This work was supported by National Heart, Lung and Blood Institute grants R01088554, T32HL007284 and American Heart Association predoctoral fellowship 14PRE20420024. We especially thank the St. Jude Cellular Imaging Shared Research Core, including Randall Wakefield and Linda Horner for the electron microscopy images, Adam Straub for assistance with representative immunofluorescence images,&#160;and Pooneh Bagher for her valuable feedback on the manuscript protocol.

1. Plating Cells for the VCCC
<ol>
	<li>Culture large 225 cm2 flasks of EC and SMC under sterile conditions at 37 &#176;C, until 70-90% confluent. Ensure that more EC than SMC are used; if using primary human EC and SMC, typically 3 large flasks of EC to 1 large flask of SMC.
	<ol>
		<li>Use primary cells at lower passages and do not use past passage 10. Choose the culture media based upon cells to be co-cultured. For primary human coronary artery EC, use EBM-2 EC basal media with EGM2 MV growth factors. For primary human coronary SMC, use SmBM SMC basal media with SmGM-2 growth factors. Prior to use with cultured cells, add growth factors to the media.</li>
	</ol>
	</li>
	<li>Construction of the VCCC Day 1: Spray large (150 mm) Petri dishes and lids with disinfectant (e.g., Cavicide) and wipe away with a paper towel or lint-free wipes. Then spray the Petri dishes with 70% ethanol (EtOH) and place them in the hood to air dry.</li>
	<li>Open the plate of filter inserts (see the Table of Materials) under sterile conditions, and coat the SMC side (bottom side of filter) of the insert with fibronectin solution. Keeping the insert in the provided 6-well plate, pipet the fibronectin solution through the side slits to the bottom of the plate, making sure that the bottom half of the filter is covered (no more than 1 mL).</li>
	<li>After 30 min of treatment with fibronectin solution at room temperature in the cell culture hood, take inserts out of the plate, vacuum any excess solution, and invert, placing into the bottom half of the clean Petri dish. Set aside. Be sure not to disturb the filter insert membrane when suctioning excess solution; this can be avoided by gently tilting the plate, and vacuuming the solution off the edge of the insert.</li>
	<li>Trypsinize a 225 cm2 flask of SMC with 3 mL of pre-warmed 2X Trypsin-EDTA solution, and once the cells have lifted off the plate, add 9 mL SMC media to neutralize the trypsin (3:1, media:trypsin). Transfer to a conical tube, mix well, and pipet 10 &#181;L onto a hemocytometer to determine cell number.
	<ol>
		<li>Count the number of cells in 5 x 5 boxes (i.e., 18), multiply this by 10,000 to get the number of cells in 1 mL (180,000). Then multiply by 12 mL (total volume of cell suspension, 2.16 million SMC). Ensure that there are enough SMC to plate the desired number of wells. 
		NOTE: For example, 1 well should have 75,000 SMC and thus 18 wells would have 1.35 million total SMC. Divide 1.35 million cells needed by 180,000 cells per 1 mL and this results in 7.5 mL of the cell suspension that will be needed for plating. Then calculate the final volume needed (18 wells x 750 &#181;L = 13.5 mL). Thus, take 7.5 mL of cell suspension, bring the volume up to 13.5 mL with pre-warmed SMC media, and mix well.</li>
	</ol>
	</li>
	<li>Plate 750 &#181;L of the cell suspension containing approximately 75,000 SMC onto the bottom side of each filter, being careful to not spill any of the media and cell solution. Place the lid on top and carefully transfer the Petri dish with the inserts into the incubator at 37 &#176;C overnight. 
	NOTE: The optimum cell density for other cell types will need to be empirically determined.</li>
	<li>On day 2, fill clean 6-well dishes with 2 mL of fresh, pre-warmed SMC media. Remove the plates from the incubator. Take the inserts out one at a time and remove the media by suction, then place into the 6-well dish with SMC media.</li>
	<li>Coat the opposite side of the filter (EC side) with 1 mL of a 0.5% bovine gelatin solution for at least 30 min at 37 &#176;C.</li>
	<li>Trypsinize the 225 cm2 flask(s) of EC with 3 mL of pre-warmed 2X Trypsin-EDTA (10x diluted in sterile Dulbecco&#39;s PBS) solution, with gentle tapping of the flask in order to lift the cells off the plate. 
	NOTE: It may be necessary to place the flask back into the incubator at 37 &#176;C for 1-3 min.
	<ol>
		<li>Once the cells have lifted off the plate, add 9 mL EC media to neutralize the trypsin (3:1). Transfer to a conical tube, pool together, mix well, and pipet 10 &#181;L onto a hemocytometer to determine cell number.
		<ol>
			<li>Count the number of cells in 5 x 5 boxes (i.e., 60), multiply this by 10,000 to get the number of cells in 1 mL (600,000). Then multiply by 12 mL (total volume of cell suspension, 7.2 million EC). Ensure that there are enough EC to plate the desired number of wells. 
			NOTE: Approximately 360,000 EC are required per well. For example, 18 wells require 6.48 million EC. Divide 6.48 million cells needed by 600,000 EC per 1 mL and this results in 10.8 mL of the cell suspension that will be needed for plating. Then calculate the final volume needed (18 wells x 1 mL = 18 mL). Thus, take 10.8 mL of cell suspension, bring the volume up to 18 mL with pre-warmed EC media, and gently resuspend.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Plate 1 mL of 360,000 EC onto the top side of each insert filter. Let the cells incubate undisturbed at 37 &#176;C for 24 h. Replace medium on Day 4. Harvest on Day 5.</li>
</ol>
2. Harvesting Fractions from the VCCC
<ol>
	<li>Perform the isolation in a 4 &#176;C room and keep all instruments on ice.</li>
	<li>Keep in mind the number of individual inserts being isolated and add lysis buffer to a 50 mL tube labeled MEJ (for the isolated filters). Then label two 10 mm dishes; one for EC and one for SMC.</li>
	<li>Adjust the volume of lysis buffer depending on how concentrated the lysate needs to be; for 15 inserts, 500 &#181;L of lysis buffer for the MEJ tube and 250 &#181;L per Petri dish is recommended.</li>
	<li>Work with one 6-well plate of inserts at a time. Suction off the medium in the cell culture hood and then transport to a cold room on ice. 
	NOTE: The following instructions are for 1 insert isolation.</li>
	<li>Pipet 10 &#181;L of 1x PBS onto the SMC side of the insert.</li>
	<li>Use the cell lifter to scrape down the cells to one end of the insert, and transfer the cells from the lifter to the SMC Petri dish that has lysis buffer in it.</li>
	<li>Once the cell lifter has touched the lysis buffer in the dish, do not allow it to touch the insert filter until it has been completely wiped and dried off on paper towels. Repeat the scraping of the SMC filter at least one to two times to fully remove the remaining SMC cell debris.</li>
	<li>Turn the insert over and pipet 10 &#181;L of 1x PBS into the upper/EC side of the insert filter. Repeat steps 2.6 and 2.7 with a cell scraper and the EC Petri dish.</li>
	<li>Collect the remaining cell and PBS slurry from the EC side of the filter with a pipette and transfer to the EC Petri dish with lysis buffer. Be very thorough in scraping the cells off both sides of the filter as to leave minimal EC/SMC contamination in the MEJ fraction.</li>
	<li>Carefully use the scalpel to cut out the filter from the plastic insert. Do this by cutting 70-80% of it away from the plastic and use forceps to pull the filter completely off the plastic insert structure. Put the filter in the 50 mL conical tube labeled MEJ with lysis buffer, and ensure it sits in the lysis buffer and remains wet.</li>
	<li>Repeat the isolation steps, in groups of 6, until all of the inserts have been processed.</li>
	<li>Ensure that different treatment groups are kept in separate tubes and dishes.</li>
	<li>Vortex the 50 mL MEJ conical tube on full strength for 15 s or until mixed well. Remove the filters from the MEJ conical with forceps, dragging them along the side walls of the tube as to leave the maximum amount of proteins and liquid in the tube. After this point, discard the filters.</li>
	<li>Once all the filters have been removed from the MEJ conical tube, do a quick spin (10-15 s at max speed ~16,000 x g) in a centrifuge to pull all the liquid and proteins to the bottom of the tube. Transfer the contents of the MEJ tube and the EC and SMC Petri dishes into separate, smaller centrifuge tubes.</li>
	<li>Optional: Sonicate the contents of the MEJ/SMC/EC tubes on setting 4 for three 10 s pulses on ice or homogenize gently by hand.</li>
	<li>Centrifuge at max speed (~16,000 x g) 15 min at 4 &#176;C. Pipet out the supernatant and assay the lysate for protein content using the bicinchoninic assay method.</li>
</ol>
3. Harvesting the VCCC for En Face Immunofluorescence
<ol>
	<li>On Day 5 of VCCC incubation, remove the inserts from the incubator. Working in the cell culture hood, suction off the SMC media from each of the lower wells of the insert filter and rinse two times with 1x PBS. Repeat with the EC side.</li>
	<li>Keeping the inserts intact and in the plate, add 4% paraformaldehyde (PFA) for 10-20 min in a walk-in 4 &#176;C room on a gentle shaker. Wash both sides of the insert filter with 1x PBS for 5 min and repeat two times. 
	CAUTION: PFA is toxic and must be handled carefully.</li>
	<li>Perform the blocking, primary and secondary antibody incubations in the plate, ensuring that both sides of the filter are kept in the antibody + blocking solution (9 mL PBS, 25 &#181;L Triton X100, 50 mg bovine serum albumin, 500 &#181;L normal serum)</li>
	<li>To mount the filter on a slide, cut the filter out of the plastic insert with a scalpel mounted on a handle and place on a microscope slide with mounting medium.</li>
</ol>
4. Harvesting the VCCC for Transverse Immunofluorescence
<ol>
	<li>On Day 5 of VCCC incubation, remove the inserts from the incubator. Suction off the media from both sides of the insert filter in the cell culture hood and rinse each side of the filter two times with 1x PBS.</li>
	<li>Keeping the inserts intact and in the 6-well plate, add 4% PFA and incubate overnight in a walk-in 4 &#176;C room on a gentle shaker. 
	CAUTION: PFA is toxic and must be handled carefully.</li>
</ol>
5. Embedding the VCCC for Transverse Immunofluorescence
<ol>
	<li>After filter inserts have been fixed approximately for 24 h in 4% PFA, transfer to 70% EtOH for at least 24 h, but up to several weeks.</li>
	<li>Process the filters on a long (overnight) run on an automated tissue processor. Use the program as follows: 70% EtOH for 30 min, 85% EtOH 30 min, 95% EtOH 30 min, 100% EtOH 30 min, 100% EtOH 45 min, 100% EtOH 45 min, Xylene 30 min, Xylene 30 min, Xylene 45 min, 60 &#176;C paraffin 30 min, 60 &#176;C paraffin 30 min, 60 &#176;C paraffin 45 min.</li>
	<li>After processing, transfer the harvested filters to the embedding station into liquid paraffin, keeping the filter in the cassette.</li>
	<li>Remove the filter from the cassette, carefully using forceps to hold it just at the edge. With scissors, cut the filter in half, forming two semi-circular shaped halves. Lay each of the 2 halves on the cold plate portion of the embedding station just long enough to fill the embedding mold with liquid paraffin.</li>
	<li>Once filled with liquid paraffin, very briefly touch the embedding mold to the cold plate, then embed each half of the filter into embedding mold (cut side down); this ensures that during sectioning, one will be cutting from the center of the filter towards the edge.</li>
	<li>During embedding, use forceps to hold the filter in place to prevent the halves from falling into each other until the paraffin is cool enough for them to stand alone. Move the embedding mold to a cold plate until completely solidified. 
	NOTE: Cooling blocks on ice trays helps to prevent wrinkled sections.</li>
</ol>
6. Sectioning and Staining the VCCC for Transverse Immunofluorescence
<ol>
	<li>For the paraffin blocks sectioning and slide mounting: cut 5 &#181;m sections, place the sections into a 42 &#176;C water bath, and pick up the sections using positively charged slides. Then keep slides overnight in a 42 &#176;C oven to dry.</li>
	<li>Before beginning the rehydration steps, bake the slides at 60 &#176;C to melt the paraffin and help adhere the VCCC sections to the slide. Cool briefly, then rehydrate the slides through two changes of xylene, 100% EtOH, 95% EtOH, one change of 70% EtOH, and two changes of distilled water.</li>
	<li>Avoid antigen retrieval methods that involve microwaving or heating as sections may fall off the slide.</li>
	<li>Perform the standard blocking for 1 h at room temperature, then the overnight incubation with the primary antibody at 4 &#176;C, followed by washing steps and secondary antibody for 1 h at room temperature. Mount with the mounting media and coverslip.</li>
</ol>

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The authors have nothing to disclose.

The VCCC has a number of advantages in terms of recapitulating MEJs in vitro, but there are points of discussion when determining if the VCCC can be utilized for specific application. For example, if different cell types are to be used with this model, it will need to be optimized. We have used primary human cells but it is possible to isolate cells from bovine aortas24, or wildtype or knockout mice, and use them in the VCCC1,5<su...

In small diameter resistance arteries, a thin internal elastic lamina (IEL) separates endothelial (EC) and smooth muscle cells (SMC). The cells can project through holes in this elastic matrix and make direct cytoplasmic connections via gap junction channels1,2. This unique structure is termed the myoendothelial junction (MEJ). The MEJ is a small (approximately 0.5 &#181;m x 0.5 &#181;m, depending on the vascular bed), cellular projection composed predominantly of endothelial cells but may originate from smooth muscle as well3,4. A number of investigators have demonstrated the immense complexity of signaling networks that occur selectively at the MEJ, rendering it an especially important location for facilitating bi-directional signaling between endothelium and smooth muscle5,6,7,8,9,10.
However, a mechanistic dissection of signaling pathways at the MEJ is difficult in an intact artery. Because the MEJ is a cellular projection, it is not currently possible to isolate an in vivo MEJ from the vascular wall. For this reason, the VCCC model1 was developed. Importantly, the VCCC replicates physiological endothelial morphology11 and polarization of signaling between the apical and MEJ portions of the cell12. It was this unique model that facilitated the discovery that alpha hemoglobin is in the endothelial cell, polarized to the MEJ. This is in contrast to conventionally cultured endothelial cells which do not express alpha hemoglobin13. For researchers interested in microvascular endothelium, it may be more appropriate to use endothelial cells that have been cultured in the VCCC, particularly if the intent is to dissect signaling pathways that occur in small diameter resistance arteries.
Using a sturdy plastic insert containing a filter with small diameter pores (0.4 &#181;m in diameter) to co-culture two distinct cell types prevents the cells from migrating between layers. It results in a 10 &#181;m distance between the cells, which is significantly longer than in vivo, but still replicates many of the in vivo characteristics of MEJs, including protein localization and second messenger signaling1,14. In addition, the VCCC allows for targeting of cell type-specific signaling via the addition of an agonist or antagonist to the specific cellular compartment. For example, loading the EC with BAPTA-AM to chelate calcium, and stimulating the SMC with phenylephrine14. In contrast to other descriptions of co-culture models15,16,17,18,19,20,21, this provides instructions on isolating the distinct fractions for mRNA and protein, including the distinct MEJ fraction within the filter pores. The addition of this technique to the VCCC allows for specific investigation of changes in mRNA localization or transcription22, protein phosphorylation12,23, and protein activity12. This article will describe plating of EC on top of the filter and SMC on the bottom, although it is possible to culture the two cell types in different conformations11,18,24.

<table>
	<tbody>
		<tr>
			<td>24mm diameter, 0.4&#181;m Transwell Polyester membrane filter insert</td>
			<td>Corning</td>
			<td>3450</td>
			<td>This is one 6 well plate of inserts</td>
		</tr>
		<tr>
			<td>225cm2 flask</td>
			<td>Corning</td>
			<td>431082</td>
			<td></td>
		</tr>
		<tr>
			<td>6 well plates (without filter inserts)</td>
			<td>Corning</td>
			<td>3506</td>
			<td></td>
		</tr>
		<tr>
			<td>Primary human coronary artery endothelial cells</td>
			<td>Lonza</td>
			<td>CC-2585</td>
			<td></td>
		</tr>
		<tr>
			<td>Primary human coronary artery smooth muscle cells</td>
			<td>Lonza</td>
			<td>CC-2583</td>
			<td></td>
		</tr>
		<tr>
			<td>EBM-2 EC Basal media</td>
			<td>Lonza</td>
			<td>CC-3156</td>
			<td></td>
		</tr>
		<tr>
			<td>EC growth factors (EGM2 MV Single quots)</td>
			<td>Lonza</td>
			<td>CC-4147</td>
			<td>Add to basal media before use</td>
		</tr>
		<tr>
			<td>SmBM SMC basal media</td>
			<td>Lonza</td>
			<td>CC-3181</td>
			<td></td>
		</tr>
		<tr>
			<td>SMC growth factors (SmGM-2 SingleQuot)</td>
			<td>Lonza</td>
			<td>CC-4149</td>
			<td>Add to basal media before use</td>
		</tr>
		<tr>
			<td>0.5% Trypsin-EDTA 10X</td>
			<td>Gibco</td>
			<td>15400-054</td>
			<td>Dilute to 2X with sterile dPBS</td>
		</tr>
		<tr>
			<td>Hemocytometer</td>
			<td>VWR</td>
			<td>15170-208</td>
			<td></td>
		</tr>
		<tr>
			<td>large petri dishes for SMC Plating (150mm)</td>
			<td>Corning</td>
			<td>353025</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine gelatin</td>
			<td>Sigma</td>
			<td>G-9382</td>
			<td></td>
		</tr>
		<tr>
			<td>Human fibronectin</td>
			<td>Corning</td>
			<td>356008</td>
			<td>5&#181;g/ml of fibronectin and store at -20</td>
		</tr>
		<tr>
			<td>10X Hanks&#39; Buffered salt solution (HBSS)</td>
			<td>Gibco</td>
			<td>14065-056</td>
			<td>Dilute to 1x with sterile water</td>
		</tr>
		<tr>
			<td>10X Dulbecco&#39;s Phosphate buffered saline (dPBS)</td>
			<td>Gibco</td>
			<td>14200-075</td>
			<td>Dilute to 1x with sterile water</td>
		</tr>
		<tr>
			<td>Disposable Curved Scalpel (30mm cutting edge) with handle</td>
			<td>Amazon</td>
			<td>MDS15210</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Fisher</td>
			<td>17-467-201</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell lifter</td>
			<td>Corning</td>
			<td>3008</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell scraper</td>
			<td>BD Falcon</td>
			<td>353085</td>
			<td></td>
		</tr>
		<tr>
			<td>50mL conical tube</td>
			<td>Corning</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>10mm petri dishes</td>
			<td>Falcon</td>
			<td>35-1008</td>
			<td></td>
		</tr>
		<tr>
			<td>Rneasy mini kit</td>
			<td>Qiagen</td>
			<td>74104</td>
			<td></td>
		</tr>
		<tr>
			<td>RNA lysis reagent</td>
			<td>Qiagen</td>
			<td>79306</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma</td>
			<td>158127</td>
			<td>Make 4% solution with dPBS</td>
		</tr>
		<tr>
			<td>70% ethanol</td>
			<td>Fisher</td>
			<td>04-355-305</td>
			<td></td>
		</tr>
		<tr>
			<td>Cavicide disinfectant</td>
			<td>Fisher</td>
			<td>131000</td>
			<td></td>
		</tr>
		<tr>
			<td>RIPA protein lysis buffer</td>
			<td>Sigma</td>
			<td>R0278</td>
			<td></td>
		</tr>
		<tr>
			<td>Sonic dismembranator</td>
			<td>Fisher</td>
			<td>Model 50</td>
			<td></td>
		</tr>
		<tr>
			<td>1X PBS for harvesting EC/SMC and immunocytochemistry</td>
			<td>Gibco</td>
			<td>10010023</td>
			<td>does not need to be sterile</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Embedding/Sectioning/Staining</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffin</td>
			<td>Fisher</td>
			<td>P31-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Automated tissue processer</td>
			<td>Excelsior</td>
			<td>ES</td>
			<td></td>
		</tr>
		<tr>
			<td>Embedding station + cold plate</td>
			<td>Leica</td>
			<td>HistoCore Arcadia</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Tek Accu-Edge Microtome blade</td>
			<td>VWR</td>
			<td>25608-961 or 25608-964</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>Thermo Fisher</td>
			<td>22-230-900</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>Thermo Fisher</td>
			<td>12-548-5E</td>
			<td></td>
		</tr>
		<tr>
			<td>Prolong Gold mounting medium</td>
			<td>Thermo Fisher</td>
			<td>P36931</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Other Solutions</td>
			<td></td>
			<td></td>
			<td>**Perform all steps in sterile conditions if using the solution on live cells</td>
		</tr>
		<tr>
			<td>Sterile dPBS</td>
			<td></td>
			<td></td>
			<td>Autoclave water to sterilize. Using the sterile water make DPBS (450mL sterile water: 50mL 10X DPBS), mix and filter sterilize (0.2&#181;m filter). 
			Final concentration: 1X</td>
		</tr>
		<tr>
			<td>sterile HBSS</td>
			<td></td>
			<td></td>
			<td>Autoclave water to sterilize. Using the sterile water make HBSS (450mL sterile water: 50mL 10X HBSS), mix and filter sterilize (0.2&#181;m filter). 
			Final concentration: 1X</td>
		</tr>
		<tr>
			<td>Fibronectin stock solution</td>
			<td></td>
			<td></td>
			<td>Use 50mL of sterile water to fibronectin to make stock of 1mL aliquots, then freeze until needed 
			Final concentration: 100&#181;g/mL</td>
		</tr>
		<tr>
			<td>Fibronectin to coat SMC side of VCCC</td>
			<td></td>
			<td></td>
			<td>thaw one aliquot and add 1mL of fibronectin (100&#181;g/mL) to 19mLs of 1X HBSS to reach a final concentration of 5&#181;g/mL. 
			Final concentration: 5&#181;g/mL</td>
		</tr>
		<tr>
			<td>2x Trypsin EDTA solution</td>
			<td></td>
			<td></td>
			<td>Dilute the 10X 0.5% Trypsin-EDTA to 2X with 1X dPBS (40mL DPBS: 10mL 10X Trypsin) . 
			Final concentration: 2X</td>
		</tr>
		<tr>
			<td>Blocking/antibody solution</td>
			<td></td>
			<td></td>
			<td>(9mL 1X PBS, 25mL Triton X100, 50mg bovine serum albumin, 500mL normal serum)</td>
		</tr>
		<tr>
			<td>Bovine gelatin to coat EC VCCC</td>
			<td></td>
			<td></td>
			<td>Mix 0.5g bovine gelatin into 100mL distilled water. Autoclave prior to first use and store at room temperature. Keep in sterile conditions. 
			Final concentration: 0.50%</td>
		</tr>
		<tr>
			<td>1X PBS for harvesting cells</td>
			<td></td>
			<td></td>
			<td>does not need to be sterile</td>
		</tr>
	</tbody>
</table>

a cell culture model of resistance arteries

The myoendothelial junction (MEJ), a unique signaling microdomain in small diameter resistance arteries, exhibits localization of specific proteins and signaling processes that can control vascular tone and blood pressure. As it is a projection from either the endothelial or smooth muscle cell, and due to its small size (on average, an area of ~1 &#181;m2), the MEJ is difficult to study in isolation. However, we have developed a cell culture model called the vascular cell co-culture (VCCC) that allows for in vitro MEJ formation, endothelial cell polarization, and dissection of signaling proteins and processes in the vascular wall of resistance arteries. The VCCC has a multitude of applications and can be adapted to suit different cell types. The model consists of two cell types grown on opposite sides of a filter with 0.4 &#181;m pores in which the in vitro MEJs can form. Here we describe how to create the VCCC via plating of cells and isolation of endothelial, MEJ, and smooth muscle fractions, which can then be used for protein isolation or activity assays. The filter with intact cell layers can be fixed, embedded, and sectioned for immunofluorescent analysis. Importantly, many of the discoveries from this model have been confirmed using intact resistance arteries, underscoring its physiological relevance.

The filter inserts used for the VCCC have small, 0.4 &#181;m holes. Figure 1A, an en face view of the VCCC filter inset, shows the pores in which the MEJ can form in vitro. The schematic depicts the smooth muscle cells plated on the lower side of the filter one day before the endothelial cells are plated on the opposite side (Figure 1B). Once the cells are plated, the EC, MEJ, and SMC fractions can be isolated t...

Watch this Scientific Journal Video about A Cell Culture Model of Resistance Arteries at JoVE.com

A Cell Culture Model of Resistance Arteries

A cell culture model of resistance arteries is described, allowing for the dissection of signaling pathways in endothelium, smooth muscle, or between endothelium and smooth muscle (the myoendothelial junction). The selective application of agonists or protein isolation, electron microscopy, or immunofluorescence can be utilized using this cell culture model.

The myoendothelial junction (MEJ), a unique signaling microdomain in small diameter resistance arteries, exhibits localization of specific proteins and ...

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7.8K Views.  University of Virginia School of Medicine. A cell culture model of resistance arteries is described, allowing for the dissection of signaling pathways in endothelium, smooth muscle, or between endothelium and smooth muscle (the myoendothelial junction). The selective application of agonists or protein isolation, electron microscopy, or immunofluorescence can be utilized using this cell culture model.

Video: A Cell Culture Model of Resistance Arteries

A Mouse Model of in Utero Transplantation

The transplantation of stem cells and viruses in utero has tremendous potential for treating congenital disorders in the human fetus. For example, in utero transplantation (IUT) of hematopoietic stem cells has been used to successfully treat patients with severe combined immunodeficiency.1,2 In several other conditions, however, IUT has been attempted without success.3 Given these mixed results, the availability of an efficient non-human model to study the biological sequelae of stem cell transplantation and gene therapy is critical to advance this field. We and others have used the mouse model of IUT to study factors affecting successful engraftment of in utero transplanted hematopoietic stem cells in both wild-type mice4-7 and those with genetic diseases.8,9 The fetal environment also offers considerable advantages for the success of in utero gene therapy. For example, the delivery of adenoviral10, adeno-associated viral10, retroviral11, and lentiviral vectors12,13 into the fetus has resulted in the transduction of multiple organs distant from the site of injection with long-term gene expression. in utero gene therapy may therefore be considered as a possible treatment strategy for single gene disorders such as muscular dystrophy or cystic fibrosis. Another potential advantage of IUT is the ability to induce immune tolerance to a specific antigen. As seen in mice with hemophilia, the introduction of Factor IX early in development results in tolerance to this protein.14 
In addition to its use in investigating potential human therapies, the mouse model of IUT can be a powerful tool to study basic questions in developmental and stem cell biology. For example, one can deliver various small molecules to induce or inhibit specific gene expression at defined gestational stages and manipulate developmental pathways. The impact of these alterations can be assessed at various timepoints after the initial transplantation. Furthermore, one can transplant pluripotent or lineage specific progenitor cells into the fetal environment to study stem cell differentiation in a non-irradiated and unperturbed host environment.
The mouse model of IUT has already provided numerous insights within the fields of immunology, and developmental and stem cell biology. In this video-based protocol, we describe a step-by-step approach to performing IUT in mouse fetuses and outline the critical steps and potential pitfalls of this technique.

A Mouse Model of in Utero Transplantation

The mouse model of in utero transplantation is a versatile tool that can be used to study the potential clinical applications of stem cell transplantation and gene therapy in the fetus. In this protocol, we present a general approach to performing this technique

The transplantation of stem cells and viruses in utero has tremendous potential for treating congenital disorders in the human fetus.  For example, in ...

Multi-photon Imaging of Tumor Cell Invasion in an Orthotopic Mouse Model of Oral Squamous Cell Carcinoma

Loco-regional invasion of head and neck cancer is linked to metastatic risk and presents a difficult challenge in designing and implementing patient management strategies. Orthotopic mouse models of oral cancer have been developed to facilitate the study of factors that impact invasion and serve as model system for evaluating anti-tumor therapeutics. In these systems, visualization of disseminated tumor cells within oral cavity tissues has typically been conducted by either conventional histology or with in vivo bioluminescent methods. A primary drawback of these techniques is the inherent inability to accurately visualize and quantify early tumor cell invasion arising from the primary site in three dimensions. Here we describe a protocol that combines an established model for squamous cell carcinoma of the tongue (SCOT) with two-photon imaging to allow multi-vectorial visualization of lingual tumor spread. The OSC-19 head and neck tumor cell line was stably engineered to express the F-actin binding peptide LifeAct fused to the mCherry fluorescent protein (LifeAct-mCherry). Fox1nu/nu mice injected with these cells reliably form tumors that allow the tongue to be visualized by ex-vivo application of two-photon microscopy. This technique allows for the orthotopic visualization of the tumor mass and locally invading cells in excised tongues without disruption of the regional tumor microenvironment. In addition, this system allows for the quantification of tumor cell invasion by calculating distances that invaded cells move from the primary tumor site. Overall this procedure provides an enhanced model system for analyzing factors that contribute to SCOT invasion and therapeutic treatments tailored to prevent local invasion and distant metastatic spread. This method also has the potential to be ultimately combined with other imaging modalities in an in vivo setting.

A comprehensive overview of the techniques involved in generating a mouse model of oral cancer and quantitative monitoring of tumor invasion within the tongue through multi-photon microscopy of labeled cells is presented. This system can serve as a useful platform for the molecular assessment and drug efficacy of anti-invasive compounds.

Loco-regional invasion of head and neck cancer is linked to metastatic risk and presents a difficult challenge in designing and implementing patient ...

Processing of Human Reduction Mammoplasty and Mastectomy Tissues for Cell Culture

Experimental examination of normal human mammary epithelial cell (HMEC) behavior, and how normal cells acquire abnormal properties, can be facilitated by in vitro culture systems that more accurately model in vivo biology. The use of human derived material for studying cellular differentiation, aging, senescence, and immortalization is particularly advantageous given the many significant molecular differences in these properties between human and commonly utilized rodent cells1-2. Mammary cells present a convenient model system because large quantities of normal and abnormal tissues are available due to the frequency of reduction mammoplasty and mastectomy surgeries.
The mammary gland consists of a complex admixture of many distinct cell types, e.g., epithelial, adipose, mesenchymal, endothelial. The epithelial cells are responsible for the differentiated mammary function of lactation, and are also the origin of the vast majority of human breast cancers. We have developed methods to process mammary gland surgical discard tissues into pure epithelial components as well as mesenchymal cells3. The processed material can be stored frozen indefinitely, or initiated into primary culture. Surgical discard material is transported to the laboratory and manually dissected to enrich for epithelial containing tissue. Subsequent digestion of the dissected tissue using collagenase and hyaluronidase strips stromal material from the epithelia at the basement membrane. The resulting small pieces of the epithelial tree (organoids) can be separated from the digested stroma by sequential filtration on membranes of fixed pore size. Depending upon pore size, fractions can be obtained consisting of larger ductal/alveolar pieces, smaller alveolar clusters, or stromal cells. We have observed superior growth when cultures are initiated as organoids rather than as dissociated single cells. Placement of organoids in culture using low-stress inducing media supports long-term growth of normal HMEC with markers of multiple lineage types (myoepithelial, luminal, progenitor)4-5. Sufficient numbers of cells can be obtained from one individual's tissue to allow extensive experimental examination using standardized cell batches, as well as interrogation using high throughput modalities.
Cultured HMEC have been employed in a wide variety of studies examining the normal processes governing growth, differentiation, aging, and senescence, and how these normal processes are altered during immortal and malignant transformation4-15,16. The effects of growth in the presence of extracellular matrix material, other cell types, and/or 3D culture can be compared with growth on plastic5,15. Cultured HMEC, starting with normal cells, provide an experimentally tractable system to examine factors that may propel or prevent human aging and carcinogenesis.

A method to process human mammary surgical discard material is described. Processed tissue, in the form of organoids, can be stored frozen indefinitely or placed in culture for long-term growth. This method enables experimental examination of normal human epithelial cell biology, and the effects of exogenous perturbations.

Experimental  examination of normal human mammary epithelial cell (HMEC) behavior, and how  normal cells acquire abnormal properties, can be facilitated ...

Videomorphometric Analysis of Hypoxic Pulmonary Vasoconstriction of Intra-pulmonary Arteries Using Murine Precision Cut Lung Slices

Acute alveolar hypoxia causes pulmonary vasoconstriction (HPV) - also known as von Euler-Liljestrand mechanism - which serves to match lung perfusion to ventilation. Up to now, the underlying mechanisms are not fully understood. The major vascular segment contributing to HPV is the intra-acinar artery. This vessel section is responsible for the blood supply of an individual acinus, which is defined as the portion of lung distal to a terminal bronchiole. Intra-acinar arteries are mostly located in that part of the lung that cannot be selectively reached by a number of commonly used techniques such as measurement of the pulmonary artery pressure in isolated perfused lungs or force recordings from dissected proximal pulmonary artery segments1,2. The analysis of subpleural vessels by real-time confocal laser scanning luminescence microscopy is limited to vessels with up to 50 &#181;m in diameter3.
We provide a technique to study HPV of murine intra-pulmonary arteries in the range of 20-100 &#181;m inner diameters. It is based on the videomorphometric analysis of cross-sectioned arteries in precision cut lung slices (PCLS). This method allows the quantitative measurement of vasoreactivity of small intra-acinar arteries with inner diameter between 20-40 &#181;m which are located at gussets of alveolar septa next to alveolar ducts and of larger pre-acinar arteries with inner diameters between 40-100 &#181;m which run adjacent to bronchi and bronchioles. In contrast to real-time imaging of subpleural vessels in anesthetized and ventilated mice, videomorphometric analysis of PCLS occurs under conditions free of shear stress. In our experimental model both arterial segments exhibit a monophasic HPV when exposed to medium gassed with 1% O2 and the response fades after 30-40 min at hypoxia.

Hypoxic pulmonary vasoconstriction (HPV) is an important physiological phenomenon by which at alveolar hypoxia lung perfusion is matched to ventilation. The major vascular segment contributing to HPV is the intra-acinar artery. Here, we describe our protocol for the analysis of HPV of murine pulmonary vessels with diameters of 20-100 &mu;m.

Acute alveolar hypoxia causes pulmonary vasoconstriction (HPV) - also known as von Euler-Liljestrand mechanism - which serves to match lung perfusion to ...

Assessing Myogenic Response and Vasoactivity In Resistance Mesenteric Arteries Using Pressure Myography

Small resistance arteries constrict and dilate respectively in response to increased or decreased intraluminal pressure; this phenomenon known as myogenic response is a key regulator of local blood flow. In isobaric conditions small resistance arteries develop sustained constriction known as myogenic tone (MT), which is a major determinant of systemic vascular resistance (SVR). Hence, ex vivo&#160;pressurized preparations of small resistance arteries are major tools to study microvascular function in near-physiological states. To achieve this, a freshly isolated intact segment of a small resistance artery (diameter ~260 &#956;m) is mounted onto two small glass cannulas and pressurized. These arterial preparations retain most in vivo characteristics and permit assessment of vascular tone in real-time. Here we provide a detailed protocol for assessing vasoactivity in pressurized small resistance mesenteric arteries from rats; these arteries develop sustained vasoconstriction - approximately 25% of maximal diameter - when pressurized at 70 mmHg. These arterial preparations may be used to study the effect of investigational compounds on relationship between intra-arterial pressure and vasoactivity and determine changes in microvascular function in animal models of various diseases.

Pressure myography is used to assess vasoactivity of small arteries that develop sustained constriction when pressurized. This manuscript provides a detailed protocol to assess in isolated segments of small mesenteric arteries from rats, vasoactivity and the effect of intraluminal pressure on vascular diameter.

Small resistance arteries constrict and dilate respectively in response to increased or decreased intraluminal pressure; this phenomenon known as myogenic ...

Assessment of Perigenital Sensitivity and Prostatic Mast Cell Activation in a Mouse Model of Neonatal Maternal Separation

Chronic prostatitis/chronic pelvic pain syndrome (CP/CPPS) has a lifetime prevalence of 14% and is the most common urological diagnosis for men under the age of 50, yet it is the least understood and studied chronic pelvic pain disorder. A significant subset of patients with chronic pelvic pain report having experienced early life stress or abuse, which can markedly affect the functioning and regulation of the hypothalamic-pituitary-adrenal (HPA) axis. Mast cell activation, which has been shown to be increased in both urine and expressed prostatic secretions of CP/CPPS patients, is partially regulated by downstream activation of the HPA axis. Neonatal maternal separation (NMS) has been used for over two decades to study the outcomes of early life stress in rodent models, including changes in the HPA axis and visceral sensitivity. Here we provide a detailed protocol for using NMS as a preclinical model of CP/CPPS in male C57BL/6 mice. We describe the methodology for performing NMS, assessing perigenital mechanical allodynia, and histological evidence of mast cell activation. We also provide evidence that early psychological stress can have long-lasting effects on the male urogenital system in mice.

We are measuring perigenital mechanical sensitivity and mast cell activation in the prostate of male C57BL/6 mice that underwent an early life stress paradigm &#8211; neonatal maternal separation, in order to induce a preclinical model of chronic prostatitis/chronic pelvic pain syndrome.

Chronic prostatitis/chronic pelvic pain syndrome (CP/CPPS) has a lifetime prevalence of 14% and is the most common urological diagnosis for men under the ...

A Human Glioblastoma Organotypic Slice Culture Model for Study of Tumor Cell Migration and Patient-specific Effects of Anti-Invasive Drugs

Glioblastoma (GBM) continues to carry an extremely poor clinical prognosis despite surgical, chemotherapeutic, and radiation therapy. Progressive tumor invasion into surrounding brain parenchyma represents an enduring therapeutic challenge. To develop anti-migration therapies for GBM, model systems that provide a physiologically relevant background for controlled experimentation are essential. Here, we present a protocol for generating slice cultures from human GBM tissue obtained during surgical resection. These cultures allow for ex vivo experimentation without passaging through animal xenografts or single cell cultures. Further, we describe the use of time-lapse laser scanning confocal microscopy in conjunction with cell tracking to quantitatively study the migratory behavior of tumor cells and associated response to therapeutics. Slices are reproducibly generated within 90 min of surgical tissue acquisition. Retrovirally-mediated fluorescent cell labeling, confocal imaging, and tumor cell migration analyses are subsequently completed within two weeks of culture. We have successfully used these slice cultures to uncover genetic factors associated with increased migratory behavior in human GBM. Further, we have validated the model&#39;s ability to detect patient-specific variation in response to anti-migration therapies. Moving forward, human GBM slice cultures are an attractive platform for rapid ex vivo assessment of tumor sensitivity to therapeutic agents, in order to advance personalized neuro-oncologic therapy.

Current ex vivo models of glioblastoma (GBM) are not optimized for physiologically relevant study of human tumor invasion. Here, we present a protocol for generation and maintenance of organotypic slice cultures from fresh human GBM tissue. A description of time-lapse microscopy and quantitative cell migration analysis techniques is provided.

Glioblastoma (GBM) continues to carry an extremely poor clinical prognosis despite surgical, chemotherapeutic, and radiation therapy. Progressive tumor ...

Isolation and Culture of Primary Endothelial Cells from Canine Arteries and Veins

Cardiovascular disease is studied in both human and veterinary medicine. Endothelial cells have been used extensively as an in vitro model to study vasculogenesis, (tumor) angiogenesis, and atherosclerosis. The current standard for in vitro research on human endothelial cells (ECs) is the use of Human Umbilical Vein Endothelial Cells (HUVECs) and Human Umbilical Artery Endothelial Cells (HUAECs). For canine endothelial research, only one cell line (CnAOEC) is available, which is derived from canine aortic endothelium. Although currently not completely understood, there is a difference between ECs originating from either arteries or veins. For a more direct approach to in vitro functionality studies on ECs, we describe a new method for isolating Canine Primary Endothelial Cells (CaPECs) from a variety of vessels. This technique reduces the chance of contamination with fast-growing cells such as fibroblasts and smooth muscle cells, a problem that is common in standard isolation methods such as flushing the vessel with enzymatic solutions or mincing the vessel prior to digestion of the tissue containing all cells. The technique we describe was optimized for the canine model, but can easily be utilized in other species such as human.

Novel isolation methods of primary endothelial cells from blood vessels are needed. This protocol describes a new technique that completely inverts blood vessels of interest, exposing only the endothelial side to enzymatic digestion. The resulting pure endothelial cell culture can be used to study cardiovascular diseases, disease modelling, and angiogenesis.

Cardiovascular disease is studied in both human and veterinary medicine. Endothelial cells have been used extensively as an in vitro model to study ...

Evaluation of Vascular Control Mechanisms Utilizing Video Microscopy of Isolated Resistance Arteries of Rats

This protocol describes the use of in vitro television microscopy to evaluate vascular function in isolated cerebral resistance arteries (and other vessels), and describes techniques for evaluating tissue perfusion using Laser Doppler Flowmetry (LDF) and microvessel density utilizing fluorescently labeled Griffonia simplicifolia (GS1) lectin. Current methods for studying isolated resistance arteries at transmural pressures encountered in vivo and in the absence of parenchymal cell influences provide a critical link between in vivo studies and information gained from molecular reductionist approaches that provide limited insight into integrative responses at the whole animal level. LDF and techniques to selectively identify arterioles and capillaries with fluorescently-labeled GS1 lectin provide practical solutions to enable investigators to extend the knowledge gained from studies of isolated resistance arteries. This paper describes the application of these techniques to gain fundamental knowledge of vascular physiology and pathology in the rat as a general experimental model, and in a variety of specialized genetically engineered &#34;designer&#34; rat strains that can provide important insight into the influence of specific genes on important vascular phenotypes. Utilizing these valuable experimental approaches in rat strains developed by selective breeding strategies and new technologies for producing gene knockout models in the rat, will expand the rigor of scientific premises developed in knockout mouse models and extend that knowledge to a more relevant animal model, with a well understood physiological background and suitability for physiological studies because of its larger size.

This manuscript describes in vitro video microscopy protocols for evaluating vascular function in rat cerebral resistance arteries. The manuscript also describes techniques for evaluating microvessel density with fluorescently labeled lectin and tissue perfusion using Laser Doppler Flowmetry.

This protocol describes the use of in vitro television microscopy to evaluate vascular function in isolated cerebral resistance arteries (and other ...

Muscle Imbalances: Testing and Training Functional Eccentric Hamstring Strength in Athletic Populations

Many hamstring injuries that occur during physical activity occur while the muscles are lengthening, during eccentric hamstring muscle actions. Opposite of these eccentric hamstring actions are concentric quadriceps actions, where the larger and likely stronger quadriceps straighten the knee. Therefore, to stabilize the lower limbs during movement, the hamstrings must eccentrically combat against the strong knee-straightening torque of the quadriceps. As such, eccentric hamstring strength expressed relative to concentric quadricep strength is commonly referred to as the &#34;functional ratio&#34; as most movements in sports require simultaneous concentric knee extension and eccentric knee flexion. To increase the strength, resiliency, and functional performance of the hamstrings, it is necessary to test and train the hamstrings at different eccentric speeds. The main purpose of this work is to provide instructions for measuring and interpreting eccentric hamstring strength. Techniques for measuring the functional ratio using isokinetic dynamometry are provided and sample data will be compared. Additionally, we briefly describe how to address hamstring strength deficiencies or unilateral strength differences using exercises that specifically focus on increasing eccentric hamstring strength.

The hamstrings are a group of muscles that are sometimes problematic for athletes, resulting in soft tissue injury in the lower limbs. To prevent such injuries, functional training of the hamstrings requires intensive eccentric contractions. Additionally, hamstring function should be tested in relation to quadricep function at different contraction speeds.

Many hamstring injuries that occur during physical activity occur while the muscles are lengthening, during eccentric hamstring muscle actions. Opposite ...