The authors wish to acknowledge support from R37 DK045729 (to JLM), R01 AR060837 (to HX) and the University of Michigan Gastrointestinal Research Center Molecular Core P30 DK034933.

All animal experiments described were approved by the University of Michigan&#39;s Committee on the Use and Care of Animals.
1. Preparation of Sterile Poly-D-lysine (PDL) and Laminin Solutions
<ol>
	<li>At least one day prior to cell isolation, prepare poly-D-lysine (PDL) and laminin coated plates. 
	NOTE: Both 6-well and 12-well plates were prepared depending on the experimental goals. Typically, 12-well plates were used for the quantitative analysis using western blots; whereas, 6-well plates were used to hold autoclaved (sterile) coverslips for immunohistochemistry. 24-well plates were used to culture EGCs for Ca2+ flux imaging.</li>
	<li>Dilute stock solutions with ultrapure tissue culture grade, DNase-free and RNase-free water in a tissue culture hood with HEPA-filtered air.</li>
	<li>Sterilizing glass coverslips
	<ol>
		<li>Hold the coverslip with sterile forceps and immerse coverslips in a beaker of 100% ethanol for 15 min. Allow the ethanol to evaporate in the tissue culture hood and then place in the 6-well plates.</li>
		<li>Alternatively, place several coverslips individually on 2 mm thick blotting paper in a plastic container and then autoclave.</li>
	</ol>
	</li>
</ol>
2. Coating Plates
NOTE: perform these steps under a sterile tissue culture laminar flow hood.
<ol>
	<li>Thaw 1 mg/mL PDL stock and dilute 1:10 with sterile, tissue culture grade water so that the final concentration is 100 &#181;g/mL. Use 2 mL per well to coat 6-well plates, 1 mL to cover one well of a 12-well plate and 0.5 mL for one well in the 24-well plates. Store the remaining diluted PDL in 12 mL aliquots at -20 &#176;C.</li>
	<li>After allowing the PDL to coat the wells for at least 1 h, remove the PDL with a sterile pipette. Allow the plates to air dry completely under a tissue culture laminar flow hood. 
	NOTE: After removing the PDL solution with a sterile pipette, it can be used twice more within 1 week after passing through a 0.22 &#181;m filter and storing at 4 &#176;C.</li>
	<li>Thaw the laminin stock on ice to avoid gel formation. 
	NOTE: Undiluted laminin becomes solid when warmed above 8 &#176;C.</li>
	<li>Dilute the laminin stock (0.5 mg/mL) 1:50 with sterile Dulbecco&#39;s Phosphate Buffered Saline (DPBS) to obtain a final concentration of 10 &#181;g/mL. Store the diluted laminin in 12 mL aliquots at -20 &#176;C.</li>
	<li>Add diluted laminin to the wells containing dried PDL. Use 1 mL to cover the 6-well surface and 0.5 mL to cover the bottom of the 12-well plates. For coverslips, add 400 &#181;L of diluted laminin to the coverslips to achieve surface coverage.</li>
	<li>Incubate the coated plates at 37 &#176;C for 2 h, if the plate is used the same day (quick coating). If not, seal the plates with plastic wrap, and store overnight at 4 &#176;C (slow coating). 
	NOTE: Sealing is critical to avoid drying and contamination.</li>
	<li>Remove the laminin solution carefully with a pipette to avoid scratching the surface. Gently wash the wells three times with sterile DPBS.</li>
	<li>Add complete glial growth media to each well and maintain the plates at 37 &#176;C until ready to add the EGC suspension. 
	NOTE: Poly-D lysine and laminin coated plates can be stored for 3 days at 4 &#176;C. Coat the plates several days ahead and then store at 4 &#176;C, wash the plates with sterile DPBS three times. Add 2 mL of sterile DPBS to each well after the final wash. Seal the plate with parafilm and store at 4 &#176;C. It is essential to ensure that the laminin does not dry out.</li>
</ol>
3. Preparation of Isolation Solutions
<ol>
	<li>Prepare the EDTA/HEPES/DPBS dissociation solution. For 500 mL of the solution, use sterile DPBS (without calcium and magnesium) to make a final solution of 10 mM HEPES and 5 mM EDTA. To 490 mL of the DPBS, add 5 mL of 1 M HEPES buffer and 5 mL of 0.5 M EDTA stock solution. Store at 4 &#176;C until use.</li>
	<li>Prepare the glial cell resuspension media using the reagents listed in Table 1.</li>
	<li>Prepare glial growth media using the reagents listed in Table 2. 
	NOTE: Glial growth media containing GDNF can be stored for one week at 4 &#176;C.</li>
</ol>
4. Removal of the Intestinal Segment
NOTE: Mice were allowed free access to the food and water prior to euthanization by the isoflurane drop method and removal of a vital organ. C57BL/6 mice greater than 8 weeks of age were used. Both genders were used without noticeable differences in the preparation.
<ol>
	<li>Euthanize the mouse with an overdose of isoflurane and place supine in a dissecting tray. Pin the extremities and sterilize the abdomen with 70% ethanol. Tent the skin with forceps and then open the abdomen with scissors.</li>
	<li>Lift the liver and identify the distal stomach/pylorus. Then remove 7 cm of the proximal intestine. Be careful not to tear the intestine.</li>
	<li>Hold the pylorus with forceps while snipping away the mesentery.
	<ol>
		<li>Use blunt dissection with the back of the scissor to scrape away adherent pancreas.</li>
		<li>Place the intestinal segment into ice-cold DPBS without Ca2+ or Mg2+ (Figure 1A). 
		NOTE: Other segments of the intestine or colon can also be removed using the same technique.</li>
	</ol>
	</li>
	<li>Use a 5 or 10 mL syringe attached to a blunt end 20 G needle to flush out the fecal contents with ice cold DPBS (Figure 1B).</li>
	<li>Divide the intestine into 3 cm segments to remove the longitudinal muscle using the technique initially described by Smith et al.12 and as modified in Sundaresan et al.10. 
	NOTE: Intestinal segments from up to two mice are used per 30 mL of the HEPES/EDTA/DPBS isolation solution.</li>
	<li>Break off about 4 cm to separate the wooden stick from the cotton tip. Soak the wooden stick in DPBS (Figure 1C).</li>
	<li>Slip one of the intestinal segments smoothly onto the wetted stick (Figure 2A-C).</li>
	<li>Remove adherent mesentery still attached using needle-nose forceps (Figure 2D).</li>
	<li>Use a clean razor blade or scalpel to make two longitudinal nicks along the intestine where the mesentery was attached (Figure 2E).
	<ol>
		<li>Wet the cotton tip with DPBS. Rub the wetted cotton tip longitudinally along the muscle to loosen the longitudinal muscle/myenteric plexus (LMMP). Then move the cotton tip horizontally to tease away the LMMP from the circular muscle along the length of the intestinal segment during LMMP removal.</li>
		<li>Hold the tip of the stick between the thumb and forefinger while stabilizing the segment with the middle and fourth finger (Figure 2F). When complete, the LMMP will easily peel off the intestinal segment.</li>
	</ol>
	</li>
	<li>Discard the LMMP stripped from the intestinal tube and attached to the cotton swab unless harvesting EGCs from the myenteric plexus is desired.</li>
	<li>Use a razor blade to flay open the intestinal segment longitudinally to remove from the wooden stick.</li>
	<li>Store the stripped intestinal tissue (predominantly mucosa and submucosa plus circular muscle) in DPBS on ice. Repeat steps 4.4&#8211;4.11 until all intestinal segments are prepared.</li>
</ol>
5. Removal of the Epithelial Mucosa
<ol>
	<li>Cut the intestines into smaller pieces of ~0.5 cm with fine scissors.</li>
	<li>Place the tissue in a sterile 50 mL conical centrifuge tube containing 30 mL of ice-cold EDTA/HEPES/DPBS solution.</li>
	<li>Rock the tissue-containing solution at 4 &#176;C for 10 min at ~60 tilts per min.</li>
	<li>Pre-moisten a 5 mL plastic pipette by pipetting the EDTA/HEPES/DPBS buffer up and down one time, and then use the pre-moistened pipette to pipette the tissue and buffer suspension up and down (trituration) 20 times to dislodge the epithelial mucosa.</li>
	<li>Collect the tissue from the EDTA incubation by pouring the mixture through a 100 &#181;m nylon cell strainer. Discard the turbid mucous-filled flow-through (Figure 3).</li>
	<li>Place the tissue retained in the strainer into a new 50 mL conical centrifuge tube containing 30 mL of ice cold EDTA/HEPES/DPBS using needle-nose forceps.</li>
	<li>Repeat the EDTA/HEPES/DPBS incubation a second time by rocking the tissue at 4 &#176;C for 10 min at ~60 tilts per min to strip off additional epithelial mucosa .</li>
	<li>Pre-moisten a 5 mL pipette with the buffer and then pipette the tissue suspension up and down 20 times to dislodge the mucous cells. 
	NOTE: The tissue will tend to stick to inside of the pipette as more of the epithelium is removed.</li>
	<li>Collect the tissue after the second incubation by pouring the mixture through a 100 &#181;m nylon cell strainer and discard the flow through. The second flow-through should be noticeably less turbid than the first (Figure 3).</li>
	<li>Repeat the 10 min EDTA incubations until the solution is nearly clear (about 3 to 4 incubations depending on the amount of tissue and the effectiveness of the trituration) (Figure 3).</li>
</ol>
6. Collection of Enteric Glial Cells from the Lamina Propria and Submucosa
<ol>
	<li>Transfer the tissue from the nylon strainer to a 15 mL conical centrifuge tube containing 5 mL of the commercially available cell recovery solution and rock for 25&#8211;30 min at 4 &#176;C.</li>
	<li>Triturate gently 10x to dissociate the enteric glial cells from the lamina propria.
	<ol>
		<li>Filter through a 40 &#181;m filter and collect the filtrate in a clean 50 mL tube.</li>
		<li>Rinse the tissue on the nylon filter with 1 mL of DPBS.</li>
		<li>Discard the tissue and transfer the ~5&#8211;6 mL of filtrate to a clean 15 mL conical centrifuge tube.</li>
		<li>Spin the filtrate at 2,000 x g in a swinging bucket centrifuge for 5 min at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Re-suspend the glial cell-containing pellet in at least 1 mL of resuspension buffer by gently pipetting the pellet up and down with a 500 &#181;L pipette tip. Avoid introducing bubbles. 
	NOTE: Adjust the resuspension volume as needed for the number of wells and plates. For example, 1.2 mL for one 6-well plate; 2.4 mL for two 6-well plates; 2.4 mL for one 12-well plate.</li>
	<li>Pipette 200 &#181;L of the cell suspension into each well of the 6-well or 100 &#181;L for a 12-well PDL-laminin coated plates containing glial growth media.</li>
	<li>Do not disturb the dishes after adding the cells to the plates and placing in the 37 &#176;C incubator to allow the maximum number of cells to adhere. 
	NOTE: A healthy preparation of cells will attach within 6&#8211;8 h but wait for at least 24 h before changing the media by gently pipetting off the media along with non-adherent cells.</li>
	<li>Use the cells for experiments 3&#8211;4 days after they are placed into culture (Figure 4). Perform immunofluorescent analysis using antibodies of interest to specific protein markers (Table 3). For example, GFAP, S100b and p75NTR or Sox10 was used here. E-cadherin or alpha smooth muscle actin antibodies were used to assess the degree of contamination from other cell types.</li>
</ol>
7. Immunofluorescent Staining
<ol>
	<li>Aspirate the media off the cultures and rinse twice with PBS.</li>
	<li>Fix the cultures for 20 min at room temperature with 4% paraformaldehyde.</li>
	<li>Remove the fixative and then rinse 3 times with PBS.</li>
	<li>Permeabilize the cells with PBS containing 0.2% Triton X-100 at room temperature.</li>
	<li>Incubate the permeabilized cells with blocking solutions comprised of anti-chicken IgY, anti-rabbit or anti-mouse IgG for 2 h at room temperature.</li>
	<li>Incubate the cells with primary antibodies overnight, e.g., GFAP (1:1,000).
	<ol>
		<li>Rinse the cells 3 times with PBS. If performing colocalization proceed to Step 7.7.</li>
	</ol>
	</li>
	<li>Incubate the next set of primary antibodies for at least 2 h at room temperature, but preferably overnight at 4 &#176;C. Use a 1:1000 dilution of rabbit anti-S100 or a 1:500 dilution of mouse anti-p75NTR. 
	NOTE: All antibodies are diluted in PBS.</li>
	<li>Rinse the cells 3 times with PBS to remove primary antibodies. Then incubate the rinsed cells with the appropriate fluorescently-tagged secondary antibodies for 2 h at room temperature (Table 3).</li>
	<li>Rinse 3 times with PBS to remove the secondary antibodies.</li>
	<li>Mount the coverslips onto slides using 1&#8211;2 drops of the mounting media with DAPI and view the cells under a fluorescent microscope.</li>
</ol>
8. Flow Cytometry
<ol>
	<li>Remove adherent glial cells for flow cytometry by rinsing the cells once with DPBS and then add 0.25% trypsin-EDTA per manufacturer&#39;s protocol. Incubate the cells in the trypsin-EDTA solution at 37 &#176;C for 3 min. Terminate the digestion by collecting the cells in complete glial cell growth media.</li>
	<li>Collect the cells by centrifuging at 400 x g for 5 min, and then re-suspend the cells in PBS.</li>
	<li>Fix the cells with 4% paraformaldehyde for 10 min and then permeabilize with 0.2% Triton X-100 in PBS as described in Step 7.4.</li>
	<li>Block the cells with PBS containing 1% BSA and the tissue specific immunoglobulin for 20 min, e.g., donkey anti-chicken IgY (IgG) (H+L) (1:1,000) or donkey anti-goat IgG (H+L) (1:1,000).</li>
	<li>Incubate cells with the primary antibodies GFAP (1:2,000), E-cadherin (1:400), &#945;-smooth muscle actin (1:500), or Pgp9.5 (1:500) overnight at 4 &#176;C.</li>
	<li>Wash away the antibody with PBS.
	<ol>
		<li>Incubate the antigen-antibody complex with fluorescently-tagged secondary antibodies incubated at room temperature for 30 min.</li>
		<li>Use a separate aliquot of cells incubated with secondary antibody to serve as the isotype control. Both populations of cells are washed and resuspended in PBS prior to flow cytometry.</li>
		<li>Analyze the two populations of cells on a flow cytometer by gating on the live cells.</li>
	</ol>
	</li>
</ol>
9. Preparing Mouse Tissues from the hGFAP-Cre:tdTomato Mice
NOTE: The Lox-STOP-Lox-tdTomato cDNA is expressed from the ROSA locus22,23 (Jackson Labs, #007914) and generates endogenous fluorescence when bred to a mouse expressing the Cre recombinase (hGFAP-Cre). In situ analysis is performed by generating frozen tissue sections.
<ol>
	<li>Fix intestinal tissue in 4% paraformaldehyde for 1 h and then dehydrate overnight in 1 M sucrose in PBS.</li>
	<li>Embed the tissue in commercially purchased optimal cutting tissue (OCT) compound comprised of 10% polyvinyl alcohol and 4.2% polyethylene glycol, and then snap freeze in liquid nitrogen.</li>
	<li>Prepare 5 &#181;m cryosections and then mount using an antifade mounting media with DAPI.</li>
	<li>For flow cytometry, prepare EGCs from the hGFAP-Cre:tdTomato+ mice as described in Steps 4.1&#8211;6.5.</li>
	<li>Trypsinize the cells from the plate after 3 days in cultures as in Step 8.1 and then fix for 10 min in 4% paraformaldehyde.</li>
	<li>Analyze cells isolated from Cre Negative mice in parallel with cells isolated from the hGFAP-Cre:tdTomato+ mice on a flow cytometer. 
	NOTE: Gates were constructed to avoid dead cells and debris.</li>
</ol>
10. Ca2+ Flux Imaging
<ol>
	<li>Incubate EGCs plated on a 24-well plate with 3 &#181;M Fluo-4-AM at 37 &#176;C for 30 min.</li>
	<li>Image the Ca2+ signal in commercially purchased Tyrode&#39;s solution using a spinning disk confocal microscope and an excitation wavelength of 480 nm (F480) after adding CCK or gastrin peptide. Image analysis was reported in Sundaresan et al.10 .</li>
</ol>

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The authors have nothing to disclose.

EGCs play important roles in gut homeostasis, and it is essential to isolate and study them in vitro. In this protocol, a simple method for isolating EGCs from the lamina propria of the adult mouse intestine was introduced to study enteric glial function.
Removing the adherent mesentery and LMMP with a cotton swab removes some of the inter-myenteric glia residing between the longitudinal and circular muscle, increases the accessibility of buffers to the submucosal surface and removes ...

Interest in the function of enteric glial cells (EGCs) has steadily increased due to their recognized roles in the gut integrity and homeostasis1,2. In addition, EGCs vary according to their location along the length of the GI tract3,4. EGCs release various trophic factors including glial cell-derived neurotrophic factor (GDNF), contribute to gut motility1,5 and respond to microbial byproducts6,7. Studies have indicated that the EGC population is heterogeneous and that their function varies depending on whether they are submucosal or reside within the myenteric plexus1,7. For example, EGCs within the submucosa contribute to tight junctions8. Differential GFAP expression and phosphorylation in EGCs have been linked to Parkinson&#39;s Disease, suggesting their possible link to the gut phenotype of this disorder9. Recently, it was observed that the loss of the nuclear protein menin in isolated cultures of EGCs from the proximal intestinal submucosa was sufficient to induce expression of the hormone gastrin10. As a result, it was proposed that EGCs might be the origin of duodenal gastrinomas, a type of neuroendocrine tumor10. Collectively, these examples underscore the relevance of studying the behavior and function of isolated EGCs in neuropathic disorders and cancer11.
The challenge in the field remains how to isolate and study either or both EGC populations in vitro. Lineage trace experiments demonstrated that EGCs in the submucosa and lamina propria originate from progenitor cells in the myenteric plexus7. Although there are several published isolation protocols available to generate cultures of myenteric EGCs12,13,14,15,16,17,18,19, none specifically targets isolation of the submucosal/lamina propria EGC population. Existing protocols for EGC isolation specifically use a combination of the mechanical separation or the microdissection of the smooth muscle combined with enzymatic dissociation, eventually discarding the mucosal cell layer.
The goal of this manuscript is to demonstrate the steps to non-enzymatically isolate primary EGCs from the lamina propria for in vitro studies. Since there are no markers that specifically distinguish myenteric EGCs from those in the submucosa, the spatial separation of the epithelial mucosa from the smooth muscle was exploited to isolate submucosal EGCs. In addition, by combining EDTA chelation with non-enzymatic dissociation, EGCs were isolated from the submucosa in contrast to the smooth muscle, which was discarded along with the associated inter-myenteric EGCs. Further separation of the submucosa and lamina propria EGCs occurred by culturing the cells on glial cell-friendly substrates, e.g., poly-D-lysine and laminin.

<table>
	<tbody>
		<tr>
			<td>Poly-D lysine (1 mg/ml stock)</td>
			<td>Sigma</td>
			<td>A-003-E</td>
			<td>Dilute 1:10</td>
		</tr>
		<tr>
			<td>Laminin (0.5 mg/ml stock)</td>
			<td>Sigma</td>
			<td>L4544</td>
			<td>Dilute to 10 &#181;g/mL on ICE</td>
		</tr>
		<tr>
			<td>EDTA (0.5M)</td>
			<td>Lonza</td>
			<td>51201</td>
			<td>Dilute 1:100 in DPBS</td>
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			<td>HEPES (1 M)</td>
			<td>Corning</td>
			<td>36216004</td>
			<td>Dilute 1:100 in DPBS</td>
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		<tr>
			<td>Cell Recovery Solution</td>
			<td>Corning</td>
			<td>354253</td>
			<td></td>
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			<td>Dulbecco&#39;s Phosphate Buffered Saline (DPBS)</td>
			<td>HyClone</td>
			<td>SH30028.02</td>
			<td></td>
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		<tr>
			<td>DMEM/F-12</td>
			<td>Thermo Fisher Scientific</td>
			<td>11320033</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (100X)</td>
			<td>Life Technologies</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>Gentamicin (50mg/mL stock)</td>
			<td>Life Technologies</td>
			<td>15750060</td>
			<td></td>
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			<td>GDNF (10 &#181;g stock)</td>
			<td>Sigma</td>
			<td>SRP3200</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Glutamine (200 mM stock)</td>
			<td>Life Technologies</td>
			<td>25030-081</td>
			<td></td>
		</tr>
		<tr>
			<td>Chicken anti-GFAP</td>
			<td>Thermo Fisher Scientific</td>
			<td>PA1-10004</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-a-Smooth Muscle Actin&#160;</td>
			<td>Abcam</td>
			<td>ab112022</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti-Pgp9.5&#160;</td>
			<td>Novus Biologicals</td>
			<td>NB600-1160</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-E-cadherin&#160;</td>
			<td>R&#38;D Systems</td>
			<td>AF748</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit S100&#160;</td>
			<td>Abcam</td>
			<td>ab34686</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse p75 NTR&#160;</td>
			<td>Millipore</td>
			<td>MAB5592</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 Goat Anti-Chicken IgY</td>
			<td>Invitrogen</td>
			<td>A-11039</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 Donkey Anti-Goat IgG</td>
			<td>Invitrogen</td>
			<td>A-11055</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 568 Goat Anti-Mouse IgG</td>
			<td>Invitrogen</td>
			<td>A-11004</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 594 Donkey Anti-Rabbit IgG</td>
			<td>Invitrogen</td>
			<td>R-37119</td>
			<td></td>
		</tr>
		<tr>
			<td>Prolong Gold antifade Reagent with DAPI</td>
			<td>Thermo Fisher Scientific</td>
			<td>P36931</td>
			<td></td>
		</tr>
		<tr>
			<td>Fungizone (Amphotericin B) 250 &#181;g/ml</td>
			<td>Life Technologies</td>
			<td>15290-018</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Fura-2-AM</td>
			<td>Invitrogen</td>
			<td>F-14201</td>
			<td></td>
		</tr>
		<tr>
			<td>CCK peptide</td>
			<td>Anaspec, Fremont, CA</td>
			<td>AS-20741</td>
			<td></td>
		</tr>
		<tr>
			<td>Gastrin peptide (Gastrin-17)</td>
			<td>Abbiotec, Bloomington, IN</td>
			<td>350188</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylon Mesh Celll Strainer (100 &#181;m)</td>
			<td>Fisher Scientific</td>
			<td>22363549</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylon Mesh Celll Strainer (40 &#181;m)</td>
			<td>Fisher Scientific</td>
			<td>22363547</td>
			<td></td>
		</tr>
		<tr>
			<td>Disposable Serologic Pipet 5 ml</td>
			<td>Fisher Scientific</td>
			<td>13-678-11D</td>
			<td></td>
		</tr>
		<tr>
			<td>0.25% Trypsin-EDTA (1X)</td>
			<td>Life Technologies</td>
			<td>25200-056</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolation of enteric glial cells from the submucosa and lamina propria of the adult mouse

The enteric nervous system (ENS) consists of neurons and enteric glial cells (EGCs) that reside within the smooth muscle wall, submucosa and lamina propria. EGCs play important roles in gut homeostasis through the release of various trophic factors and contribute to the integrity of the epithelial barrier. Most studies of primary enteric glial cultures use cells isolated from the myenteric plexus after enzymatic dissociation. Here, a non-enzymatic method to isolate and culture EGCs from the intestinal submucosa and lamina propria is described. After manual removal of the longitudinal muscle layer, EGCs were liberated from the lamina propria and submucosa using sequential HEPES-buffered EDTA incubations followed by incubation in commercially available non-enzymatic cell recovery solution. The EDTA incubations were sufficient to strip most of the epithelial mucosa from the lamina propria, allowing the cell recovery solution to liberate the submucosal EGCs. Any residual lamina propria and smooth muscle was discarded along with the myenteric glia. EGCs were easily identified by their ability to express glial fibrillary acidic protein (GFAP). Only about 50% of the cell suspension contained GFAP+ cells after completing tissue incubations and prior to plating on the poly-D-lysine/laminin substrate. However, after 3 days of culturing the cells in glial cell-derived neurotrophic factor (GDNF)-containing culture media, the cell population adhering to the substrate-coated plates comprised of &#62;95% enteric glia. We created a hybrid mouse line by breeding a hGFAP-Cre mouse to the ROSA-tdTomato reporter line to track the percentage of GFAP+ cells using endogenous cell fluorescence. Thus, non-myenteric enteric glia can be isolated by non-enzymatic methods and cultured for at least 5 days.

Preps were considered unsuccessful if GFAP+ cells did not adhere and spread within 24 h (Figure 4A). The number of glial cells could not be determined until after 24 h when the cells adhered and showed evidence of spreading into flat aggregates (Figure 4B). Cells at the edge of the clusters tended to extend long processes and expressed classic glial markers, e.g., GFAP, S100b and p75NTR (F...

Watch this Scientific Journal Video about Isolation of Enteric Glial Cells from the Submucosa and Lamina Propria of the Adult Mouse at JoVE.com

Isolation of Enteric Glial Cells from the Submucosa and Lamina Propria of the Adult Mouse

Here, we describe the isolation of enteric-glial cells from the intestinal-submucosa using sequential EDTA incubations to chelate divalent cations and then incubation in non-enzymatic cell recovery solution. Plating the resultant cell suspension on poly-D-lysine and laminin results in a highly enriched culture of submucosal glial cells for functional analysis.

The enteric nervous system (ENS) consists of neurons and enteric glial cells (EGCs) that reside within the smooth muscle wall, submucosa and lamina ...

isolation-enteric-glial-cells-from-submucosa-lamina-propria-adult

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12.9K Views.  University of Michigan. Here, we describe the isolation of enteric-glial cells from the intestinal-submucosa using sequential EDTA incubations to chelate divalent cations and then incubation in non-enzymatic cell recovery solution. Plating the resultant cell suspension on poly-D-lysine and laminin results in a highly enriched culture of submucosal glial cells for functional analysis.

Video: Isolation of Enteric Glial Cells from the Submucosa and Lamina Propria of the Adult Mouse

Isolation of Retinal Stem Cells from the Mouse Eye

The adult mouse retinal stem cell (RSC) is a rare quiescent cell found within the ciliary epithelium (CE) of the mammalian eye1,2,3. The CE is made up of non-pigmented inner and pigmented outer cell layers, and the clonal RSC colonies that arise from a single pigmented cell from the CE are made up of both pigmented and non-pigmented cells which can be differentiated to form all the cell types of the neural retina and the RPE. There is some controversy about whether all the cells within the spheres all contain at least some pigment4; however the cells are still capable of forming the different cell types found within the neural retina1-3. In some species, such as amphibians and fish, their eyes are capable of regeneration after injury5, however; the mammalian eye shows no such regenerative properties. We seek to identify the stem cell in vivo and to understand the mechanisms that keep the mammalian retinal stem cells quiescent6-8, even after injury as well as using them as a potential source of cells to help repair physical or genetic models of eye injury through transplantation9-12. Here we describe how to isolate the ciliary epithelial cells from the mouse eye and grow them in culture in order to form the clonal retinal stem cell spheres. Since there are no known markers of the stem cell in vivo, these spheres are the only known way to prospectively identify the stem cell population within the ciliary epithelium of the eye.

In this video, we will demonstrate how to isolate retinal stem cells from the ciliary epithelium of the mouse eye and grow them in culture to form clonal retinal spheres. The spheres that are isolated possess the cardinal properties of stem cells: self-renewal and multipotentiality.

The adult mouse retinal stem cell (RSC) is a rare quiescent cell found within the ciliary epithelium (CE) of the mammalian eye1,2,3. The CE is made up of ...

Isolation and Culture of Cells from the Nephrogenic Zone of the Embryonic Mouse Kidney

Embryonic development of the kidney has been extensively studied both as a model for epithelial-mesenchymal interaction in organogenesis and to gain understanding of the origins of congenital kidney disease. More recently, the possibility of steering na&iuml;ve embryonic stem cells toward nephrogenic fates has been explored in the emerging field of regenerative medicine. Genetic studies in the mouse have identified several pathways required for kidney development, and a global catalog of gene transcription in the organ has recently been generated <a href="http://www.gudmap.org/">http://www.gudmap.org/</a>, providing numerous candidate regulators of essential developmental functions. Organogenesis of the rodent kidney can be studied in organ culture, and many reports have used this approach to analyze outcomes of either applying candidate proteins or knocking down the expression of candidate genes using siRNA or morpholinos. However, the applicability of organ culture to the study of signaling that regulates stem/progenitor cell differentiation versus renewal in the developing kidney is limited as cultured organs contain a compact extracellular matrix limiting diffusion of macromolecules and virus particles. To study the cell signaling events that influence the stem/progenitor cell niche in the kidney we have developed a primary cell system that establishes the nephrogenic zone or progenitor cell niche of the developing kidney ex vivo in isolation from the epithelial inducer of differentiation. Using limited enzymatic digestion, nephrogenic zone cells can be selectively liberated from developing kidneys at E17.5. Following filtration, these cells can be cultured as an irregular monolayer using optimized conditions. Marker gene analysis demonstrates that these cultures contain a distribution of cell types characteristic of the nephrogenic zone in vivo, and that they maintain appropriate marker gene expression during the culture period. These cells are highly accessible to small molecule and recombinant protein treatment, and importantly also to viral transduction, which greatly facilitates the study of candidate stem/progenitor cell regulator effects. Basic cell biological parameters such as proliferation and cell death as well as changes in expression of molecular markers characteristic of nephron stem/progenitor cells in vivo can be successfully used as experimental outcomes. Ongoing work in our laboratory using this novel primary cell technique aims to uncover basic mechanisms governing the regulation of self-renewal versus differentiation in nephron stem/progenitor cells.

In this report we describe a method for the isolation and culture of the progenitor cell niche from the embryonic mouse kidney that can be used to study signaling pathways regulating stem/progenitor cells of the developing kidney. These cultured cells are highly accessible to small molecule and recombinant protein treatment, and importantly also to viral transduction, which allows efficient manipulation of candidate pathways.

Embryonic development of the kidney has been extensively studied both as a model for epithelial-mesenchymal interaction in organogenesis and  to gain ...

Generation of Mice Derived from Induced Pluripotent Stem Cells

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also produce a source of patient-matched cells for regenerative therapies. iPSCs may be generated using multiple protocols and derived from multiple cell sources. Once generated, iPSCs are tested using a variety of assays including immunostaining for pluripotency markers, generation of three germ layers in embryoid bodies and teratomas, comparisons of gene expression with embryonic stem cells (ESCs) and production of chimeric mice with or without germline contribution2. Importantly, iPSC lines that pass these tests still vary in their capacity to produce different differentiated cell types2. This has made it difficult to establish which iPSC derivation protocols, donor cell sources or selection methods are most useful for different applications.
The most stringent test of whether a stem cell line has sufficient developmental potential to generate all tissues required for survival of an organism (termed full pluripotency) is tetraploid embryo complementation (TEC)3-5. Technically, TEC involves electrofusion of two-cell embryos to generate tetraploid (4n) one-cell embryos that can be cultured in vitro to the blastocyst stage6. Diploid (2n) pluripotent stem cells (e.g. ESCs or iPSCs) are then injected into the blastocoel cavity of the tetraploid blastocyst and transferred to a recipient female for gestation (see Figure 1). The tetraploid component of the complemented embryo contributes almost exclusively to the extraembryonic tissues (placenta, yolk sac), whereas the diploid cells constitute the embryo proper, resulting in a fetus derived entirely from the injected stem cell line.
Recently, we reported the derivation of iPSC lines that reproducibly generate adult mice via TEC1. These iPSC lines give rise to viable pups with efficiencies of 5-13%, which is comparable to ESCs3,4,7 and higher than that reported for most other iPSC lines8-12. These reports show that direct reprogramming can produce fully pluripotent iPSCs that match ESCs in their developmental potential and efficiency of generating pups in TEC tests. At present, it is not clear what distinguishes between fully pluripotent iPSCs and less potent lines13-15. Nor is it clear which reprogramming methods will produce these lines with the highest efficiency. Here we describe one method that produces fully pluripotent iPSCs and "all- iPSC" mice, which may be helpful for investigators wishing to compare the pluripotency of iPSC lines or establish the equivalence of different reprogramming methods.

Generating induced pluripotent stem cell (iPSC) lines produces lines of differing developmental potential even when they pass standard tests for pluripotency. Here we describe a protocol to produce mice derived entirely from iPSCs, which defines the iPSC lines as possessing full pluripotency1.

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also ...

Isolation and Culture of Dental Epithelial Stem Cells from the Adult Mouse Incisor

Understanding the cellular and molecular mechanisms that underlie tooth regeneration and renewal has become a topic of great interest1-4, and the mouse incisor provides a model for these processes. This remarkable organ grows continuously throughout the animal&#39;s life and generates all the necessary cell types from active pools of adult stem cells housed in the labial (toward the lip) and lingual (toward the tongue) cervical loop (CL) regions. Only the dental stem cells from the labial CL give rise to ameloblasts that generate enamel, the outer covering of teeth, on the labial surface. This asymmetric enamel formation allows abrasion at the incisor tip, and progenitors and stem cells in the proximal incisor ensure that the dental tissues are constantly replenished. The ability to isolate and grow these progenitor or stem cells in vitro allows their expansion and opens doors to numerous experiments not achievable in vivo, such as high throughput testing of potential stem cell regulatory factors. Here, we describe and demonstrate a reliable and consistent method to culture cells from the labial CL of the mouse incisor.

The continuously growing mouse incisor provides a model for studying renewal of dental tissues from dental epithelial stem cells (DESCs). A robust system for consistently and reliably obtaining these cells from the incisor and expanding them in vitro is reported here.

Understanding the cellular and molecular mechanisms that underlie tooth regeneration and renewal has become a topic of great interest1-4, and the mouse ...

Isolation of Murine Valve Endothelial Cells

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and surrounded by a monolayer of valve endothelial cells (VECs). VECs play essential roles in establishing the valve structures during embryonic development, and are important for maintaining life-long valve integrity and function. In contrast to a continuous endothelium over the surface of healthy valve leaflets, VEC disruption is commonly observed in malfunctioning valves and is associated with pathological processes that promote valve disease and dysfunction. Despite the clinical relevance, focused studies determining the contribution of VECs to development and disease processes are limited. The isolation of VECs from animal models would allow for cell-specific experimentation. VECs have been isolated from large animal adult models but due to their small population size, fragileness, and lack of specific markers, no reports of VEC isolations in embryos or adult small animal models have been reported. Here we describe a novel method that allows for the direct isolation of VECs from mice at embryonic and adult stages. Utilizing the Tie2-GFP reporter model that labels all endothelial cells with Green Fluorescent Protein (GFP), we have been successful in isolating GFP-positive (and negative) cells from the semilunar and atrioventricular valve regions using fluorescence activated cell sorting (FACS). Isolated GFP-positive VECs are enriched for endothelial markers, including CD31 and von Willebrand Factor (vWF), and retain endothelial cell expression when cultured; while, GFP-negative cells exhibit molecular profiles and cell shapes consistent with VIC phenotypes. The ability to isolate embryonic and adult murine VECs allows for previously unattainable molecular and functional studies to be carried out on a specific valve cell population, which will greatly improve our understanding of valve development and disease mechanisms.

The ability to isolate heart valve endothelial cells (VECs) is critical for understanding mechanisms of valve development, maintenance, and disease. Here we describe the isolation of VECs from embryonic and adult Tie2-GFP mice using FACS that will allow for studies determining the contribution of VECs in developmental and disease processes.

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and ...

Isolation and Primary Culture of Mouse Aortic Endothelial Cells

The vascular endothelium is essential to normal vascular homeostasis. Its dysfunction participates in various cardiovascular disorders. The mouse is an important model for cardiovascular disease research. This study demonstrates a simple method to isolate and culture endothelial cells from the mouse aorta without any special equipment. To isolate endothelial cells, the thoracic aorta is quickly removed from the mouse body, and the attached adipose tissue and connective tissue are removed from the aorta. The aorta is cut into 1 mm rings. Each aortic ring is opened and seeded onto a growth factor reduced matrix with the endothelium facing down. The segments are cultured in endothelial cell growth medium for about 4 days. The endothelial sprouting starts as early as day 2. The segments are then removed and the cells are cultured continually until they reach confluence. The endothelial cells are harvested using neutral proteinase and cultured in endothelial cell growth medium for another two passages before being used for experiments. Immunofluorescence staining indicated that after the second passage the majority of cells were double positive for Dil-ac-LDL uptake, Lectin binding, and CD31 staining, the typical characteristics of endothelial cells. It is suggested that cells at the second to third passages are suitable for in vitro and in vivo experiments to study the endothelial biology. Our protocol provides an effective means of identifying specific cellular and molecular mechanisms in endothelial cell physiopathology.

The vascular endothelial cells play a significant role in many important cardiovascular disorders. This article describes a simple method to isolate and expand endothelial cells from the mouse aorta without using any special equipment. Our protocol provides an effective means of identifying mechanisms in endothelial cell physiopathology.

The vascular endothelium is essential to normal vascular homeostasis. Its dysfunction participates in various cardiovascular disorders. The mouse is an ...

Isolation and Culture of Primary Mouse Keratinocytes from Neonatal and Adult Mouse Skin

The keratinocyte (KC) is the predominant cell type in the epidermis, the outermost layer of the skin. Epidermal KCs play a critical role in providing skin defense by forming an intact skin barrier against environmental insults, such as UVB irradiation or pathogens, and also by initiating an inflammatory response upon those insults. Here we describe methods to isolate KCs from neonatal mouse skin and from adult mouse tail skin. We also describe culturing conditions using defined growth supplements (dGS) in comparison to chelexed fetal bovine serum (cFBS). Functionally, we show that both neonatal and adult KCs are highly responsive to high calcium-induced terminal differentiation, tight junction formation and stratification. Additionally, cultured adult KCs are susceptible to UVB-triggered cell death and can release large amounts of TNF upon UVB irradiation. Together, the methods described here will be useful to researchers for the setup of in vitro models to study epidermal biology in the neonatal mouse and/or the adult mouse.

Epidermal keratinocytes form a functional skin barrier and are positioned at the front line of host defense against external environmental insults. Here we describe methods for isolation and primary culture of epidermal keratinocytes from neonatal and adult mouse skin, and induction of terminal differentiation and UVB-triggered inflammatory response from keratinocytes.

The keratinocyte (KC) is the predominant cell type in the epidermis, the outermost layer of the skin. Epidermal KCs play a critical role in providing skin ...

Isolation, Characterization, and Differentiation of Cardiac Stem Cells from the Adult Mouse Heart

Myocardial infarction (MI) is a leading cause of morbidity and mortality around the world. A major goal of regenerative medicine is to replenish the dead myocardium after MI. Although several strategies have been used to regenerate myocardium, stem cell therapy remains a major approach to replenish the dead myocardium of an MI heart. Accumulating evidence suggests the presence of resident cardiac stem cells (CSCs) in the adult heart and their endocrine and/or paracrine effects on cardiac regeneration. However, CSC isolation and their characterization and differentiation toward myocardial cells, especially cardiomyocytes, remains a technical challenge. In the present study, we provided a simple method for the isolation, characterization, and differentiation of CSCs from the adult mouse heart. Here, we describe a density gradient method for the isolation of CSCs, where the heart is digested by a 0.2% collagenase II solution. To characterize the isolated CSCs, we evaluated the expression of CSCs/cardiac markers Sca-1, NKX2-5, and GATA4, and pluripotency/stemness markers OCT4, SOX2, and Nanog. We also determined the proliferation potential of isolated CSCs by culturing them in a Petri dish and assessing the expression of the proliferation marker Ki-67. For evaluating the differentiation potential of CSCs, we selected seven- to ten-days cultured CSCs. We transferred them to a new plate with a cardiomyocyte differentiation medium. They are incubated in a cell culture incubator for 12 days, while the differentiation medium is changed every three days. The differentiated CSCs express cardiomyocyte-specific markers: actinin and troponin I. Thus, CSCs isolated with this protocol have stemness and cardiac markers, and they have a potential for proliferation and differentiation toward cardiomyocyte lineage.

The overall goal of this article is to standardize the protocol for the isolation, characterization, and differentiation of cardiac stem cells (CSCs) from the adult mouse heart. Here, we describe a density gradient centrifugation method to isolate murine CSCs and elaborated methods for CSC culture, proliferation, and differentiation into cardiomyocytes.

Myocardial infarction (MI) is a leading cause of morbidity and mortality around the world. A major goal of regenerative medicine is to replenish the dead ...

Isolation of Myoepithelial Cells from Adult Murine Lacrimal and Submandibular Glands

The lacrimal gland (LG) is an exocrine tubuloacinar gland that secretes an aqueous layer of tear film. The LG epithelial tree is comprised of acinar, ductal epithelial, and myoepithelial cells (MECs). MECs express alpha smooth muscle actin (&#945;SMA) and have a contractile function. They are found in multiple glandular organs and are of ectodermal origin. In addition, the LG contains SMA+ vascular smooth muscle cells of endodermal origin called pericytes: contractile cells that envelop the surface of vascular tubes. A new protocol allows us to isolate both MECs and pericytes from adult murine LGs and submandibular glands (SMGs). The protocol is based on the genetic labeling of MECs and pericytes using the SMACreErt2/+:Rosa26-TdTomatofl/fl mouse strain, followed by preparation of the LG single-cell suspension for fluorescence activated cell sorting (FACS). The protocol allows for the separation of these two cell populations of different origins based on the expression of the epithelial cell adhesion molecule (EpCAM) by MECs, whereas pericytes do not express EpCAM. Isolated cells could be used for cell cultivation or gene expression analysis.

The lacrimal gland (LG) has two cell types expressing &#945;-smooth muscle actin (&#945;SMA): myoepithelial cells (MECs) and pericytes. MECs are of ectodermal origin, found in many glandular tissues, while pericytes are vascular smooth muscle cells of endodermal origin. This protocol isolates MECs and pericytes from murine LGs.

The lacrimal gland (LG) is an exocrine tubuloacinar gland that secretes an aqueous layer of tear film. The LG epithelial tree is comprised of acinar, ...

Simultaneous Isolation and Culture of Atrial Myocytes, Ventricular Myocytes, and Non-Myocytes from an Adult Mouse Heart

The isolation and culturing of cardiac myocytes from mice has been essential for furthering the understanding of cardiac physiology and pathophysiology. While isolating myocytes from neonatal mouse hearts is relatively straightforward, myocytes from the adult murine heart are preferred. This is because compared to neonatal cells, adult myocytes more accurately recapitulate cell function as it occurs in the adult heart in vivo. However, it is technically difficult to isolate adult mouse cardiac myocytes in the necessary quantities and viability, which contributes to an experimental impasse. Furthermore, published procedures are specific for the isolation of either atrial or ventricular myocytes at the expense of atrial and ventricular non-myocyte cells. Described here is a detailed method for isolating both atrial and ventricular cardiac myocytes, along with atrial and ventricular non-myocytes, simultaneously from a single mouse heart. Also provided are the details for optimal cell-specific culturing methods, which enhance cell viability and function. This protocol aims not only to expedite the process of adult murine cardiac cell isolation, but also to increase the yield and viability of cells for investigations of atrial and ventricular cardiac cells.

A method is described for the simultaneous isolation of myocytes and non-myocytes from both the atria and ventricles of a single adult mouse heart. This protocol results in consistent yields of highly viable cardiac myocytes and non-myocytes and details optimal cell-specific culture conditions for phenotyping and in vitro analysis.

The isolation and culturing of cardiac myocytes from mice has been essential for furthering the understanding of cardiac physiology and pathophysiology. ...