C,D) and calculate the average length of these neurites to give a representative average length of the DRG, as shown in Figure 4. 
	NOTE: Immunostaining has been tested on DRG explants in plain gels to show the presence of non-neuronal support cells7. DRGs are fixed in 4% paraformaldehyde (PFA) at room temperature, washed three times with 1x PBS for 15 min, and stored in 1x PBS at 4 &#176;C until immunostaining.</li>
	<li>For explants in MC devices, remove the MC device and follow the same process described above7.</li>
</ol>

Advanced In Vitro Model for Studying Neuronal Hypersensitivity in DRG Cultures

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protocol+for+dissection+and+culture+of+murine+dorsal+root+ganglia+neurons+to+study+neuropeptide+release.">Protocol for dissection and culture of murine dorsal root ganglia neurons to study neuropeptide release.</a> STAR Protocols. 2 (1), 100333 (2021).</li><li>Bai, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+high-resolution+anatomical+rat+atlas.">A high-resolution anatomical rat atlas.</a> J Anatomy. 209 (5), 707-708 (2006).</li><li>Bian, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+influence+of+hyaluronic+acid+hydrogel+crosslinking+density+and+macromolecular+diffusivity+on+human+MSC+chondrogenesis+and+hypertrophy.">The influence of hyaluronic acid hydrogel crosslinking density and macromolecular diffusivity on human MSC chondrogenesis and hypertrophy.</a> Biomaterials. 34 (2), 413-421 (2013).</li><li>Haberberger, R. V., Barry, C., Dominguez, N., Matusica, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+dorsal+root+ganglia.">Human dorsal root ganglia.</a> Front Cell Neurosci. 13, 271 (2019).</li><li>Schwaid, A. G., Krasowka-Zoladek, A., Chi, A., Cornella-Taracido, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+the+rat+and+human+dorsal+root+ganglion+proteome.">Comparison of the rat and human dorsal root ganglion proteome.</a> Scientific Rep. 8 (1), 13469 (2018).</li></ol>

The authors of this study declare that they have no conflict of interest.

This protocol outlines a method to harvest adult Sprague Dawley DRGs and culture them in 3D natural hydrogels. In contrast to this method, other approaches to harvesting DRGs from mice and rats involve isolating the spinal column. The excised spinal column is halved, and the spinal cord is removed to expose DRGs23,24,25. Damage to the spinal cord limits blood supply, which affects DRGs and internal neurons26</s...

Chronic pain is the leading cause of disability and loss of work globally1. Chronic pain affects about 20% of adults globally and imposes a significant societal and economic burden2, with total costs estimated between $560 and $635 billion every year in the United States3.
The main peripheral feature exhibited by chronic pain patients is a lowered stimulation threshold of nerves, which leads to the nervous system being more responsive to stimuli4,5. The lowered stimulation threshold can result in a painful response to a previously non-painful stimulus (allodynia) or a heightened response to a painful stimulus (hyperalgesia)6. Current chronic pain treatments have limited efficacy, and treatments that succeed in animal models often fail in human trials due to mechanistic differences in pain manifestation7. In vitro models that can more accurately mimic peripheral sensitization mechanisms have the potential to increase the translation of new therapeutics8,9. Further, by modeling key aspects of sensitized nerves in a culture system, researchers could develop a deeper understanding of the mechanisms that drive lowered thresholds and identify novel therapeutic targets that reverse them10.
The ideal in vitro platforms or microphysiological systems would incorporate the physical separation of distal neurites and dorsal root ganglia (DRG) cell body, a three-dimensional (3D) cellular environment, and the presence of native support cells to closely mimic in vivo conditions. However, a recent paper by Caparaso et al.11 shows that current DRG culture platforms lack one or more of these key features, making them insufficient in replicating in vivo conditions. Even though these platforms are easy to set up, they do not mimic the biological basis of peripheral sensitization and thus may not translate to in vivo efficacy. To address this limitation, a physiologically relevant in vitro model has been developed for the culture of dorsal root ganglia (DRG) within a hydrogel matrix with three isolated compartments to allow temporal fluidic isolation of neurites and DRG cell bodies11. This model offers both physiological and anatomical relevance, which has the potential to study peripheral sensitization of neurons in vitro.
The growing interest in the use of DRG explants in a 3D culture is due to their ability to facilitate robust neurite growth, which serves as an indirect indicator of DRG viability12. While primary neonatal or embryonic DRG explants are predominantly used in current in vitro culture platforms13,14, using explants from adult rodents provides a better model of mature neuronal physiology, which closely mimics human DRG physiology compared to explants from neonatal or embryonic rodents15. Explant DRGs refer to the preservation of the cellular and molecular tissue of native DRG tissue, primarily by maintaining native non-neuronal support cells.Herein, this protocol describes the methodology to harvest and culture DRG explants from adult Sprague Dawley rats in a multi-compartment (MC) device (Figure 1).
Efficacy has been shown in culturing DRGs from the cervical, thoracic, and lumbar spine with no observable differences in neurite growth. For this application, the objective was to elicit neurite growth into the outer compartments of the device; therefore, this article did not discriminate among DRG levels. However, if needed for a specific experiment, the DRG level can be tailored to meet experimenters&#39; needs. There are currently other compartmentalized culture models for the 3D culture of DRGs16, however, these devices do not contain preserved native non-neuronal support cells, which can limit translation. Preserving the native structure of harvested DRGs is important because it ensures the retention of non-neuronal support cells, whose interactions with DRG neurons are essential for maintaining the functional properties of these neurons. Several studies co-cultured dissociated DRGs with non-native neuronal cells such as Schwann cells to promote myelination of neurons17,18,19.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>#5 forceps</td>
			<td>Fine Science Tools</td>
			<td>11252-00</td>
			<td>For trimming and cutting DRG</td>
		</tr>
		<tr>
			<td>10x DMEM</td>
			<td>MilliporeSigma</td>
			<td>D2429</td>
			<td></td>
		</tr>
		<tr>
			<td>1x PBS (autoclaved)</td>
			<td>Prepared in lab</td>
			<td></td>
			<td>7.3 - 7.5 pH</td>
		</tr>
		<tr>
			<td>24 well plates</td>
			<td>VWR</td>
			<td>82050-892</td>
			<td>To temporarily store harvested and cut DRGs</td>
		</tr>
		<tr>
			<td>3 mL Syringe sterile, single use</td>
			<td>BD</td>
			<td>309657</td>
			<td></td>
		</tr>
		<tr>
			<td>48 well plates</td>
			<td>Greiner Bio-One</td>
			<td>677180</td>
			<td></td>
		</tr>
		<tr>
			<td>60 mm Petri dish</td>
			<td>Fisher Scientific</td>
			<td>FB0875713A</td>
			<td>To hold media for trimming and cutting</td>
		</tr>
		<tr>
			<td>Aluminium foil</td>
			<td>Fisherbrand</td>
			<td>01-213-104</td>
			<td></td>
		</tr>
		<tr>
			<td>B27 Plus 50x</td>
			<td>ThermoFisher</td>
			<td>17504044</td>
			<td>For DRG media</td>
		</tr>
		<tr>
			<td>Collagen type I</td>
			<td>Ibidi</td>
			<td>50205</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved cup Friedman Pearson Rongeur</td>
			<td>Fine Science Tools</td>
			<td>16221-14</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Dumont #3 forceps</td>
			<td>Fine Science Tools</td>
			<td>11293-00</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>ThermoFisher</td>
			<td>16000044</td>
			<td>For DRG media</td>
		</tr>
		<tr>
			<td>Form cure</td>
			<td>Form Labs</td>
			<td></td>
			<td>curing agent</td>
		</tr>
		<tr>
			<td>Form wash</td>
			<td>Form Labs</td>
			<td></td>
			<td>To wash excess resins off MC</td>
		</tr>
		<tr>
			<td>Glass bead sterilizer</td>
			<td>Fisher Scientific</td>
			<td>NC9531961</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass vials (8 mL)</td>
			<td>DWK Life Sciences (Wheaton)</td>
			<td>224724</td>
			<td></td>
		</tr>
		<tr>
			<td>GlutaMax</td>
			<td>ThermoFisher</td>
			<td>35050-061</td>
			<td>For DRG media</td>
		</tr>
		<tr>
			<td>HEPES (1M)</td>
			<td>Millipore Sigma</td>
			<td>H0887</td>
			<td></td>
		</tr>
		<tr>
			<td>High temp V2 resin</td>
			<td>FormLabs</td>
			<td>FLHTAM02</td>
			<td></td>
		</tr>
		<tr>
			<td>Hyaluronic Acid Sodium Salt</td>
			<td>MilliporeSigma</td>
			<td>53747</td>
			<td>Used to make MAHA</td>
		</tr>
		<tr>
			<td>Irgacure</td>
			<td>MilliporeSigma</td>
			<td>410896</td>
			<td></td>
		</tr>
		<tr>
			<td>Laminin</td>
			<td>R&#38;D Systems</td>
			<td>344600501</td>
			<td></td>
		</tr>
		<tr>
			<td>Large blunt-nose scissors</td>
			<td>Militex</td>
			<td>EG5-26</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Large forceps (serrated tips)</td>
			<td>Militex</td>
			<td>9538797</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Large sharp-nosed scissors</td>
			<td>Fine Science Tools</td>
			<td>14010-15</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Low Retention pipette tips</td>
			<td>Fisher Scientific</td>
			<td>02-707-017</td>
			<td>For pipetting collagen and MAHA</td>
		</tr>
		<tr>
			<td>Methacrylated hyaluronic acid (MAHA)</td>
			<td>Prepared in lab</td>
			<td>N/A</td>
			<td>85 - 115 % methacrylation</td>
		</tr>
		<tr>
			<td>Nerve Growth Factor (NGF)</td>
			<td>R&#38;D Systems</td>
			<td>556-NG-100</td>
			<td>For DRG media</td>
		</tr>
		<tr>
			<td>Neurobasal A Media</td>
			<td>ThermoFisher</td>
			<td>10888022</td>
			<td>For DRG media</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Bemis</td>
			<td>PM996</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Bemis</td>
			<td>PM996</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin (PS)</td>
			<td>EMD Millipore</td>
			<td>516106</td>
			<td>For DRG media</td>
		</tr>
		<tr>
			<td>pH test strips</td>
			<td>VWR International</td>
			<td>BDH35309.606</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips (1000 &#181;L)</td>
			<td>USA Scientific</td>
			<td>1111-2021</td>
			<td></td>
		</tr>
		<tr>
			<td>Preform 3.23.1 software</td>
			<td>Formslab</td>
			<td></td>
			<td>To upload STL file</td>
		</tr>
		<tr>
			<td>Rat</td>
			<td>Charles River</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Resin 3D printer</td>
			<td>Form Labs</td>
			<td>Form 3L</td>
			<td>3D printing MC device</td>
		</tr>
		<tr>
			<td>Small sharp-nosed scissors</td>
			<td>Fine Science Tools</td>
			<td>14094-11</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>MilliporeSigma</td>
			<td>S6014</td>
			<td></td>
		</tr>
		<tr>
			<td>Straight cup rongeur</td>
			<td>Fine Science Tools</td>
			<td>16004-16</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Straight edge spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15024-10</td>
			<td>For dissection</td>
		</tr>
		<tr>
			<td>Surgical Scaplel blade (No. 10)</td>
			<td>Fisher Scientific</td>
			<td>22-079-690</td>
			<td></td>
		</tr>
		<tr>
			<td>Syring filters, PES (0.22 &#181;m)</td>
			<td>Celltreat</td>
			<td>229747</td>
			<td></td>
		</tr>
		<tr>
			<td>Tiny spring scissors</td>
			<td>World Precision Instruments</td>
			<td>14003</td>
			<td>For trimming and cutting DRG</td>
		</tr>
		<tr>
			<td>UV lamp</td>
			<td>Analytik Jena US</td>
			<td></td>
			<td>To photocrosslink hydrogel (15 - 18 mW/cm2)</td>
		</tr>
	</tbody>
</table>

harvesting, embedding, and culturing dorsal root ganglia in multi-compartment devices to study peripheral neuronal features

The most common peripheral neuronal feature of pain is a lowered stimulation threshold or hypersensitivity of terminal nerves from the dorsal root ganglia (DRG). One proposed cause of this hypersensitivity is associated with the interaction between immune cells in the peripheral tissue and neurons. In vitro models have provided foundational knowledge in understanding how these mechanisms result in nociceptor hypersensitivity. However, in vitro models face the challenge of translating efficacy to humans. To address this challenge, a physiologically and anatomically relevant in vitro model has been developed for the culture of intact dorsal root ganglia (DRGs) in three isolated compartments in a 48-well plate. Primary DRGs are harvested from adult Sprague Dawley rats after humane euthanasia. Excess nerve roots are trimmed, and the DRG is cut into appropriate sizes for culture. DRGs are then grown in natural hydrogels, enabling robust growth in all compartments. This multi-compartment system offers anatomically relevant isolation of the DRG cell bodies from neurites, physiologically relevant cell types, and mechanical properties to study the interactions between neural and immune cells. Thus, this culture platform provides a valuable tool for investigating treatment isolation strategies, ultimately leading to an improved screening approach for predicting pain.

The present protocol described a technique to harvest and culture DRG from adult Sprague Dawley rats in a multi-compartment (MC) device. As shown&#160;in Figure 1, DRG harvested from adult rats was trimmed and cut into ~0.5 mm. The trimmed and cut DRGs were then embedded in a hydrogel in the soma region of the MC device (Figure 2) and cultured for 27 days before neurite quantification. DRG was cultured in plain gel to serve as the control. The c...

Author Spotlight: In Vitro and In Vivo Models to Enhance Chronic Pain Treatment Efficacy

This protocol provides a technique to harvest and culture explanted dorsal root ganglion (DRG) from adult Sprague Dawley rats in a multi-compartment (MC) device.

Harvesting, Embedding, and Culturing Dorsal Root Ganglia in Multi-compartment Devices to Study Peripheral Neuronal Features

The most common peripheral neuronal feature of pain is a lowered stimulation threshold or hypersensitivity of terminal nerves from the dorsal root ganglia ...

harvesting-embedding-culturing-dorsal-root-ganglia-multi-compartment

Research

JoVE Journal

Bioengineering

733 Views.  University of Nebraska-Lincoln. This protocol provides a technique to harvest and culture explanted dorsal root ganglion (DRG) from adult Sprague Dawley rats in a multi-compartment (MC) device.

Video: Author Spotlight: In Vitro and In Vivo Models to Enhance Chronic Pain Treatment Efficacy

Demonstrating the Uses of the Novel Gravitational Force Spectrometer to Stretch and Measure Fibrous Proteins

The study of macromolecular structure has become critical to the elucidation of molecular mechanisms and function. There are several limited, but important bioinstruments capable of testing the force dependence of structural features in proteins. Scale has been a limiting parameter on how accurately researchers can peer into the nanomechanical world of molecules, such as nucleic acids, enzymes, and motor proteins that perform life-sustaining work. Atomic force microscopy (AFM) is well tuned to determine native structures of fibrous proteins with a distance resolution on par with electron microscopy. However, in AFM force studies, the forces are typically much higher than a single molecule might experience 1, 2. Optical traps (OT) are very good at determining the relative distance between the trapped beads and they can impart very small forces 3. However, they do not yield accurate absolute lengths of the molecules under study. Molecular simulations provide supportive information to such experiments, but are limited in the ability to handle the same large molecular sizes, long time frames, and convince some researchers in the absence of other supporting evidence2, 4.
The gravitational force spectrometer (GFS) fills a critical niche in the arsenal of an investigator by providing a unique combination of abilities. This instrument is capable of generating forces typically with 98% or better accuracy from the femtonewton range to the nanonewton range. The distance measurements currently are capable of resolving the absolute molecular length down to five nanometers, and relative bead pair separation distances with a precision similar to an optical trap. Also, the GFS can determine stretching or uncoiling where the force is near equilibrium, or provide a graded force to juxtapose against any measured structural changes. It is even possible to determine how many amino acid residues are involved in uncoiling events under physiological force loads 2. Unlike in other methods where there is extensive force calibration that must precede any assay, the GFS requires no such force calibration 5. By complementing the strengths of other methods, the GFS will bridge gaps in understanding the nanomechanics of vital proteins and other macromolecules.

This is a step-by step guide showing the purpose, operation, and representative results from the novel gravitational force spectrometer.

The study of macromolecular structure has become critical to the elucidation of molecular mechanisms and function. There are several limited, but ...

Impulsive Pressurization of Neuronal Cells for Traumatic Brain Injury Study

A novel impulsive cell pressurization experiment has been developed using a Kolsky bar device to investigate blast-induced traumatic brain injury (TBI). We demonstrate in this video article how blast TBI-relevant impulsive pressurization is applied to the neuronal cells in vitro. This is achieved by using well-controlled pressure pulse created by a specialized Kolsky bar device, with complete pressure history within the cell pressurization chamber recorded. Pressurized neuronal cells are inspected immediately after pressurization, or further incubated to examine the long-term effects of impulsive pressurization on neurite/axonal outgrowth, neuronal gene expression, apoptosis, etc. We observed that impulsive pressurization at about 2 MPa induces distinct neurite loss relative to unpressurized cells. Our technique provides a novel method to investigate the molecular/cellular mechanisms of blast TBI, via impulsive pressurization of brain cells at well-controlled pressure magnitude and duration.

A novel impulsive cell pressurization experiment has been developed using a Kolsky bar device to investigate the molecular/cellular mechanisms of blast-induced traumatic brain injury.

A novel impulsive cell pressurization experiment has been developed using a Kolsky bar device to investigate blast-induced traumatic brain injury (TBI). ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Density Gradient Multilayered Polymerization (DGMP): A Novel Technique for Creating Multi-compartment, Customizable Scaffolds for Tissue Engineering

Complex tissue culture matrices, in which types and concentrations of biological stimuli (e.g. growth factors, inhibitors, or small molecules) or matrix structure (e.g. composition, concentration, or stiffness of the matrix) vary over space, would enable a wide range of investigations concerning how these variables affect cell differentiation, migration, and other phenomena. The major challenge in creating layered matrices is maintaining the structural integrity of layer interfaces without diffusion of individual components from each layer1. Current methodologies to achieve this include photopatterning2-3, lithography4, sequential functionalization5, freeze drying6, microfluidics7, or centrifugation8, many of which require sophisticated instrumentation and technical skills. Others rely on sequential attachment of individual layers, which may lead to delamination of layers9. 
DGMP overcomes these issues by using an inert density modifier such as iodixanol to create layers of varying densities10. Since the density modifier can be mixed with any prepolymer or bioactive molecule, DGMP allows each scaffold layer to be customized. Simply varying the concentration of the density modifier prevents mixing of adjacent layers while they remain aqueous. Subsequent single step polymerization gives rise to a structurally continuous multilayered scaffold, in which each layer has distinct chemical and mechanical properties. The density modifier can be easily removed with sufficient rinsing without perturbation of the individual layers or their components. This technique is therefore well suited for creating hydrogels of various sizes, shapes, and materials.
A protocol for fabricating a 2D-polyethylene glycol (PEG) gel, in which alternating layers incorporate RGDS-350, is outlined below. We use PEG because it is biocompatible and inert. RGDS, a cell adhesion peptide11, is used to demonstrate spatial restriction of a biological cue, and the conjugation of a fluorophore (Alexa Fluor 350) enables us to visually distinguish various layers. This procedure can be adapted for other materials (e.g. collagen, hyaluronan, etc.) and can be extended to fabricate 3D gels with some modifications10.

Density Gradient Multilayered Polymerization DGMP: A Novel Technique for Creating Multi-compartment, Customizable Scaffolds for Tissue Engineering

Here we describe a unique strategy for creating biocompatible, layered matrices with continuous interfaces between distinct layers for tissue engineering. Such a scaffold could provide an ideal customizable environment to modulate cell behavior by various biological, chemical or mechanical cues

Complex tissue culture matrices, in which types and concentrations of biological stimuli (e.g. growth factors, inhibitors, or small molecules) or matrix ...

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D culture of cells in collagen type I gels helps to stabilize cell morphology and function, which is necessary for creating microfluidic tissue models in microdevices. Translating traditional 3D culture techniques for tissue culture plates to microfluidic devices is often difficult because of the limited channel dimensions. In this method, we describe a technique for modifying native type I collagen to generate polycationic and polyanionic collagen solutions that can be used with layer-by-layer deposition to create ultrathin collagen assemblies on top of cells cultured in microfluidic devices. These thin collagen layers stabilize cell morphology and function, as shown using primary hepatocytes as an example cell, allowing for the long term culture of microtissues in microfluidic devices.

The creation of functional microtissues within microfluidic devices requires the stabilization of cell phenotypes by adapting traditional cell culture techniques to the limited spatial dimensions in microdevices. Modification of collagen allows the layer-by-layer deposition of ultrathin collagen assemblies that can stabilize primary cells, such as hepatocytes, as microfluidic tissue models.

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D ...

Fabricating Cotton Analytical Devices

A robust, low-cost analytical device should be user-friendly, rapid, and affordable. Such devices should also be able to operate with scarce samples and provide information for follow-up treatment. Here, we demonstrate the development of a cotton-based urinalysis (i.e., nitrite, total protein, and urobilinogen assays) analytical device that employs a lateral flow-based format, and is inexpensive, easily fabricated, rapid, and can be used to conduct multiple tests without cross-contamination worries. Cotton is composed of cellulose fibers with natural absorptive properties that can be leveraged for flow-based analysis. The simple but elegant fabrication process of our cotton-based analytical device is described in this study. The arrangement of the cotton structure and test pad takes advantage of the hydrophobicity and absorptive strength of each material. Because of these physical characteristics, colorimetric results can persistently adhere to the test pad. This device enables physicians to receive clinical information in a timely manner and shows great potential as a tool for early intervention.

To investigate simple fabrication approaches for multiple assay needs, we created a fluid-absorbing channel system made of cotton material. This device was used to establish a multiple detection platform, and solve contamination issues that commonly affect lateral flow-based biomedical devices, for clinical urinalysis of nitrite, total protein, and urobilinogen.

A robust, low-cost analytical device should be user-friendly, rapid, and affordable. Such devices should also be able to operate with scarce samples and ...

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol adhesive through a metal stencil, the overall amount of adhesive used in assembling paper microfluidic devices can be significantly reduced. We show on a simple 4-layer planar paper microfluidic device that the optimal adhesive application technique and device construction style depends heavily on desired performance characteristics. By moderately increasing the overall area of a device, it is possible to dramatically decrease the wicking time and increase device success rates while also reducing the amount of adhesive required to keep the device together. Such adhesive application also causes the adhesive to form semi-permanent bonds instead of permanent bonds between paper layers, enabling single-use devices to be non-destructively disassembled after use. Nonplanar 3D origami devices also benefit from the semi-permanent bonds during folding, as it reduces the likelihood that unrelated faces may accidently stick together. Like planar devices, nonplanar structures see reduced wicking times with patterned adhesive application vs uniformly applied adhesive.

We demonstrate the use of patterned aerosol adhesives to construct 3D paper microfluidic devices. This method of adhesive application forms semi-permanent bonds between layers, enabling single-use devices to be non-destructively disassembled after use and to ease folding complex nonplanar structures.

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol ...

Automated Robotic Liquid Handling Assembly of Modular DNA Devices

Recent advances in modular DNA assembly techniques have enabled synthetic biologists to test significantly more of the available &#34;design space&#34; represented by &#34;devices&#34; created as combinations of individual genetic components. However, manual assembly of such large numbers of devices is time-intensive, error-prone, and costly. The increasing sophistication and scale of synthetic biology research necessitates an efficient, reproducible way to accommodate large-scale, complex, and high throughput device construction.
Here, a DNA assembly protocol using the Type-IIS restriction endonuclease based Modular Cloning (MoClo) technique is automated on two liquid-handling robotic platforms. Automated liquid-handling robots require careful, often times tedious optimization of pipetting parameters for liquids of different viscosities (e.g. enzymes, DNA, water, buffers), as well as explicit programming to ensure correct aspiration and dispensing of DNA parts and reagents. This makes manual script writing for complex assemblies just as problematic as manual DNA assembly, and necessitates a software tool that can automate script generation. To this end, we have developed a web-based software tool, http://mocloassembly.com, for generating combinatorial DNA device libraries from basic DNA parts uploaded as Genbank files. We provide access to the tool, and an export file from our liquid handler software which includes optimized liquid classes, labware parameters, and deck layout. All DNA parts used are available through Addgene, and their digital maps can be accessed via the Boston University BDC ICE Registry. Together, these elements provide a foundation for other organizations to automate modular cloning experiments and similar protocols.
The automated DNA assembly workflow presented here enables the repeatable, automated, high-throughput production of DNA devices, and reduces the risk of human error arising from repetitive manual pipetting. Sequencing data show the automated DNA assembly reactions generated from this workflow are ~95% correct and require as little as 4% as much hands-on time, compared to manual reaction preparation.

Here, an automated workflow to perform modular DNA &#34;device&#34; assembly using a modular cloning DNA assembly method on liquid-handling robots is presented. The protocol uses an intuitive software tool for generating liquid handler picklists for combinatorial DNA device library generation, which we demonstrate using two liquid handling platforms.

Recent advances in modular DNA assembly techniques have enabled synthetic biologists to test significantly more of the available &#34;design space&#34; ...

Melt Electrospinning Writing of Three-dimensional Poly(&#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology with great potential for biomedical applications. The technique facilitates the direct deposition of biocompatible polymer fibers to fabricate well-ordered scaffolds in the sub-micron to micro scale range. The establishment of a stable, viscoelastic, polymer jet between a spinneret and a collector is achieved using an applied voltage and can be direct-written. A significant benefit of a typical porous scaffold is a high surface-to-volume ratio which provides increased effective adhesion sites for cell attachment and growth. Controlling the printing process by fine-tuning the system parameters enables high reproducibility in the quality of the printed scaffolds. It also provides a flexible manufacturing platform for users to tailor the morphological structures of the scaffolds to their specific requirements. For this purpose, we present a protocol to obtain different fiber diameters using melt electrospinning writing (MEW) with a guided amendment of the parameters, including flow rate, voltage and collection speed. Furthermore, we demonstrate how to optimize the jet, discuss often experienced technical challenges, explain troubleshooting techniques and showcase a wide range of printable scaffold architectures.

This protocol serves as a comprehensive guideline to fabricate scaffolds via electrospinning with polymer melts in a direct writing mode. We systematically outline the process and define the appropriate parameter settings for achieving targeted scaffold architectures.

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology ...

Development of Microfluidic Devices to Study the Elongation Capability of Tip-growing Plant Cells in Extremely Small Spaces

In vivo, tip-growing plant cells need to overcome a series of physical barriers; however, researchers lack the methodology to visualize cellular behavior in such restrictive conditions. To address this issue, we have developed growth chambers for tip-growing plant cells that contain a series of narrow, micro-fabricated gaps (~1 &#181;m) in a poly-dimethylsiloxane (PDMS) substrate. This transparent material allows the user to monitor tip elongation processes in individual cells during microgap penetration by time-lapse imaging. Using this experimental platform, we observed morphological changes in pollen tubes as they penetrated the microgap. We captured the dynamic changes in the shape of a fluorescently labeled vegetative nucleus and sperm cells in a pollen tube during this process. Furthermore, we demonstrated the capability of root hairs and moss protonemata to penetrate the 1 &#181;m gap. This in vitro platform can be used to study how individual cells respond to physically constrained spaces and may provide insights into tip-growth mechanisms.

We describe a method to investigate the capability of tip-growing plant cells, including pollen tubes, root hairs, and moss protonemata, to elongate through extremely narrow gaps (~1 &#181;m) in a microfluidic device.

In vivo, tip-growing plant cells need to overcome a series of physical barriers; however, researchers lack the methodology to visualize cellular behavior ...