In loving memory of Rich West, a brilliant scientist, colleague, and dear friend.&#160;We would like to thank the University of Delaware Flow Cytometry and BioImaging Core as part of the Delaware Biotechnology Institute for their ongoing contributions to our studies. We would also like to thank Emma Hudgins for the careful review and editing of the manuscript. Our work is supported by the National Institute of General Medical Sciences P20GM113125-6564 (I.S. Fancher). This project was also supported by the Delaware INBRE program, with a grant from the NIGMS (P20GM103446) from the National Institutes of Health and the State of Delaware (I.S. Fancher), and a University of Delaware General University Research grant (I.S. Fancher). This content is solely the responsibility of the authors and does not necessarily represent the official views of NIH.

The use of animals in these studies was approved by the University of Delaware, Institutional Animal Care and Use Committee (#1372).
1. Tissue dissection and cleaning
NOTE: 10- to 12-week-old C57BL/6J mice are used in the video protocol. Please refer to Figure 1 for the schematic and the desired outcome of tissue dissection and cleaning arteries and refer to the Table of Materials for the list of supplies and manufacturer information.
<ol>
	<li>Euthanize the mouse by CO2 asphyxiation followed by cervical dislocation.</li>
	<li>Secure the mouse using dissection pins and spray the ventral surface with 70% ethanol.</li>
	<li>Isolation of subcutaneous adipose tissues located by each hindlimb 
	&#8203;NOTE: Dissecting tools needed: straight Bonn scissors (9 cm) and blunt, curved Graefe forceps.
	<ol>
		<li>Use forceps to lift the skin right above the genitals and make a small (2 mm) incision in the skin.</li>
		<li>Insert the tip of the scissors into the incision site and make a 4-5 cm midline incision, beginning at the initial incision and dissecting cranially to the base of the sternum. Be cautious to not penetrate the abdominal cavity.</li>
		<li>Make a 1 cm lateral incision, extending laterally from the midline immediately below the forelimb.</li>
		<li>Make a diagonal 1 cm incision between the hindlimb and genitals.</li>
		<li>Make small cuts along the midline skin incisions to peel away the associated connective tissue and expose the subcutaneous adipose depot. Pin the skin down.</li>
		<li>Identify the &#34;C-shaped&#34; subcutaneous adipose tissue and artery/vein that runs perpendicular to the abdominal cavity.</li>
		<li>Start from the bottom of the C shape near the genitals and gently grasp the adipose tissue to expose the connective tissues. Make small cuts in the connective tissue to separate the fat from the skin. Avoid disturbing the exposed vasculature.</li>
		<li>Continue dissecting away the connective tissue until the subcutaneous adipose tissue can be removed intact. Carefully separate the exposed artery from the surrounding connective tissue.</li>
		<li>Store the isolated subcutaneous adipose in HEPES buffer on ice until ready to isolate the vascular bed of interest.</li>
		<li>Repeat steps 1.3.3.-1.3.5. for the remaining posterior subcutaneous adipose depot.</li>
	</ol>
	</li>
	<li>Isolation of the mesenteric adipose depot 
	NOTE: Dissecting tools needed: straight Bonn scissors (9 cm), Graefe forceps, a pair of #5 forceps, straight iris scissors, and curved Bonn scissors (9 cm).
	<ol>
		<li>Use the Graefe forceps to lift the thin peritoneal cavity wall above the urinary bladder and make a small incision using straight scissors.</li>
		<li>Slowly lift the penetrated peritoneal cavity wall, carefully insert the straight iris scissors into the incision site, and make a 4-5 cm incision from the urinary bladder to the sternum.</li>
		<li>Make two 1 cm long horizontal incisions with the straight iris scissors, extending laterally from the midline to immediately above the hindlimb and below the forelimb. Use the Graefe forceps to peel back the abdominal musculature and expose the peritoneal viscera.</li>
		<li>Repeat step 1.4.3. on the opposite side.</li>
		<li>Use the pair of #5 forceps to lift the intestines out of the visceral cavity to reveal the mesenteric adipose.</li>
		<li>Use the curved scissors and #5 forceps to separate the adipose along the colon, starting from the caecum to where the colon descends from view.</li>
		<li>Return to the caecum and use the curved scissors and #5 forceps to separate the adipose along the small intestine to the pancreas. Remove a small part of the pancreas to completely isolate the mesenteric adipose. 
		&#8203;NOTE: Removing a small part of the pancreas along with the mesenteric adipose may help in cleaning the arteries as it provides stability during cleaning.</li>
		<li>Store the isolated mesenteric adipose in HEPES buffer on ice until ready to isolate the mesenteric arteries.</li>
	</ol>
	</li>
	<li>Isolation of the respective arteries from subcutaneous and mesenteric adipose tissues 
	NOTE: Dissecting tools needed: a dissecting dish and stereoscope with a light source, a pair of #55 forceps, and a pair of #5 forceps.
	<ol>
		<li>Place ~10 mL of cold HEPES buffer into the dissecting dish and transfer the adipose tissue to the dish using the #5 forceps.</li>
		<li>Use the stereoscope at 1x magnification to view and position the adipose tissue to expose the artery/vein pair(s) (Figure 2).</li>
		<li>Use the #55 forceps to carefully remove the parenchymal adipose from the artery. Remove small pieces of adipose at a time and carefully observe artery/vein pairs under the stereoscope to identify and avoid damaging the vasculature of interest. 
		NOTE: Adjust the magnification and light source accordingly to clean the arteries. Increase the magnification and light intensity to better view the arteries as adipose is removed. Fine cleaning of the arteries should be done under 4x-5x magnification. It is important to distinguish between connective tissues (collagens), veins, and arteries. Connective tissues and collagen fibers have a string-like appearance without branches, are striated, and have a whitish color. Unlike collagen fibers and connective tissues, arteries and veins are transparent and have multiple branches with a defined lumen. Small arteries and veins may appear similar in appearance when free of parenchymal tissue. The presence of blood in the lumen can further help one to identify the arteries from veins. Veins have thinner blood vessel walls and a larger lumen and contain more blood compared to comparably sized arteries.</li>
		<li>Repeat step 1.5.3. until the arteries are completely void of parenchymal adipose tissue.</li>
		<li>Use #5 forceps to transfer the cleaned arteries to a new tube with fresh HEPES buffer and keep on ice until ready for digestion.</li>
	</ol>
	</li>
</ol>
2. Digestion of isolated arteries for isolation of endothelial cells
NOTE: Please refer to Figure 2 for the schematic of tissue digestion and isolation of endothelial cells and the Table of Materials for a complete list of supplies needed for the protocol.
<ol>
	<li>Add 1 mg each of dispase and elastase to each 2 mL microcentrifuge tube. Add 2 mL of dissociation solution to each tube. Vortex and incubate at 37 &#176;C until completely dissolved. Use #5 forceps to transfer the cleaned arteries (step 1.5.5.) to the respective sample tubes. The final concentration of each enzyme is 0.5 mg/mL.</li>
	<li>Incubate at 37 &#176;C for 1 h with gentle agitation by inversion every 10-15 min, such that the arteries are suspended in solution to maintain consistent and uniform contact between the enzymes and arteries.</li>
	<li>Weigh 1 mg of collagenase type I per sample and add it to new tubes.</li>
	<li>Allow the arteries to settle at the bottom of the tube. Remove 500 &#181;L of the buffer from the top without disturbing the arteries and transfer it into designated tubes containing collagenase. Vortex and return the solution to the respective samples in the original sample tubes. The final concentration of collagenase type I is 0.5 mg/mL.</li>
	<li>Incubate at 37&#160;&#176;C for 15 min. Ensure that the arteries separate into pieces following brief shaking of the sample tube. If the arteries do not separate into pieces following perturbation, incubate in 5 min increments until the arterial breakdown is visible.</li>
	<li>Using a glass pipette, vigorously triturate the digested tissue 10x-15x to mechanically dissociate the cells.</li>
	<li>Pass the solution and digested tissue through a 70 &#181;m cell strainer or a flow cytometry tube strainer to obtain single-cell suspensions.</li>
</ol>
3. Staining of endothelial cells prior to flow cytometry
NOTE: The staining and processing of endothelial cells are carried out on ice to improve cell viability, unless instructed. The 5 mL polystyrene round-bottom tubes are centrifuged at 1,163 x g for 3 min at room temperature (RT).
<ol>
	<li>Centrifuge the single-cell suspensions at 1,163 x g for 3 min at RT.</li>
	<li>Remove the supernatant and resuspend the cell pellet in 1 mL of PBS + 5% BSA for 30 min at RT. 
	NOTE: Researchers should consider blocking FC receptors depending on the cell type, target of interest, and antibody used18,19.</li>
	<li>Add 1 &#181;L of cell viability stain (405 nm excitation) (Table of Materials) and incubate in the dark for 30 min at RT.</li>
	<li>Centrifuge the cell suspension at 1,163 x g for 3 min at RT. Resuspend the cell pellet in 200 &#181;L of PBS + 1% BSA.</li>
	<li>To the samples, add primary antibodies conjugated to CD31 (CD31-PE, 0.75 &#181;g/sample) and CD45 (CD45-FITC, 2.5 &#181;g/sample) and incubate on a rocker in the fridge for 15 min after covering the samples with aluminum foil. 
	NOTE : To identify and/or obtain a pure population of endothelial cells, probe for both CD31 and CD45 using CD31- and CD45-conjugated primary antibodies with distinct excitation/emission fluorescence and gate/sort cells with a CD31+CD45&#8722; expression profile. Compensation may be required depending on the fluorophores used18,20,21.</li>
	<li>Wash the cells by adding 1 mL of PBS + 1% BSA directly to the cell suspension.</li>
	<li>Centrifuge the cell suspension at 1,163 x g for 3 min at RT. Remove the supernatant and resuspend in 200 &#181;L of 1% formaldehyde with 0.1% BSA. Store in the fridge covered with aluminum foil until ready for flow cytometry. 
	NOTE: Cell yields are improved by reducing the wash and centrifugation steps. These may need to be altered depending on the outcome approach. The sample in fixative should be analyzed within 24 h.</li>
</ol>

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The authors have no conflicts of interest.

Endothelial dysfunction is a precursor to severe disease states, likely driving the development of atherosclerosis, hypertension, and stroke3,23. While the identified mechanisms underlying endothelial dysfunction in a given pathological condition are many, distinct vascular beds are likely to be differentially influenced by pathological conditions4,24. Furthermore, different cardiovascular risk factors (e.g., obesity, hypertension, dyslipidemia, smoking, diabetes) induce dysfunction through a variety of distinct mechanisms23,25. Therefore, it is critical to isolate endothelial cell populations from established animal models of disease or accessible human tissue and perform assays on cells immediately removed from the in vivo environment. Isolating cells in this way has a unique advantage over studying cells in culture in that they are void of culture-induced phenotypic changes5,6. Moreover, including a heterogenous endothelial cell population, as observed in vivo26,27 (and that can be further separated using flow-assisted cell sorting), from a live organism better informs the in vivo environment. Finally, this method is applicable to the investigation of numerous animal models and potentially human tissue and can be used to generate primary cell culture lines, if such a need is warranted.
The critical step in this protocol, and the step that will need most adjusting for different vascular beds or cell types not presented here, is the digestion of vascular tissue. This step must be optimized for cell health without diminishing cell yield. The specific enzymes used and the duration for digestion are critical to optimizing cell health and yield to be able to adequately perform downstream assays. For the identification of membrane expression of CD36, as detected by flow cytometry in subcutaneous and mesenteric endothelial cells (Figure 4), a modified version of the digestion protocol originally designed for patch-clamp electrophysiology28 was developed. This included an extension of the collagenase I digestion time to 30 min to increase cell yields to better meet the demands of flow cytometry vs. what is required for patch-clamp studies. As this modification may impact cell health to some extent, a cell viability stain was used in the flow cytometry analyses to ensure that only viable endothelial cells were assessed following this protocol (Figure 3). It is recommended that studies aimed at isolating cells from vascular tissue should include a marker for cell viability prior to assessing the desired approach.
The described protocol is sufficient to liberate viable endothelial cells for analysis and downstream applications from &#8804;1 mg arterial samples derived from mice; however, if the arteries of interest were to be isolated from different tissues or organisms (e.g., humans) that would result in significantly different arterial masses, one would have to optimize the digestion enzyme content and the duration of incubation to efficiently isolate the cell population of interest. For some vascular beds that begin with even smaller amounts of starting tissue than the subcutaneous and mesenteric beds presented here (e.g., coronary arteries), pooling of arteries from several mice may be necessary per sample. Indeed, this may be a necessary step for any given vascular bed given that the outcome approach requires a relatively high cell yield. A major limitation is that, due to the nature of variations per sample digestion, cell yields can sometimes be significantly different across batches even when from the same vascular bed. This can cause issues when analyzing data and should be accounted for via normalization when appropriate. For instance, CD36 expression was extremely variable in the raw data due to alterations in cell yields across individual samples of subcutaneous and mesenteric adipose arteries. Therefore, raw data was normalized to CD31+CD45&#8722; mean fluorescence intensity with the assumption that this method would correct for batch differences (Figure 4). Of course, depending on the approach, more sophisticated statistical analyses and normalization methods may be required.
In summary, this paper presents a method to dissect, isolate, and digest subcutaneous and mesenteric arteries from mice to investigate the expression and/or function of endothelial targets of interest. With modifications to the presented protocol, different vascular beds and cell types (e.g., smooth muscle cells) can be investigated. This protocol, as a foundation for a plethora of available experimental approaches, has the potential to advance the understanding of vascular cell biology and mechanisms of vascular dysfunction.

Endothelial cells are well recognized for their important roles in a variety of physiological processes, including barrier functions, angiogenesis, and vascular tone regulation1,2. Although endothelial cell dysfunction is well documented in promoting atherosclerosis, hypertension, diabetes, etc., the underlying mechanisms driving endothelial dysfunction remain poorly understood and likely differ between distinct vascular beds2,3,4. The effort to unravel these pathological mechanisms of endothelial cell dysfunction is challenged by the limited access to a pure population of endothelial cells from tissues and/or the phenotypic changes of endothelial cells in culture5,6. Therefore, being able to isolate and perform analyses on endothelial cells from various vascular beds in their native form will give insight into the processes of cardiovascular disease.
This protocol presents the procedure to dissect subcutaneous and mesenteric adipose tissues from mice, followed by isolation of their respective arterial vasculature. Functionally viable endothelial cells lining the arterial walls are liberated using a specific cocktail of enzymes. Special care was taken to optimize the conditions of the digestion protocol to obtain a sufficient yield of endothelial cells from &#60;1 mg of starting tissue while keeping biomolecular markers intact for analysis.&#160;Isolated endothelial cells are next identified using flow cytometry. The presence of CD31 (PECAM) is used to primarily identify endothelial cells. Due to the expression of CD31 in other cell types, including several of hematopoietic origin that also express CD45, the purity of the isolated endothelial cells was further enhanced by the exclusion of cells expressing both CD31 and CD455,6,7,8. Moreover, depending on the research question and the downstream applications to be employed, researchers should consider selecting an extensive panel of positive and negative selection markers to optimize the purity of the cell population of interest.
Though the technique of dissecting and isolating adipose depot arteries has been used in the previous publications9,10,11,12,13, a detailed protocol describing the isolation of embedded arteries has yet to be presented. Demonstrating the technique for the isolation and digestion of arteries from different vascular beds from a given species of interest will provide options for researchers to evaluate freshly isolated vascular cells&#160;from arteries of interest and allow them to perform a wide range of functional tests on specific cell types. Tests may include but are not limited to flow cytometry for cell sorting and membrane protein expression5,8, electrophysiology for ion channel activity9, molecular profiling (proteomics/genomic analyses, etc.)14,15, and the generation of primary cell lines for drug screening in vitro16,17.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Tissue dissection tools</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>5 forceps (2)</td>
			<td>Fine Science Tools</td>
			<td>11252-00</td>
			<td>For isolation of mesenteric adipose depot and arteries transfer</td>
		</tr>
		<tr>
			<td>55 forceps (2)</td>
			<td>Fine Science Tools</td>
			<td>11295-51</td>
			<td>For parenchymal adipose removal</td>
		</tr>
		<tr>
			<td>Curved Bonn scissors</td>
			<td>Fine Science Tools</td>
			<td>14061-10</td>
			<td>For isolation of mesenteric adipose depot</td>
		</tr>
		<tr>
			<td>Graefe forceps</td>
			<td>Fine Science Tools</td>
			<td>11051-10</td>
			<td>For general dissection steps and isolation of subcutaneous adipose depot</td>
		</tr>
		<tr>
			<td>Straight Bonn scissor</td>
			<td>Fine Science Tools</td>
			<td>14060-09</td>
			<td>For general dissection steps and isolation of subcutaneous adipose depot</td>
		</tr>
		<tr>
			<td>Stereoscope with light source</td>
			<td>Laxco</td>
			<td>Z230PT40</td>
			<td>For parenchymal adipose removal and artery isolation</td>
		</tr>
		<tr>
			<td>Digestive enzymes for Artery Digestion</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase Type I</td>
			<td>Wothington-BioChem</td>
			<td>LS004194</td>
			<td></td>
		</tr>
		<tr>
			<td>Dispase (Neutral Protease)</td>
			<td>Wothington-BioChem</td>
			<td>LS02110</td>
			<td></td>
		</tr>
		<tr>
			<td>Elastase</td>
			<td>Wothington-BioChem</td>
			<td>LS002292</td>
			<td></td>
		</tr>
		<tr>
			<td>Solutions Recipes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES Buffer, pH 7.4</td>
			<td></td>
			<td></td>
			<td>Final concentration</td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>J.T.Baker</td>
			<td>1332-1</td>
			<td>2 mM</td>
		</tr>
		<tr>
			<td>Dextrose (D-glucose) anhydrous</td>
			<td>Fisher</td>
			<td>D16-500</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Fisher</td>
			<td>BP310-500</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>Magnesium Chloride</td>
			<td>Fisher</td>
			<td>M33-500</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>Potassium Chloride</td>
			<td>Fisher</td>
			<td>P217-500</td>
			<td>5 mM</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Oxoid</td>
			<td>LP0005</td>
			<td>145 mM</td>
		</tr>
		<tr>
			<td>Dissociation Solution, pH 7.30-7.40</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dextrose (D-glucose) anhydrous</td>
			<td>Fisher</td>
			<td>D16-500</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Fisher</td>
			<td>BP310-500</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>Magnesium Chloride</td>
			<td>Fisher</td>
			<td>M33-500</td>
			<td>2 mM</td>
		</tr>
		<tr>
			<td>Potassium Chloride</td>
			<td>Fisher</td>
			<td>P217-500</td>
			<td>56 mM</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Oxoid</td>
			<td>LP0005</td>
			<td>55 mM</td>
		</tr>
		<tr>
			<td>Sodium L-Glutamate monohydrate</td>
			<td>TCI</td>
			<td>G0188</td>
			<td>80 mM</td>
		</tr>
		<tr>
			<td>Flow Cytometry Analyses</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL Polystyrene round-bottom tube with cell-strainer cap</td>
			<td>Falcon</td>
			<td>352235</td>
			<td></td>
		</tr>
		<tr>
			<td>CD31-PE</td>
			<td>Miltenyi Biotec</td>
			<td>130-119-653</td>
			<td>0.75&#181;g/sample</td>
		</tr>
		<tr>
			<td>CD36-APC</td>
			<td>R&#38;D Systems</td>
			<td>AF2519</td>
			<td>2.5&#181;g/ sample</td>
		</tr>
		<tr>
			<td>CD45-FITC</td>
			<td>BioLegend</td>
			<td>103108</td>
			<td>2.5&#181;g/ sample</td>
		</tr>
		<tr>
			<td>Live/Dead Fixable Violet Dead Cell Staining kit (cell viability stain)</td>
			<td>Invitrogen</td>
			<td>L34955</td>
			<td>1&#181;L/sample</td>
		</tr>
	</tbody>
</table>

isolation and identification of vascular endothelial cells from distinct adipose depots for downstream applications

Vascular endothelial cells lining the wall of the vascular system play important roles in a variety of physiological processes, including vascular tone regulation, barrier functions, and angiogenesis. Endothelial cell dysfunction is a hallmark predictor and major driver for the progression of severe cardiovascular diseases, yet the underlying mechanisms remain poorly understood. The ability to isolate and perform analyses on endothelial cells from various vascular beds in their native form will give insight into the processes of cardiovascular disease. This protocol presents the procedure for the dissection of mouse subcutaneous and mesenteric adipose tissues, followed by isolation of their respective arterial vasculature. The isolated arteries are then digested using a specific cocktail of digestive enzymes focused on liberating functionally viable endothelial cells. The digested tissue is assessed by flow cytometry analysis using CD31+/CD45&#8722; cells as markers for positive endothelial cell identification. Cells can be sorted for immediate downstream functional assays or used to generate primary cell lines. The technique of isolating and digesting arteries from different vascular beds will provide options for researchers to evaluate freshly isolated vascular cells&#160;from arteries of interest and allow them to perform a wide range of functional tests on specific cell types.

A schematic of the workflow is shown in Figure 1. The schematic highlights the protocol steps described in more detail in the protocol text. Figure 2 shows pictures of subcutaneous adipose (Figure 2A, left) following dissection and a subcutaneous artery arcade (Figure 2A, right) following cleaning of the parenchymal tissue. Figure 2 also shows the mesenteric adipose (Figure 2B, left) following dissection and the mesenteric arterial arcade (Figure 2B, right) following cleaning of the parenchymal tissue.
The protocol presented here is designed to assist researchers potentially interested in a) comparing distinct adipose depot vasculature beds in basic science approaches and/or disease models, b) the digestion of vascular beds prior to isolation of vascular cells of interest for downstream application, and c) the use of flow cytometry to identify vascular cells of interest (Figure 3) for a variety of applications including, but not limited to, protein expression analyses as performed here (Figure 4). Figure 3 details an example of our approach to identify endothelial cells from digested mouse arteries using flow cytometry. Cell preparations obtained from digested subcutaneous (Figure 3A) or mesenteric (Figure 3B) adipose arteries were stained with CD31-PE, CD45-FITC, and a cell viability dye prior to the fixation and identification of viable CD31+CD45&#8722; endothelial cells using flow cytometry, as similarly performed elsewhere8. The cell viability stain allowed for the identification of only those viable cells that survived the isolation and digestion protocol prior to fixation. The use of this method will allow researchers to assess the expression or function of viable vascular cells of interest.
To determine if endothelial cells isolated from distinct adipose depot vasculature exhibited differences in membrane protein expression, isolated cells were probed for the fatty acid translocase, CD36 (Figure 4), as this membrane protein was recently shown to be essential in the endothelial-mediated distribution of fatty acids to tissues22. Therefore, determining the relative expression differences between membrane CD36, for example, in distinct vascular beds may a) support existing data regarding differential preference for fatty acid utilization among different tissues and/or b) unveil potential novel differences in tissue distribution of fatty acids in health and disease. From the same preparations of digested vascular cells exemplified in Figure 3, CD31+CD45&#8722;CD36+ endothelial cells were identified in subcutaneous (Figure 4A) and mesenteric (Figure 4B) adipose, and CD36 expression was quantified in this population in each vascular bed. Figure 4C and Figure 4D reveal that both the percentage of endothelial cells expressing CD36 and the intensity of CD36 expression were greater in subcutaneous endothelial cells compared to that observed in mesenteric endothelial cells. These preliminary findings support the use of our methodology to identify distinct differences between vascular beds.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63999/63999fig01.jpg" /> 
Figure 1: Working scheme to isolate endothelial cells from subcutaneous and visceral adipose tissues for downstream applications. (A)10-12-week-old C57BL/6J mice were euthanized and used to demonstrate this procedure. (Bi) First, the subcutaneous adipose is removed. (Bii) Then, the mesenteric (visceral) adipose tissue is subsequently removed. The white dashed lines indicate the locations of the subcutaneous (left) and mesenteric (right) adipose depots following the dissection and removal. The respective arteries are isolated for downstream applications. (C) In this protocol, isolated arteries are next digested using a specific enzyme cocktail as a first step in liberating functionally viable endothelial cells. (D) Isolated endothelial cells are identified by probing for CD31+ and CD45&#8722; cells using conjugated antibodies prior to flow cytometry analysis. Cells with an expression profile of CD31+/CD45&#8722; are the endothelial cells of interest for additional analyses. <a href="https://www.jove.com/files/ftp_upload/63999/63999fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63999/63999fig02.jpg" /> 
Figure 2: Isolated subcutaneous and visceral adipose tissues and respective arteries. (A) Left: Isolated &#34;C-shaped&#34; subcutaneous adipose tissue from hindlimbs. A major branch of the subcutaneous adipose artery, denoted by arrow (1), is revealed when parenchymal adipose is removed. Right: Isolated subcutaneous adipose arteries. (B) Left: The mesenteric (visceral) adipose tissue is isolated from the gut. Right: The respective arterial arcade is isolated by removing the parenchymal tissue. Different scales are used between the pictures of adipose (5 mm) and arteries (1 mm). <a href="https://www.jove.com/files/ftp_upload/63999/63999fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63999/63999fig03.jpg" /> 
Figure 3: Identification of endothelial cells following arterial tissue digestion via flow cytometry. Representative plots showing cells liberated from isolated (A) subcutaneous or (B) mesenteric adipose arteries. Cells were exposed to conjugated antibodies that targeted extracellular epitopes of CD31 (PE) and CD45 (FITC) prior to fixation. Flow cytometry was used to identify the CD31+CD45&#8722; endothelial cell population. A cell viability stain was used to select only the cells that survived the isolation and digestion protocols for subsequent analyses. In addition, appropriate gating at this stage will further separate an intended cell population from contaminating cells should the size of the cell type of interest be known. <a href="https://www.jove.com/files/ftp_upload/63999/63999fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63999/63999fig04.jpg" /> 
Figure 4: Analysis of CD36 membrane expression in subcutaneous vs. mesenteric adipose artery endothelial cells via flow cytometry. Representative population plots and histograms showing endothelial CD36 (APC) membrane expression in (A) subcutaneous or (B) mesenteric adipose arteries. (C) Percentage of endothelial cells (ECs) expressing CD36 in subcutaneous vs. mesenteric ECs (n = 5, 3 males, 2 females; *p &#60; 0.05 following Student&#39;s t-test). (D) Normalized EC membrane CD36 expression in subcutaneous vs. mesenteric adipose arteries (n = 5, 3 males, 2 females; *p &#60; 0.05 following Student&#39;s t-test). <a href="https://www.jove.com/files/ftp_upload/63999/63999fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Isolation and Identification of Vascular Endothelial Cells from Distinct Adipose Depots for Downstream Applications at JoVE.com

Isolation and Identification of Vascular Endothelial Cells from Distinct Adipose Depots for Downstream Applications

This protocol details a method for the dissection of mouse adipose depots and the isolation and digestion of respective arteries to liberate and then identify the endothelial cell population. Freshly isolated cells used in downstream applications will advance the understanding of vascular cell biology and the mechanisms of vascular dysfunction.

Vascular endothelial cells lining the wall of the vascular system play important roles in a variety of physiological processes, including vascular tone ...

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2.6K Views.  University of Delaware. This protocol details a method for the dissection of mouse adipose depots and the isolation and digestion of respective arteries to liberate and then identify the endothelial cell population. Freshly isolated cells used in downstream applications will advance the understanding of vascular cell biology and the mechanisms of vascular dysfunction.

Video: Isolation and Identification of Vascular Endothelial Cells from Distinct Adipose Depots for Downstream Applications

Isolation and Culture of Pulmonary Endothelial Cells from Neonatal Mice

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells (HUVECs) have shown to be convenient, easy to obtain and culture, and thus are the most widely studied endothelial cells. However, for research focused on processes like angiogenesis, permeability or many others, microvascular endothelial cells (ECs) are a much more physiologically relevant model to study 2. Furthermore, ECs isolated from knockout mice provide a useful tool for analysis of protein function ex vivo. Several approaches to isolate and culture microvascular ECs of different origin have been reported to date 3-7, but consistent isolation and culture of pure ECs is still a major technical problem in many laboratories. Here, we provide a step-by-step protocol on a reliable and relatively simple method of isolating and culturing mouse lung endothelial cells (MLECs). In this approach, lung tissue obtained from 6- to 8-day old pups is first cut into pieces, digested with collagenase/dispase (C/D) solution and dispersed mechanically into single-cell suspension. MLECS are purified from cell suspension using positive selection with anti-PECAM-1 antibody conjugated to Dynabeads using a Magnetic Particle Concentrator (MPC). Such purified cells are cultured on gelatin-coated tissue culture (TC) dishes until they become confluent. At that point, cells are further purified using Dynabeads coupled to anti-ICAM-2 antibody. MLECs obtained with this protocol exhibit a cobblestone phenotype, as visualized by phase-contrast light microscopy, and their endothelial phenotype has been confirmed using FACS analysis with anti-VE-cadherin 8 and anti-VEGFR2 9 antibodies and immunofluorescent staining of VE-cadherin. In our hands, this two-step isolation procedure consistently and reliably yields a pure population of MLECs, which can be further cultured. This method will enable researchers to take advantage of the growing number of knockout and transgenic mice to directly correlate in vivo studies with results of in vitro experiments performed on isolated MLECs and thus help to reveal molecular mechanisms of vascular phenotypes observed in vivo.

Here, we describe a protocol for isolation and culture of murine pulmonary endothelial cells. This method comprises mechanic and enzymatic lung tissue dissociation as well as a 2-step purification process using anti-PECAM-1 and anti-ICAM-2 antibodies conjugated to magnetic beads, which produces a pure endothelial cell population of mostly microvascular origin.

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells ...

Manual Isolation of Adipose-derived Stem Cells from Human Lipoaspirates

In 2001, researchers at the University of California, Los Angeles, described the isolation of a new population of adult stem cells from liposuctioned adipose tissue that they initially termed Processed Lipoaspirate Cells or PLA cells. Since then, these stem cells have been renamed as Adipose-derived Stem Cells or ASCs and have gone on to become one of the most popular adult stem cells populations in the fields of stem cell research and regenerative medicine. Thousands of articles now describe the use of ASCs in a variety of regenerative animal models, including bone regeneration, peripheral nerve repair and cardiovascular engineering. Recent articles have begun to describe the myriad of uses for ASCs in the clinic. The protocol shown in this article outlines the basic procedure for manually and enzymatically isolating ASCs from large amounts of lipoaspirates obtained from cosmetic procedures. This protocol can easily be scaled up or down to accommodate the volume of lipoaspirate and can be adapted to isolate ASCs from fat tissue obtained through abdominoplasties and other similar procedures.

In 2001, researchers at UCLA described the isolation of a population of adult stem cells, termed Adipose-derived Stem Cells or ASCs, from adipose tissue. This article outlines the isolation of ASCs from lipoaspirates using a manual, enzymatic digestion protocol using collagenase.

In 2001, researchers at the University of  California, Los Angeles, described the isolation of a new population of adult  stem cells from liposuctioned ...

Isolation of Murine Valve Endothelial Cells

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

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

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

Isolation and Primary Culture of Mouse Aortic Endothelial Cells

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

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

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

Identification and Dissection of Diverse Mouse Adipose Depots

Adipose tissues are complex organs with a wide array of functions, including storage and mobilization of energy in response to local and global needs, uncoupling of metabolism to generate heat, and secretion of adipokines to regulate whole-body homeostasis and immune responses. Emerging research is identifying important regional differences in the developmental, molecular, and functional profiles of adipocytes located in discrete depots throughout the body. Different properties of the depots are medically relevant since metabolic diseases often demonstrate depot-specific effects. This protocol will provide investigators with a detailed anatomic atlas and dissection guide for the reproducible and accurate identification and excision of diverse mouse adipose tissues. Standardized dissection of discrete adipose depots will allow detailed comparisons of their molecular and metabolic characteristics and contributions to local and systemic pathologic states under various nutritional and environmental conditions.

Adipocytes exist in discrete depots and have diverse roles within their unique microenvironments. As regional differences in adipocyte character and function are uncovered, standardized identification and isolation of depots is crucial for advancement of the field. Herein, we present a detailed protocol for the excision of various mouse adipose depots.

Adipose tissues are complex organs with a wide array of functions, including storage and mobilization of energy in response to local and global needs, ...

Isolation and Culture of Primary Aortic Endothelial Cells from Miniature Pigs

Xenotransplantation is a promising way to resolve the shortage of human organs for patients with end-stage organ failure, and the pig is considered as a suitable organ source. Immune rejection and coagulation are two major hurdles for the success of xenotransplantation. Vascular endothelial cell (EC) injury and dysfunction are important for the development of the inflammation and coagulation responses in xenotransplantation. Thus, isolation of porcine aortic endothelial cells (pAECs) is necessary for investigating the immune rejection and coagulation responses. Here, we have developed a simple enzymatic approach for the isolation, characterization, and expansion of highly purified pAECs from miniature pigs. First, the miniature pig was anaesthetized with ketamine, and a length of aorta was excised. Second, the endothelial surface of aorta was exposed to 0.005% collagenase IV digestive solution for 15 min. Third, the endothelial surface of the aorta was scraped in only one direction with a cell scraper (&#60;10 times), and was not compressed during the process of scraping. Finally, the isolated pAECs of Day 3, and after passage 1 and passage 2, were identified by flow cytometry with an anti-CD31 antibody. The percentages of CD31-positive cells were 97.4% &#177; 1.2%, 94.4% &#177; 1.1%, and 92.4% &#177; 1.7% (mean &#177; SD), respectively. The concentration of Collagenase IV, the digestive time, the direction, and frequency and intensity of scraping are critical for decreasing fibroblast contamination and obtaining high-purity and a large number of ECs. In conclusion, our enzymatic method is a highly-effctive method for isolating ECs from the miniature pig aorta, and the cells can be expanded in vitro to investigate the immune and coagulation responses in xenotransplantation.

An effective enzymatic method for isolation of primary porcine aortic endothelial cells (pAECs) from miniature pigs is described. The isolated primary pAECs can be used to investigate the immune and coagulation response in xenotransplantation.

Xenotransplantation is a promising way to resolve the shortage of human organs for patients with end-stage organ failure, and the pig is considered as a ...

Adipose Tissue-Derived Stem Cell Isolation and Characterization

Isolation and Identification of Mesenchymal Stem Cells Derived from Adipose Tissue of Sprague Dawley Rats

Adult mesenchymal cells have revolutionized molecular and cell biology in recent decades. They can differentiate into different specialized cell types, in addition to their great capacity for self-renewal, migration, and proliferation. Adipose tissue is one of the least invasive and most accessible sources of mesenchymal cells. It has also been reported to have higher yields compared to other sources, as well as superior immunomodulatory properties. Recently, different procedures for obtaining adult mesenchymal cells from different tissue sources and animal species have been published. After evaluating the criteria of some authors, we standardized a methodology that is applicable to different purposes and easily reproducible. A pool of stromal vascular fraction (SVF) from perirenal and epididymal adipose tissue allowed us to develop primary cultures with optimal morphology and functionality. The cells were observed adhered to the plastic surface for 24 h, and exhibited a fibroblast-like morphology, with prolongations and a tendency to form colonies. Flow cytometry (FC) and immunofluorescence (IF) techniques were used to assess the expression of the membrane markers CD105, CD9, CD63, CD31, and CD34. The ability of adipose-derived stem cells (ASCs) to differentiate into the adipogenic lineage was also assessed using a cocktail of factors (4 &#181;M insulin, 0.5 mM 3-methyl-iso-butyl-xanthine, and 1 &#181;M dexamethasone). After 48 h, a gradual loss of fibroblastoid morphology was observed, and at 12 days, the presence of lipid droplets positive to oil red staining was confirmed. In summary, a procedure is proposed to obtain optimal and functional ASC cultures for application in regenerative medicine.

Author Spotlight: Isolation and Identification of Mesenchymal Stem Cells Derived from Adipose Tissue of Sprague Dawley Rats  

This protocol describes a methodology for isolating and identifying adipose tissue-derived mesenchymal stem cells (MSCs) from Sprague Dawley rats.

Adult mesenchymal cells have revolutionized molecular and cell biology in recent decades. They can differentiate into different specialized cell types, in ...

Semi-Automated Isolation of Murine White Adipose Tissue

Semi-Automated Isolation of the Stromal Vascular Fraction from Murine White Adipose Tissue Using a Tissue Dissociator

The in vitro study of white, brown, and beige adipocyte differentiation enables the investigation of cell-autonomous functions of adipocytes and their mechanisms. Immortalized white preadipocyte cell lines are publicly available and widely used. However, the emergence of beige adipocytes in white adipose tissue in response to external cues is difficult to recapitulate to the full extent using publicly available white adipocyte cell lines. Isolation of the stromal vascular fraction (SVF) from murine adipose tissue is commonly executed to obtain primary preadipocytes and perform adipocyte differentiation. However, mincing and collagenase digestion of adipose tissue by hand can result in experimental variation and is prone to contamination. Here, we present a modified semi-automated protocol that utilizes a tissue dissociator for collagenase digestion to achieve easier isolation of the SVF, with the aim of reducing experimental variation, reducing contamination, and increasing reproducibility. The obtained preadipocytes and differentiated adipocytes can be used for functional and mechanistic analyses.

Author Spotlight: Semi-Automated Isolation of the Stromal Vascular Fraction from Murine White Adipose Tissue Using a Tissue Dissociator  

This protocol describes the semi-automated isolation of the stromal vascular fraction (SVF) from murine adipose tissue to obtain preadipocytes and achieve adipocyte differentiation in vitro. Using a tissue dissociator for collagenase digestion reduces experimental variation and increases reproducibility.

The in vitro study of white, brown, and beige adipocyte differentiation enables the investigation of cell-autonomous functions of adipocytes and their ...

Culturing Preadipocytes Isolated from Murine Perivascular Adipose Tissue

Isolation, Culture, and Adipogenic Induction of Stromal Vascular Fraction-derived Preadipocytes from Mouse Periaortic Adipose Tissue

Perivascular adipose tissue (PVAT) is an adipose tissue depot that surrounds blood vessels and exhibits the phenotypes of white, beige, and brown adipocytes. Recent discoveries have shed light on the central role of PVAT in regulating vascular homeostasis and participating in the pathogenesis of cardiovascular diseases. A comprehensive understanding of PVAT properties and regulation is of great importance for the development of future therapies. Primary cultures of periaortic adipocytes are valuable for studying PVAT function and the crosstalk between periaortic adipocytes and vascular cells. This paper presents an economical and feasible protocol for the isolation, culture, and adipogenic induction of stromal vascular fraction-derived preadipocytes from mouse periaortic adipose tissue, which can be useful for modeling adipogenesis or lipogenesis in vitro. The protocol outlines tissue processing and cell differentiation for culturing periaortic adipocytes from young mice. This protocol will provide the technological cornerstone at the bench side for the investigation of PVAT function.

Author Spotlight: The Significance of Isolation, Culture, and Adipogenic Induction of SVF-Derived Preadipocytes from Mouse Perivascular Adipose Tissue 

Here, we describe the isolation, culture, and adipogenic induction of stromal vascular fraction-derived preadipocytes from mouse periaortic adipose tissue, allowing for the study of perivascular adipose tissue function and its relationship with vascular cells.

Perivascular adipose tissue (PVAT) is an adipose tissue depot that surrounds blood vessels and exhibits the phenotypes of white, beige, and brown ...

Single Cell Nuclei Isolation from IMAT for Studying Gene Expression Profiles 

Isolation of Nuclei from Human Intermuscular Adipose Tissue and Downstream Single-Nuclei RNA Sequencing

Intermuscular adipose tissue (IMAT) is a relatively understudied adipose depot located between muscle fibers. IMAT content increases with age and BMI and is associated with metabolic and muscle degenerative diseases; however, an understanding of the biological properties of IMAT and its interplay with the surrounding muscle fibers is severely lacking. In recent years, single-cell and nuclei RNA sequencing have provided us with cell type-specific atlases of several human tissues. However, the cellular composition of human IMAT remains largely unexplored due to the inherent challenges of its accessibility from biopsy collection in humans. In addition to the limited amount of tissue collected, the processing of human IMAT is complicated due to its proximity to skeletal muscle tissue and fascia. The lipid-laden nature of the adipocytes makes it incompatible with single-cell isolation. Hence, single nuclei RNA sequencing is optimal for obtaining high-dimensional transcriptomics at single-cell resolution and provides the potential to uncover the biology of this depot, including the exact cellular composition of IMAT. Here, we present a detailed protocol for nuclei isolation and library preparation of frozen human IMAT for single nuclei RNA sequencing. This protocol allows for the profiling of thousands of nuclei using a droplet-based approach, thus providing the capacity to detect rare and low-abundant cell types.

Author Spotlight: Deciphering the Cellular Mysteries of Intermuscular Adipose Tissue in Humans

The biology of intermuscular adipose tissue (IMAT) is largely unexplored due to the limited accessibility of human tissue. Here, we present a detailed protocol for nuclei isolation and library preparation of frozen human IMAT for single nuclei RNA sequencing to identify the cellular composition of this unique adipose depot.

Intermuscular adipose tissue (IMAT) is a relatively understudied adipose depot located between muscle fibers. IMAT content increases with age and BMI and ...

Methods Article

Isolation and Identification of Vascular Endothelial Cells from Distinct Adipose Depots for Downstream Applications