This work was supported by a) the National Institutes of Health/NIAMS (grant numbers R01 AR062507 and R01 AR053860 to RS), b) the University of Alabama at Birmingham Dental Academic Research Training (DART) grant (number T90DE022736 (PI MacDougall)) to SBP from the National Institute of Dental and Craniofacial Research/National Institutes of Health, c) a UAB Global Center for Craniofacial, Oral and Dental Disorders (GC-CODED) Pilot and Feasibility grant to SBP and d) the National Institute of Dental and Craniofacial Research/National Institutes of Health K99 DE024406 grant to SBP.

All experiments with mice were approved by the UAB Institutional Animal Care and Use Committee (IACUC).
1. Plate Preparation
NOTE: Coverslips can be used to image DP cells at the end of the assay. Be sure the plate lid is on during all incubation and rinsing steps outside of the sterile tissue culture hood to prevent contamination during sample processing.
<ol>
	<li>Coverslip preparation
	<ol>
		<li>Autoclave circular coverslips.</li>
		<li>Filter ultrapure water under the hood.</li>
		<li>Transfer the coverslips to a 24-well plate and cover each one with 400 &#956;L of 0.1 mg/mL poly-D-lysine.</li>
		<li>Allow the coverslips to soak, immersed for 5 min, on a rocker at 12 rpm or orbital shaker at 40-50 rpm. Be sure lid is on to prevent contamination.</li>
		<li>Rinse the coverslips with filtered ultrapure water for about 5 min on a rocker at 12 rpm or shaker at 50 rpm. Repeat.</li>
		<li>Allow the coverslips to dry for at least 2 h under the hood with the plate lid off to facilitate evaporation. These coverslips should be used within 48 h.</li>
		<li>Seed DP cells atop coverslips and process at the end of the assay for immunofluorescence (section 3.3).</li>
	</ol>
	</li>
	<li>Coating filters
	<ol>
		<li>Dilute laminin to 10 &#956;g/mL. Filter the diluted laminin under the hood.</li>
		<li>Pipette 450-500 &#956;L of 10 &#956;g/mL laminin into each well of a 24-well plate.</li>
		<li>Place transwell filter, 3 &#956;m porosity, in a well so that it contacts the laminin solution and leave it in a 37 &#176;C incubator for either 2 h or overnight. Filter pores should allow some of the solution to diffuse and coat both the top and bottom of the filter. These filters should be used within 48 h or be refrigerated at 4 &#176;C for up to 1 week.</li>
	</ol>
	</li>
</ol>
2. Cell Plating with Optional Genetic Manipulation
<ol>
	<li>Mice: Genetic alteration of DP cells can be performed (but is not required) to study mesenchymal-neuronal interactions. See sections 2.3 and 2.4 for suggested genetic variations.</li>
	<li>DP dissection, dispersion and plating (Figure 1) 
	NOTE: For DP dissection, use ultra-fine, straight-edge forceps. The ultra-fine edges will allow the user to wedge the forcep edge between the mineralized structure and DP tissue.
	<ol>
		<li>Harvest P5-P8 mice. At this stage, the teeth should be mineralizing, and the root is open.</li>
		<li>Anesthetize the neonates via hypothermia by placing them in a dish in the 4 &#176;C refrigerator until they no longer move or respond to touch. Euthanize neonates by decapitation and in accordance with IACUC procedures at the designated facility.</li>
		<li>Prepare a 3-5 mL aliquot of 0.25% trypsin-EDTA in a 50 mL conical tube to collect the DP from each mouse. This will facilitate the digestion of dental pulp from postnatal mice. Use more than 3 mL if digesting tissue from more than 10 postnatal mice.</li>
		<li>Place the head on a disposable underpad so that the mouth is toward the ceiling and the base of the neck is flat on the work surface. Use a razor blade in a sawing motion to separate the mandible from the maxilla.</li>
		<li>Optionally remove the tongue either with scissors or with forceps to allow easier access to the molars.</li>
		<li>Place the opened head in a dish atop a sterile gauze pad and place the specimen under a dissecting microscope (Figure 1D).</li>
		<li>Remove the alveolar bone tissue surrounding first molars. Submandibular teeth are not fully erupted at this point. Insert forceps into alveolar opening and tease the tissue away from the tooth toward the buccal (cheek) or lingual (tongue) side of the mouth. Maxillary teeth will require full removal of the cleft around the tooth for exposure and removal.</li>
		<li>Gently transfer the submandibular and maxillary first molars (M1s) to a separate cell culture dish with 1x phosphate-buffered saline (PBS).</li>
		<li>Repeat 2.2.4-2.2.8 until all M1s are collected. Keep the dish containing the M1s on ice during the harvesting.</li>
		<li>Remove the Enamel Outer Organ (EOE) surrounding the outside of each M1. This can alternatively be done after step 2.2.11.</li>
		<li>With a set of forceps, rotate the M1 so the cusps are down and the open root is exposed. There will be an oval opening on the bottom of the tooth, and opaque DP tissue encapsulated by a thin layer of dentin and enamel.</li>
		<li>Using the tip of the forceps, gently loosen the DP by running one arm of the forceps around the internal circumference of the mineralized tissue. Remove DP tissue out of the mineralized structure and transfer it to a third dish containing 1x PBS. Remove the EOE if it was not already separated (Figure 1E).</li>
		<li>Transfer all DP tissue to 0.25% trypsin-EDTA in a 50 mL conical tube. Vortex the mixture and place in a 37 &#176;C warm water bath for 10 min. This can be done with the same forceps or with long, vial forceps. The tissue will be difficult to disperse and will require vortexing every 3-4 min. Do NOT exceed 10 min trypsinization since the trypsin can damage cell membranes.</li>
		<li>Under a sterile hood, add warmed co-culture media (Table 1) to a final ratio of at least 1:1 media to trypsin to inactivate the enzyme. Larger ratios are acceptable if more tissue dispersion is desired.</li>
		<li>Pipette the media up and down multiple times with a 10 mL pipette to further disperse the DP in the media. Be careful to avoid large bubbles. Complete dispersion is nearly impossible due to the sticky nature of the tissue. However, it is also not necessary since cells will migrate outward from the tissue once plated.</li>
		<li>Transfer 1 mL of the dispersed DP to each well of a 24-well tissue culture plate (Figure 1F).</li>
		<li>Place the plate in an incubator at 37 &#176;C and allow cells to attach and migrate out from the undispersed tissue for 48 h before changing media. Primary cells need to be plated at relatively high concentrations in order to reach 85-90% confluence within 1 week. If this is not be achieved after 1 week, discard the plate.</li>
	</ol>
	</li>
	<li>Optional Genetic Manipulation of DP Cells
	<ol>
		<li>To alter cell signaling pathways, harvest DP cells from genetic knockout mice or from mice in which a gene of interest is flanked by loxP sites. In the latter case, the gene can be deleted using Adenovirus-Cre-GFP (Ad-Cre-GFP) recombinase to remove the flanked gene, as described below. Use Adenovirus-eGFP (Ad-eGFP) as a control virus to ensure that the viral infection does not cause a cellular response. 
		NOTE: Ad-eGFP is controlled by the CMV promoter enhancer, which is very strong. Ad-Cre-GFP is regulated by an IRES, an internal regulatory region between the Cre and GFP, which is not very strong. This results in brighter fluorescence in Ad-eGFP cells than Ad-Cre-GFP cells. Confirm equivalent infection levels based on the total numbers of cells fluorescing, not on the levels of cellular fluorescence.</li>
		<li>Prepare 500 &#956;L of media containing virus and 10 &#956;g/mL of polybrene per well. Polybrene assists with viral infection in cells near confluence10. This protocol is based on a multiplicity of infection (MOI) of 100 for Ad-eGFP and 200 Ad-Cre-GFP for efficient gene infection with little or no effect on cell viability. The estimated cell number is 4 x 104 cells/well of a 24-well plate: 
		<img alt="Equation 1" src="/files/ftp_upload/60809/60809equ01.jpg" /></li>
		<li>Mix media with brief vortexing or pipetting and add 500 &#956;L to each well.</li>
		<li>After 24 h, add an additional 500 &#956;L of co-culture media that does not contain additional polybrene or virus.</li>
		<li>After a total of 48 h, aspirate virus-containing media and replace with fresh co-culture media. At this point, TG neurons can be added atop transwell filters.</li>
	</ol>
	</li>
	<li>Trigeminal neuron dissection, dispersion and plating 
	NOTE: In this protocol, the imaging of neurite outgrowth in co-culture with DP cells was optimized using adolescent (6-week-old) B6.Cg-Tg(Thy1-YFP)16Jrs/J mice. Central and peripheral nervous systems of Thy1-YFP mice have a yellow fluorescent protein (YFP) tag whose expression begins around P6-P10 in neurons and increases exponentially throughout the nervous system during postnatal and adult life11,12. YFP and GFP have conserved sequences that allow these nerves to be stained with anti-GFP antibodies, resulting in a pan-neuronal stain. Ultimately, these mice allow for better visualization and quantification of the neurons used and grown in cell culture.
	<ol>
		<li>Euthanize adolescent mice with carbon dioxide followed by cervical dislocation.</li>
		<li>Decapitate the mice and remove the skin from the skull. Be sure to include equivalent numbers of males and females.</li>
		<li>Insert the tip of a pair of micro-dissecting scissors into the base of the skull. Cut along the sagittal suture of the skull (Figure 1A).</li>
		<li>Make four small horizontal cuts: two along the coronal sutures by the ears, and two along the lambdoid sutures at the base of the skull. This should create two flaps of bone.</li>
		<li>Use the forceps to peel back the two flaps of bone. This should reveal the brain.</li>
		<li>Remove the brain. Transfer the head to a tissue culture dish with 1x PBS and put under the microscope.</li>
		<li>Locate the TG ganglia, which is easily visible in rodents13, housed in the dura matter between the brain and bone of the maxillary process (Figure 1B).</li>
		<li>Cut the three branches that travel to the eyes, maxillae and mandible and transfer the ganglia to cold 1x PBS using straight-edge fine forceps. Keep the dish containing the TG ganglia on ice during harvesting.</li>
		<li>Once all TG bundles are harvested, transfer ganglia to a 50 mL conical tube containing 5 mg/mL sterile-filtered collagenase type II using vial forceps.</li>
		<li>Vortex the collagenase with the TG bundle and place the tube in a 37 &#176;C water bath for 25-30 min. During this time, take the conical tube out of the water bath, vortex, and return to the bath every 5-10 min.</li>
		<li>Centrifuge the collagenase-TG neuron solution for 2 min at 643 x g.</li>
		<li>Under a tissue culture hood, gently aspirate the collagenase with a micropipette.</li>
		<li>Add 5 mL of 1% sterile-filtered trypsin type II and vortex. Place the conical tube in a 37 &#176;C water bath for 5 min.</li>
		<li>Centrifuge the trypsin-TG mix for 5 min at 643 x g. Remove the top portion of trypsin with a micropipette, so that the TG neurons are not removed. There will still be liquid in the tube.</li>
		<li>Add enough media to deactivate the remaining trypsin (at a 1:1 or lower ratio of trypsin to media).</li>
		<li>Count the number of cells and dilute the solution to 200,000 cells/mL (250 &#956;L of cell-containing 50,000 cells).</li>
		<li>Place coated transwell filters from section 1.2 into wells with DP.</li>
		<li>Dilute the cell-containing solution so that there are 200,000 cells/mL. Pipette 250 &#956;L onto the transwell filter, and culture the cells at 37 &#176;C overnight (Figure 1F).</li>
		<li>The next day, replace the media with 1 mL of Co-culture media with 1 &#956;M uridine and 15 &#956;M 5&#39;-fluor-2&#39;deoxyuridine to stop the over-proliferation of mesenchymal cells that may prevent neurite outgrowth. Optional: Add growth factors or inhibitors in this media if attempting further manipulation.</li>
		<li>Culture the cells for additional desired time points. This protocol was optimized for 5 total days of culture with the media changed only on day 2 to add mitotic inhibitors. Longer time periods will require additional media changes.</li>
	</ol>
	</li>
</ol>
3. Sample Collection and Processing
<ol>
	<li>Trigeminal staining
	<ol>
		<li>Pipette 1 mL aliquots of sterile 1x PBS into a 24-well plate under a tissue culture hood for each transwell filter to be processed.</li>
		<li>Remove the liquid atop the filter be removed with a 200-1,000 &#956;L pipette, being careful to leave cell layers intact. Attached cells should remain attached and unattached cells will come loose during this gentle pipetting. A vacuum filter can damage the cell layer and is not recommended for aspiration.</li>
		<li>Transfer the filter to the 1x PBS plate, being sure to remove the top layer of media as mentioned in the previous step.</li>
		<li>Place the plate with all transwell filters and lids on a rocker at 12 rpm or 40-50 rpm on an orbital shaker for 10 min.</li>
		<li>Aspirate the PBS, including the top layer as described above, add 500 &#956;L of 1xPBS and place on the rocker or orbital shaker for an additional 5-10 min rinse.</li>
		<li>Aspirate the 1x PBS once more and replace with 4% paraformaldehyde (PFA). Use at least 500 &#956;L so that the entire filter surface is submerged. Place the plate on a rocker at 12 rpm or orbital shaker at 40-50 rpm at room temperature for 1 h.</li>
		<li>Remove the PFA and rinse the plates twice for 5-10 min each on a rocker with 500 &#956;L of 1x PBS.</li>
		<li>Block with 10% bovine serum albumin (BSA) + 5% goat/donkey serum, depending on the secondary antibody host animal, in 0.05% Tween-1xPBS (PBST). Use 450-500 &#956;L of solution so that the filter is submerged in the liquid. 
		NOTE: At this stage, these plates can be stored at 4 &#176;C for several months to process at a later time. Use parafilm to seal the plate to ensure evaporation does not occur and check regularly for evaporation so that that filter surfaces remain submerged.</li>
		<li>Remove the block and add 450-500 &#956;L of primary antibody in 1% BSA-PBST without an additional rinse. Incubate at 4 &#176;C overnight, gently rocking.</li>
		<li>Remove primary antibody solution and rinse with 500 &#956;L of 1xPBST on a rocker, 3 times, at room temperature</li>
		<li>Remove the PBST, add secondary antibody and incubate on a rocker overnight at 4 &#176;C wrapped in aluminum foil to protect fluorophores from light degradation. Rinse again, 3 times, with 1x PBST on a rocker at room temperature. Replace with 1x PBS. 
		NOTE: At this point, the plate can be stored for several months at 4 &#176;C if wrapped in aluminum foil to prevent fluorophore degradation. Use parafilm to seal the plate to ensure evaporation does not occur and check regularly for evaporation so that that filter surfaces remain submerged. It is best to image within one month.</li>
		<li>For optimal imaging, utilize Thy1-YFP mouse neurons with an anti-GFP antibody to specifically stain the neuronal afferents well above background levels. Use Anti-Neurofilament 200 to precisely stain axonal structures if Thy1-YFP mice are not available.</li>
		<li>Image as desired. Filters do not need to be plated and can instead be placed atop a coverslip or slide for imaging with an inverted microscope. 
		NOTE: Areas of the filter will not contain afferent structures. Take several images with a z-stack depth that captures all afferent structures. Use stitching software to visualize the neurite outgrowth over large areas, or (preferably) entire filter regions.</li>
	</ol>
	</li>
	<li>RNA, protein, and media collection
	<ol>
		<li>While the transwell filters are processing, collect the media in RNAse/DNAse-free tubes and freeze for later assays (ELISA, proteomics, etc.).</li>
		<li>Immediately after media collection, aliquot lysis buffer or Radio Immuno Precipitation Assay (RIPA) buffer with proteinase and phosphatase inhibitors into the wells. This protocol was optimized for 100 &#956;L/well in a 24-well plate.</li>
		<li>Allow buffers to lyse cells for 5 min, then scrape each filter with a new pipette tip, and collect the cell samples in RNAse/DNAse-free tubes. Freeze the lysis for future assays (semi-quantitative and/or quantitative PCR, Western blot, etc.).</li>
	</ol>
	</li>
	<li>Optional immunofluorescence of dental pulp cells:
	<ol>
		<li>Gently lift coverslips from Section 1.1 with forceps and transfer them to different wells with 4% PFA for 1 h with rocking.</li>
		<li>Aspirate PFA and rinse the coverslips twice with 1x PBS. Follow this with permeabilization, blocking and immunofluorescence for markers of interest with standard immunofluorescence techniques.</li>
	</ol>
	</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activating+transcription+factor+3+mRNA+is+upregulated+in+primary+cultures+of+trigeminal+ganglion+neurons.">Activating transcription factor 3 mRNA is upregulated in primary cultures of trigeminal ganglion neurons.</a> Molecular Brain Research. 118 (1-2), 156-159 (2003).</li><li>Lillesaar, C., Arenas, E., Hildebrand, C., Fried, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Responses+of+rat+trigeminal+neurones+to+dental+pulp+cells+or+fibroblasts+overexpressing+neurotrophic+factors+in+vitro.">Responses of rat trigeminal neurones to dental pulp cells or fibroblasts overexpressing neurotrophic factors in vitro.</a> Neuroscience. 119 (2), 443-451 (2003).</li><li>Lillesaar, C., Eriksson, C., Fried, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rat+tooth+pulp+cells+elicit+neurite+growth+from+trigeminal+neurones+and+express+mRNAs+for+neurotrophic+factors+in+vitro.">Rat tooth pulp cells elicit neurite growth from trigeminal neurones and express mRNAs for neurotrophic factors in vitro.</a> Neuroscience Letters. 308 (3), (2001).</li><li>Lillesaar, C., Eriksson, C., Johansson, C. S., Fried, K., Hildebrand, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tooth+pulp+tissue+promotes+neurite+outgrowth+from+rat+trigeminal+ganglia+in+vitro.">Tooth pulp tissue promotes neurite outgrowth from rat trigeminal ganglia in vitro.</a> Journal of neurocytology. 28 (8), 663-670 (1999).</li><li>Chmilewsky, F., Ayaz, W., Appiah, J., About, I., Chung, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nerve+Growth+Factor+Secretion+From+Pulp+Fibroblasts+is+Modulated+by+Complement+C5a+Receptor+and+Implied+in+Neurite+Outgrowth.">Nerve Growth Factor Secretion From Pulp Fibroblasts is Modulated by Complement C5a Receptor and Implied in Neurite Outgrowth.</a> Scientific reports. 6, 31799 (2016).</li><li>Pagella, P., Neto, E., Jim&#195;&#169;nez-Rojo, L., Lamghari, M., Mitsiadis, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microfluidics+co-culture+systems+for+studying+tooth+innervation.">Microfluidics co-culture systems for studying tooth innervation.</a> Frontiers in Physiology. 5, 326 (2014).</li><li>Miura, T., Yokokawa, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+culture+on+a+chip:+Developmental+biology+applications+of+self-organized+capillary+networks+in+microfluidic+devices.">Tissue culture on a chip: Developmental biology applications of self-organized capillary networks in microfluidic devices.</a> Development, Growth &amp; Differentiation. 58 (6), 505-515 (2016).</li></ol>

The authors have nothing to disclose.

The daily activities of the oral cavity require that teeth sense external stimuli and internal inflammation in order to permit proper usage and maintenance. However, only limited information is available regarding the signals that drive the developmental processes of tooth innervation. This protocol provides a method to isolate and co-culture primary DP cells and TG neurons in order to study the cross-communication between the two populations. Several variables were optimized and leave open further avenues of research, a...

Tooth innervation allows teeth to sense pressure, temperature and inflammation, all of which are crucial to the use and maintenance of the tooth organ. Failure to sense tooth pain associated with dental caries and trauma leads to disease progression. Thus, proper innervation is a requirement for normal tooth growth, function and care.
While most organs are fully functional and innervated by the time of birth, tooth development extends into adult life, with tooth innervation and mineralization occurring in concert during postnatal stages1,2. Interestingly, the dental pulp (DP) mesenchyme initially secretes repellant signals during embryogenesis to prevent axon entry into the developing tooth organ, which later shifts to the secretion of attractant factors as the tooth nears eruption3,4. During postnatal stages, afferent axons from the trigeminal (TG) nerve penetrate into and throughout the tooth around the time dentin deposition begins (reviewed in Pagella, P. et al.5). Several in vivo studies have demonstrated that neuronal-mesenchymal interactions guide tooth innervation in mice (reviewed in Luukko, K. et al.6), but few details of the molecular mechanisms are available.
Cell co-cultures provide controlled environments in which investigators can manipulate interactions between neuronal and mesenchymal populations. Co-culture experiments make it possible to delve deeper into the signaling pathways guiding tooth innervation and development. However, several of the conventional methods used to study cells in co-culture present technical challenges. For instance, crystal violet staining of neurite outgrowth can non-specifically stain Schwann cells included in TG bundle dispersions, and there may be peaks in color intensity with relatively small responses7. Microfluidic chambers offer an attractive option, but are considerably more expensive than transwell filters8,9 and only permit the investigation of neuronal responses to DP secretions. To address these issues, we have developed a protocol that allows for: a) precise staining and imaging of TG neurite outgrowth in response to DP secretions, b) genetic modification of DP cells and/or TG neurons to investigate specific signaling pathways, and c) investigation of DP cell responses to factors secreted by TG neurons. This protocol provides the ability to precisely investigate several features of tooth innervation in the controlled environment of an in vitro co-culture assay.

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<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>5-Fluoro-2&#39;-deoxyuridine</td>
			<td>Sigma-Aldrich</td>
			<td>F0503</td>
			<td>Used as a mitotic Inhibitor at 15 &#956;M concentration in co-culture media, Day 2</td>
		</tr>
		<tr>
			<td>24 Well Cell Culture Plate</td>
			<td>Corning</td>
			<td>3524</td>
			<td>Co-culture plate</td>
		</tr>
		<tr>
			<td>Alexa-546 anti-chicken</td>
			<td>Invitrogen</td>
			<td>A-11040</td>
			<td>Secondary to stain neurite outgrowth labeled by anti-GFP antibody, 1:500 dilution</td>
		</tr>
		<tr>
			<td>Anti-GFP Antibody</td>
			<td>Aves Lab, Inc</td>
			<td>GFP-1010</td>
			<td>Primary antibody to label Thy1-YFP neurons, 1:200 dilution</td>
		</tr>
		<tr>
			<td>Anti-Neurofilament 200 antibody</td>
			<td>Sigma-Aldrich</td>
			<td>NO142</td>
			<td>Monoclonal primary antibody to label neurons, 1:1000 dilution, alternative if YFP mice are not available</td>
		</tr>
		<tr>
			<td>B6;129- Tgfbr2tm1Karl/J</td>
			<td>The Jackson Laboratory</td>
			<td>12603</td>
			<td>Tgfbr2f/f mouse model used for dental pulp cells in optimized protocol</td>
		</tr>
		<tr>
			<td>B6.Cg-Tg(Thy1-YFP)16Jrs/J</td>
			<td>The Jackson Laboratory</td>
			<td>3709</td>
			<td>Thy1-YFP mouse model genotype used for trigeminal neurons</td>
		</tr>
		<tr>
			<td>Collagenase Type II</td>
			<td>Millipore</td>
			<td>234155-100MG</td>
			<td>Used to disperse trigeminal neurons</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Gibco</td>
			<td>10437</td>
			<td>Additive to co-culture media</td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>Fine Science Tools</td>
			<td>11413-11</td>
			<td>Fine forceps for TG dissection</td>
		</tr>
		<tr>
			<td>Laminin</td>
			<td>Sigma-Aldrich</td>
			<td>L2020</td>
			<td>Coats the transwell inserts at final concentration of 10 &#956;g/ml, stock solution is assumed at 1.5 mg/ml</td>
		</tr>
		<tr>
			<td>Lysis Buffer (Buffer RLT)</td>
			<td>Qiagen</td>
			<td>79216</td>
			<td>Extracts RNA from dental pulp cells post co-culture</td>
		</tr>
		<tr>
			<td>L-Glutamine</td>
			<td>Gibco</td>
			<td>25030081</td>
			<td>Additive to co-culture media</td>
		</tr>
		<tr>
			<td>Micro-dissecting scissors</td>
			<td>Sigma-Aldrich</td>
			<td>S3146-1EA</td>
			<td>Dissection scissors to open skull</td>
		</tr>
		<tr>
			<td>Microscope Cover Glass</td>
			<td>Fisherbrand</td>
			<td>12-545-81</td>
			<td>Circlular coverslip for optional cell culturing and immunofluorescence processing</td>
		</tr>
		<tr>
			<td>Minimal Essential Medium a</td>
			<td>Gibco</td>
			<td>12571063</td>
			<td>Co-culture media base</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15070063</td>
			<td>Antibiotic additive to co-culture media</td>
		</tr>
		<tr>
			<td>Phosphatase Inhibitor</td>
			<td>Sigma-Aldrich</td>
			<td>04 906 837 001</td>
			<td>Additive to RIPA Buffer for extracting protein from dental pulp cells post co-culture</td>
		</tr>
		<tr>
			<td>Polybrene</td>
			<td>Millipore</td>
			<td>TR-1003-G</td>
			<td>Used to aid in dental pulp cell transfection</td>
		</tr>
		<tr>
			<td>Poly-D-Lysine</td>
			<td>Sigma-Aldrich</td>
			<td>P7280</td>
			<td>Coverslip coating to aid dental pulp cellular adhesion</td>
		</tr>
		<tr>
			<td>Protease Inhibitors</td>
			<td>Millipore</td>
			<td>05 892 791 001</td>
			<td>Additive to RIPA Buffer for extracting protein from dental pulp cells post co-culture</td>
		</tr>
		<tr>
			<td>RNAse/DNAse free eppendorf tubes</td>
			<td>Denville</td>
			<td>C-2172</td>
			<td>Presterilized 1.7 ml tubes for RNA, DNA or protein collection at the end of assay</td>
		</tr>
		<tr>
			<td>ThinCert Cell Culture Insert</td>
			<td>Greiner Bio-One</td>
			<td>662631</td>
			<td>Transwell inserts for trigeminal neurons in co-culture assays</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.25%)</td>
			<td>Gibco</td>
			<td>25200056</td>
			<td>Used fto disperse dental pulp cells</td>
		</tr>
		<tr>
			<td>Trypsin Type II</td>
			<td>Sigma-Aldrich</td>
			<td>T-7409</td>
			<td>Used to disperse trigeminal neurons</td>
		</tr>
		<tr>
			<td>Ultra Fine Forceps</td>
			<td>Fine Science Tools</td>
			<td>11370-40</td>
			<td>Ultra fine forceps for dissection</td>
		</tr>
		<tr>
			<td>Uridine</td>
			<td>Sigma-Aldrich</td>
			<td>U3750</td>
			<td>Used as a mitotic Inhibitor at 1 &#956;M concentration in co-culture media, Day 2</td>
		</tr>
		<tr>
			<td>Vacuum Filtration System</td>
			<td>Millipore</td>
			<td>SCNY00060</td>
			<td>Steriflip disposable filter, 50 &#956;m nylon net filter</td>
		</tr>
		<tr>
			<td>Vial forceps</td>
			<td>Fine Science Tools</td>
			<td>110006-15</td>
			<td>Long forceps for tissue transfer to conicals</td>
		</tr>
	</tbody>
</table>

a co-culture method to study neurite outgrowth in response to dental pulp paracrine signals

Tooth innervation allows teeth to sense pressure, temperature and inflammation, all of which are crucial to the use and maintenance of the tooth organ. Without sensory innervation, daily oral activities would cause irreparable damage. Despite its importance, the roles of innervation in tooth development and maintenance have been largely overlooked. Several studies have demonstrated that DP cells secrete extracellular matrix proteins and paracrine signals to attract and guide TG axons into and throughout the tooth. However, few studies have provided detailed insight into the crosstalk between the DP mesenchyme and neuronal afferents. To address this gap in knowledge, researchers have begun to utilize co-cultures and a variety of techniques to investigate these interactions. Here, we demonstrate the multiple steps involved in co-culturing primary DP cells with TG neurons dispersed on an overlying transwell filter with large diameter pores to allow axonal growth through the pores. Primary DP cells with the gene of interest flanked by loxP sites were utilized to facilitate gene deletion using an Adenovirus-Cre-GFP recombinase system. Using TG neurons from the Thy1-YFP mouse allowed for precise afferent imaging, with expression well above background levels by confocal microscopy. The DP responses can be investigated via protein or RNA collection and analysis, or alternatively, through immunofluorescent staining of DP cells plated on removable glass coverslips. Media can be analyzed using techniques such as proteomic analyses, although this will require albumin depletion due to the presence of fetal bovine serum in the media. This protocol provides a simple method that can be manipulated to study the morphological, genetic, and cytoskeletal responses of TG neurons and DP cells in response to the controlled environment of a co-culture assay.

These results show that TG neurite outgrowth was increased in the presence of primary DP cells in the underlying well compared to the control of TG neurite monoculture (Figure 2A,C). There is some assay-to-assay variability in neurite outgrowth. Thus, a TG neuron monoculture should be included in all assays as a control to detect the basal levels of neurite outgrowth. Primary cells from the Tgfbr2f/f mouse were used in this protocol after infection of Ad-Cre-GFP a...

Watch this Scientific Journal Video about A Co-Culture Method to Study Neurite Outgrowth in Response to Dental Pulp Paracrine Signals at JoVE.com

A Co-Culture Method to Study Neurite Outgrowth in Response to Dental Pulp Paracrine Signals

We describe the isolation, dispersion and plating of dental pulp (DP) primary cells with trigeminal (TG) neurons cultured atop overlying transwell filters. Cellular responses of DP cells can be analyzed with immunofluorescence or RNA/protein analysis. Immunofluorescence of neuronal markers with confocal microscopy permits the analysis of neurite outgrowth responses.

Tooth innervation allows teeth to sense pressure, temperature and inflammation, all of which are crucial to the use and maintenance of the tooth organ. ...

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Research

JoVE Journal

Developmental Biology

6.0K Views.  University of Alabama at Birmingham. We describe the isolation, dispersion and plating of dental pulp (DP) primary cells with trigeminal (TG) neurons cultured atop overlying transwell filters. Cellular responses of DP cells can be analyzed with immunofluorescence or RNA/protein analysis. Immunofluorescence of neuronal markers with confocal microscopy permits the analysis of neurite outgrowth responses.

Video: A Co-Culture Method to Study Neurite Outgrowth in Response to Dental Pulp Paracrine Signals

An Optogenetic Approach for Assessing Formation of Neuronal Connections in a Co-culture System

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are reprogrammed from human fibroblasts using episomal vectors. Colonies of iPSCs can be observed 30 days after initiation of fibroblast reprogramming. Pluripotent colonies are manually picked and grown in neural induction medium to permit differentiation into neural progenitor cells (NPCs). iPSCs rapidly convert into neuroepithelial cells within 1 week and retain the capability to self-renew when maintained at a high culture density. Primary mouse NPCs are differentiated into astrocytes by exposure to a serum-containing medium for 7 days and form a monolayer upon which embryonic day 18 (E18) rat cortical neurons (transfected with channelrhodopsin-2 (ChR2)) are added. Human NPCs tagged with the fluorescent protein, tandem dimer Tomato (tdTomato), are then seeded onto the astrocyte/cortical neuron culture the following day and allowed to differentiate for 28 to 35 days. We demonstrate that this system forms synaptic connections between iPSC-derived neurons and cortical neurons, evident from an increase in the frequency of synaptic currents upon photostimulation of the cortical neurons. This co-culture system provides a novel platform for evaluating the ability of iPSC-derived neurons to create synaptic connections with other neuronal populations.

A protocol to generate a co-culture system consisting of neurons derived from induced pluripotent stem cells (iPSCs), primary cortical neurons and astrocytes is described. This co-culture system allows detection of the formation of synaptic contacts and circuits between new, iPSC-derived neurons and pre-existing cortical neurons expressing channelrhodopsin-2.

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are ...

Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many myopathies affecting skeletal muscle function can be quickly and easily assessed in zebrafish over the first few days of embryogenesis. By 24 hr post-fertilization (hpf), wildtype zebrafish spontaneously contract their tail muscles and by 48 hpf, zebrafish exhibit controlled swimming behaviors. Reduction in the frequency of, or other alterations in, these movements may indicate a skeletal muscle dysfunction. To analyze swimming behavior and assess muscle performance in early zebrafish development, we utilize both touch-evoked escape response and locomotion assays.
Touch-evoked escape response assays can be used to assess muscle performance during short burst movements resulting from contraction of fast-twitch muscle fibers. In response to an external stimulus, which in this case is a tap on the head, wildtype zebrafish at 2 days post-fertilization (dpf) typically exhibit a powerful burst swim, accompanied by sharp turns. Our method quantifies skeletal muscle function by measuring the maximum acceleration during a burst swimming motion, the acceleration being directly proportional to the force produced by muscle contraction.
In contrast, locomotion assays during early zebrafish larval development are used to assess muscle performance during sustained periods of muscle activity. Using a tracking system to monitor swimming behavior, we obtain an automated calculation of the frequency of activity and distance in 6-day old zebrafish, reflective of their skeletal muscle function. Measurements of swimming performance are valuable for phenotypic assessment of disease models and high-throughput screening of mutations or chemical treatments affecting skeletal muscle function.

Zebrafish are an excellent model to study muscle function and disease. During early embryogenesis zebrafish begin regular muscle contractions producing rhythmic swimming behavior, which is altered when the muscle is disrupted. Here we describe a touch-evoked response and locomotion assay to examine swimming performance as a measure of muscle function.

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many ...

Application of Impermeable Barriers Combined with Candidate Factor Soaked Beads to Study Inductive Signals in the Chick

The chick embryo provides a superb vertebrate model that can be used to dissect developmental questions in a direct way. Its accessibility and robustness following surgical intervention are key experimental strengths. Mica plates were the first barriers used to prevent chick limb bud initiation1. Protocols that use aluminum foil as an impermeable barrier to wing bud or leg bud induction and or initiation are described. We combine this technique with bead placement lateral to the barrier to exogenously supply candidate endogenous factors that have been blocked by the barrier. The results are analyzed using in situ hybridization of subsequent gene expression. Our main focus is on the role of retinoic acid signaling in the induction and later initiation of the chick embryo fore and hindlimb. We use BMS 493 (an inverse agonist of retinoic acid receptors (RAR)) soaked beads implanted in the lateral plate mesoderm (LPM) to mimic the effect of a barrier placed between the somites (a source of retinoic acid (RA)) and the LPM from which limb buds grow. Modified versions of these protocols could also be used to address other questions on the origin and timing of inductive cues. Provided the region of the chick embryo is accessible at the relevant developmental stage, a barrier could be placed between the two tissues and consequent changes in development studied. Examples may be found in the developing brain, axis extension and in organ development, such as liver or kidney induction.

Protocols using impermeable barriers to block induction events between tissues of the main body axis and flank in the chick embryo required for limb formation are described. Beads soaked in candidate inductive signals are used to overcome the effect of barrier placement and analysis of gene expression confirms this.

The chick embryo provides a superb vertebrate model that can be used to dissect developmental questions in a direct way. Its accessibility and robustness ...

A 5-mC Dot Blot Assay Quantifying the DNA Methylation Level of Chondrocyte Dedifferentiation In Vitro

The dedifferentiation of hyaline chondrocytes into fibroblastic chondrocytes often accompanies monolayer expansion of chondrocytes in vitro. The global DNA methylation level of chondrocytes is considered to be a suitable biomarker for the loss of the chondrocyte phenotype. However, results based on different experimental methods can be inconsistent. Therefore, it is important to establish a precise, simple, and rapid method to quantify global DNA methylation levels during chondrocyte dedifferentiation.
Current genome-wide methylation analysis techniques largely rely on bisulfite genomic sequencing. Due to DNA degradation during bisulfite conversion, these methods typically require a large sample volume. Other methods used to quantify global DNA methylation levels include high-performance liquid chromatography (HPLC). However, HPLC requires complete digestion of genomic DNA. Additionally, the prohibitively high cost of HPLC instruments limits HPLC&#39;s wider application.
In this study, genomic DNA (gDNA) was extracted from human chondrocytes cultured with varying number of passages. The gDNA methylation level was detected using a methylation-specific dot blot assay. In this dot blot approach, a gDNA mixture containing the methylated DNA to be detected was spotted directly onto an N+ membrane as a dot inside a previously drawn circular template pattern. Compared with other gel electrophoresis-based blotting approaches and other complex blotting procedures, the dot blot method saves significant time. In addition, dot blots can detect overall DNA methylation level using a commercially available 5-mC antibody. We found that the DNA methylation level differed between the monolayer subcultures, and therefore could play a key role in chondrocyte dedifferentiation. The 5-mC dot blot is a reliable, simple, and rapid method to detect the general DNA methylation level to evaluate chondrocyte phenotype.

A 5-mC Dot Blot Assay Quantifying the DNA Methylation Level of Chondrocyte Dedifferentiation In Vitro

We present a method to quantify DNA methylation based on the 5-methylcytosine (5-mC) dot blot. We determined the 5-mC levels during chondrocyte dedifferentiation. This simple technique could be used to quickly determine the chondrocyte phenotype in ACI treatment.

The dedifferentiation of hyaline chondrocytes into fibroblastic chondrocytes often accompanies monolayer expansion of chondrocytes in vitro. The global ...

The Aortic Ring Co-culture Assay: A Convenient Tool to Assess the Angiogenic Potential of Mesenchymal Stromal Cells In Vitro

Angiogenesis is a complex, highly regulated process responsible for providing and maintaining adequate tissue perfusion. Insufficient vasculature maintenance and pathological malformations can result in severe ischemic diseases, while overly abundant vascular development is associated with cancer and inflammatory disorders. A promising form of pro-angiogenic therapy is the use of angiogenic cell sources, which can provide regulatory factors as well as physical support for newly developing vasculature.
Mesenchymal Stromal Cells (MSCs) are extensively investigated candidates for vascular regeneration due to their paracrine effects and their ability to detect and home to ischemic or inflamed tissues. In particular, first trimester human umbilical cord perivascular cells (FTM HUCPVCs) are a highly promising candidate due to their pericyte-like properties, high proliferative and multilineage potential, immune-privileged properties, and robust paracrine profile. To effectively evaluate potentially angiogenic regenerative cells, it is a requisite to test them in reliable and &#34;translatable&#34; pre-clinical assays. The aortic ring assay is an ex vivo angiogenesis model that allows for easy quantification of tubular endothelial structures, provides accessory supportive cells and extracellular matrix (ECM) from the host, excludes inflammatory components, and is fast and inexpensive to set up. This is advantageous when compared to in vivo models (e.g., corneal assay, Matrigel plug assay); the aortic ring assay can track the administered cells and observe intercellular interactions while avoiding xeno-immune rejection.
We present a protocol for a novel application of the aortic ring assay, which includes human MSCs in co-cultures with developing rat aortic endothelial networks. This assay allows for the analysis of the MSC contribution to tube formation and development through physical pericyte-like interactions and of their potency for actively migrating to sites of angiogenesis, and for evaluating their ability to perform and mediate ECM processing. This protocol provides further information on changes in MSC phenotype and gene expression following co-culture.

The Aortic Ring Co-culture Assay: A Convenient Tool to Assess the Angiogenic Potential of Mesenchymal Stromal Cells In Vitro

Here, we present a novel application of the aortic ring assay where prelabelled mesenchymal cells are co-cultured with rat aorta-derived endothelial networks. This novel method allows visualization of Mesenchymal Stromal Cells (MSCs) homing and integration with endothelial networks, quantification of network properties, and evaluation of MSC immunophenotypes and gene expression.

Angiogenesis is a complex, highly regulated process responsible for providing and maintaining adequate tissue perfusion. Insufficient vasculature ...

Organotypic Culture Method to Study the Development Of Embryonic Chicken Tissues

The embryonic chicken is commonly used as a reliable model organism for vertebrate development. Its accessibility and short incubation period makes it ideal for experimentation. Currently, the study of these developmental pathways in the chicken embryo is conducted by applying inhibitors and drugs at localized sites and at low concentrations using a variety of methods. In vitro tissue&#160;culturing is a technique that enables the study of tissues separated from the host organism, while simultaneously bypassing many of the physical limitations present when working with whole embryos, such as the susceptibility of embryos to high doses of potentially lethal chemicals. Here, we present an organotypic culturing protocol for culturing the embryonic chicken half head in vitro, which presents new opportunities for the examination of developmental processes beyond the currently established methods.

Here, we present an organotypic culturing protocol to grow embryonic chicken organs in vitro. Using this method, the development of embryonic chicken tissue can be studied, while maintaining a high degree of control over the culture environment.

The embryonic chicken is commonly used as a reliable model organism for vertebrate development. Its accessibility and short incubation period makes it ...

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by Sertoli cell junctions at the blood-testis barrier (BTB), which is the tightest tissue barrier in the mammalian body and segregates the seminiferous epithelium into two compartments, a basal and an adluminal. The BTB creates a unique microenvironment for germ cells in meiosis I/II and for the development of postmeiotic spermatids into spermatozoa via spermiogenesis. Here, we describe a reliable assay to monitor BTB integrity of mouse testis in vivo. An intact BTB blocks the diffusion of FITC-conjugated inulin from the basal to the apical compartment of the seminiferous tubules. This technique is suitable for studying gene candidates, viruses, or environmental toxicants that may affect BTB function or integrity, with an easy procedure and a minimal requirement of surgical skills compared to alternative methods.

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Here, we present a protocol to assess the blood-testis barrier integrity by injecting inulin-FITC into testes. This is an efficient in vivo method to study blood-testis barrier integrity that can be compromised by genetic and environmental elements.

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by ...

Isolation, Propagation, and Prion Protein Expression During Neuronal Differentiation of Human Dental Pulp Stem Cells

Bioethical issues related to the manipulation of embryonic stem cells have hindered advances in the field of medical research. For this reason, it is very important to obtain adult stem cells from different tissues such as adipose, umbilical cord, bone marrow and blood. Among the possible sources, dental pulp is particularly interesting because it is easy to obtain in respect of bioethical considerations. Indeed, human Dental Pulp Stem Cells (hDPSCs) are a type of adult stem cells able to differentiate in neuronal-like cells and can be obtained from the third molar of healthy patients (13-19 ages). In particular, the dental pulp was removed with an excavator, cut into small slices, treated with collagenase IV and cultured in a flask. To induce the neuronal differentiation, hDPSCs were stimulated with EGF/bFGF for 2 weeks. Previously, we have demonstrated that during the differentiation process the content of cellular prion Protein (PrPC) in hDPSCs increased. The cytofluorimetric analysis showed an early expression of PrPC that increased after neuronal differentiation process. Ablation of PrPC by siRNA PrP prevented neuronal differentiation induced by EGF/bFGF. In this paper, we illustrate that as we enhanced the isolation, separation and in vitro cultivation methods of hDPSCs with several easy procedures, more efficient cell clones were obtained and large-scale expansion of the mesenchymal stem cells (MSCs) was observed. We also show how the hDPSCs, obtained with methods detailed in the protocol, are an excellent experimental model to study the neuronal differentiation process of MSCs and subsequent cellular and molecular processes.

Here we present a protocol for human Dental Pulp Stem Cells isolation and propagation in order to evaluate the prion protein expression during neuronal differentiation process.

Bioethical issues related to the manipulation of embryonic stem cells have hindered advances in the field of medical research. For this reason, it is very ...

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C (UV-C) Damage

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational health hazard. Here, we demonstrate the exposure of primary stem cells from the mouse ocular surface to UV-C radiation. Reactive oxygen species (ROS) formation is the readout of the extent of oxidative stress/damage. In an experimental in vitro setting, it is also essential to assess the percentage of dead cells generated due to oxidative stress. In this article, we will demonstrate the 2&#39;,7&#39;-Dichlorofluoresceindiacetate (DCFDA) staining of UV-C exposed mouse primary ocular surface stem cells and their quantification based on the fluorescent images of DCFDA staining. DCFDA staining directly corresponds to ROS generation. We also demonstrate the quantification of dead and live cells by simultaneous staining with propidium iodide (PI) and Hoechst 3332 respectively and the percentage of DCFDA (ROS positive) and PI positive cells.

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C UV-C Damage

This protocol demonstrates the simultaneous detection of reactive oxygen species (ROS), live cells, and dead cells in live primary cultures from mouse ocular surface cells. 2&#39;,7&#39;-Dichlorofluoresceindiacetate, propidium iodide, and Hoechst staining are used to assess the ROS, dead cells, and live cells, respectively, followed by imaging and analysis.

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational ...

An Enhanced Green Fluorescence Protein-based Assay for Studying Neurite Outgrowth in Primary Neurons

Neurite outgrowth is a fundamental event in the formation of the neural circuits during nervous system development. Severe neurite damage and synaptic dysfunction occur in various neurodegenerative diseases and age-related degeneration. Investigation of the mechanisms that regulate neurite outgrowth would not only shed valuable light on brain developmental processes but also on such neurological disorders. Due to the low transfection efficiency, it is currently challenging to study the effect of a specific protein on neurite outgrowth in primary mammalian neurons. Here, we describe a simple method for the investigation of neurite outgrowth by the co-transfection of primary rat cortical neurons with EGFP and a protein of interest (POI). This method allows the identification of POI transfected neurons through the EGFP signal, and thus the effect of the POI on neurite outgrowth can be determined precisely. This EGFP-based assay provides a convenient approach for the investigation of pathways regulating neurite outgrowth.

In this report, we describe a simple protocol for studying neurite outgrowth in embryonic rat cortical neurons by co-transfecting with EGFP and the protein of interest.

Neurite outgrowth is a fundamental event in the formation of the neural circuits during nervous system development. Severe neurite damage and synaptic ...