The authors thank Dr. Pooneh Bagher and Dr.&#160;Erika Westcott for editorial comments in preparing the manuscript. This work was supported by the National Heart, Lung and Blood Institute of the National Institutes of Health under award numbers R37-HL041026 and R01-HL086483 to SSS and F32-HL107050 to MJS. This content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

1. Preparation of Equipment: Flow Chamber, Pipettes, and Solutions
<ol>
	<li>Flow chamber: Use a flow chamber with a 24 mm x 50 mm glass coverslip on the bottom to superfuse the isolated endothelial tubes (see Figure 1A). Before assembling the chamber, wash the glass coverslips with both 3% HCl and 70% ethanol, rinse with ultrapure water, and dry with nitrogen gas.</li>
	<li>Pipettes: Three different types of pipettes are used during the protocol: cannulation, trituration, and pinning. Both the cannulation and pinning pipettes can be made before the day of the experiment, but the trituration pipette should be made the day of as it requires knowledge of the diameter of the vessel to size the lumen appropriately.
	<ol>
		<li>Cannulation pipettes: In order to flush red blood cells from the lumen, pull borosilicate glass capillary tubes using a micropipette puller to have long, tapered ends. Break the ends with forceps to an outer diameter ~30% less than that of the arteries (~50-80 &#181;m), and fire polish the tip to be smooth (see Figure 1B). It is helpful to also angle the tips of the pipettes to approximately 45&#176;.</li>
		<li>Trituration pipettes: To mechanically dissociate the smooth muscle cells and adventitia from the endothelial tubes, make triturating pipettes from borosilicate glass capillary tubes. Prepare capillary tubes as above, and break the end with forceps. Tip: Use a glass scoring device to make it easier to break the thicker wall of the trituration pipette glass cleanly. Break the pipette to the internal diameter determined on the day of the experiment, approximately 10-20 &#181;m larger than that of the outer diameter of the vessel from which endothelial tubes will be isolated. Fire polish the broken ends until smooth (see Figure 1C).</li>
		<li>Pinning pipettes: To stabilize the endothelial tube within the flow chamber, make pinning pipettes to secure the endothelial tube against the coverslip bottom of the chamber. Pull glass capillary tubes in the same manner as the cannulation pipettes. Then break the ends with forceps to a diameter of ~100 &#181;m, and fire polish until the tips are sealed, rounded and smooth (see Figure 1D). Note: If the tip of the pinning pipette is not completely sealed, fluid will enter the lumen and may confound experimental results.</li>
	</ol>
	</li>
	<li>Solutions: Prepare all solutions in ultrapure (18.2 M&#937;) H2O and sterilize them through a 0.2 &#181;m filter. Make sure the osmolality of the solutions is between 285-300 mOsm. Solutions remain good for approximately two weeks when stored at 4 &#176;C.
	<ol>
		<li>Dissection solution: The salt solution used during the dissection of the superior epigastric artery is composed of (in mmol/L): 137 NaCl, 5 KCl, 1 MgCl2, 10 N-2-hydroxyethylpiperazine-N&#39;-2-ethanesulfonic acid (HEPES), 10 glucose, 0.01 sodium nitroprusside (SNP), and 0.1% bovine serum albumin (BSA) with a pH of 7.4. The NO donor SNP is included to help relax smooth muscle cells, thus making them easier to cannulate the artery and to facilitate smooth muscle cell removal during trituration.</li>
		<li>Dissociation solution: The salt solution used during the dissociation of the endothelial tube is composed of (in mmol/L): 137 NaCl, 5 KCl, 1 MgCl2, 10 HEPES, 10 glucose, 2 CaCl2, and 0.1% BSA, with a pH of 7.4. To a 1 ml&#160;aliquot of this buffer, add: 0.62 mg/ml&#160;papain, 1.5 mg/ml&#160;collagenase and 1.0 mg/ml&#160;dithioerythritol. Since enzymes from different suppliers can have varying levels of enzymatic activity, the product numbers of those used for this work are included for reference in the table of materials. Note: During the protocol both dissociation buffer and dissociation buffer with enzymes are used at different steps.</li>
		<li>Superfusion solution: The physiological salt solution (PSS) used for superfusing the isolated endothelial tube is composed of (in mmol/L): 137 NaCl, 5 KCl, 1 MgCl2, 10 HEPES, 10 glucose and 2 CaCl2, with a pH of 7.4.</li>
		<li>Prepare buffers and solutions before beginning surgical procedures. Remove the solutions from 4 &#176;C and aliquot 50 ml&#160;of Dissection buffer into a 200 ml&#160;Erlenmeyer flask and place it on ice. Additionally, aliquot 50 ml&#160;of Dissociation buffer without enzymes and place it on the bench top to equilibrate to room temperature. Remove the superfusion solution from 4 &#176;C and allow it to equilibrate to room temperature on the bench top.</li>
	</ol>
	</li>
</ol>
2. Animal Preparation
Make sure that all procedures and protocols involving animals are in accord with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. The procedures described here were approved by the Animal Care and Use Committee of the University of Missouri.
<ol>
	<li>Use male or female mice at least 9 weeks old and study each individually. Anesthetize a mouse with pentobarbital sodium (60 mg/kg) via intraperitoneal (i.p.) injection. Tip: Diluting the pentobarbital to 10 mg/ml in sterile saline before injection reduces the possibility of overdose.</li>
	<li>Anesthesia compromises temperature regulation, so keep the mouse warm immediately after the first injection of pentobarbital. Using a metal carrier (ventilated aluminum basket) on top of a heating plate (calibrated to ~37 &#176;C) works well to keep the temperature of the mouse stable. Alternatively, use a heat lamp, taking care to position it at an appropriate distance from the mouse. Monitor the mouse every 5-10 min&#160;until the appropriate level of anesthesia is achieved (lack of withdrawal to toe or tail pinch). A supplementary dose may be required after 15-20 min. It is best to proceed slowly and with caution to avoid overdose, especially with obese, aged, or animals otherwise stressed.</li>
	<li>Remove the hair from the ventral side by carefully shaving it with small pet hair clippers. Particular attention should be taken to avoid trauma to the animal. It is helpful to remove all hair from an area spanning the armpits to the pubic area. Remove any loosely clinging hair with a fresh alcohol swab.</li>
	<li>Position the mouse on its back, lying with both fore- and hind limbs spread out to expose as much of the abdominal wall as possible. If necessary, use laboratory tape to secure the legs in a spread-eagle manner.</li>
</ol>
3. Isolation of the Superior Epigastric Artery
<ol>
	<li>Make a small incision through the skin just above the area of the pubic region. Ensure that only the skin is being cut while avoiding damage to the underlying abdominal muscle; extend the incision laterally in each direction out to respective hindlimbs. Continue the incision rostrally along the ventral midline to the top of the rib cage and then extend the incision laterally in each direction to respective forelimbs.</li>
	<li>Gently lift the skin and use scissors to sever the connective tissue holding the skin to the underlying muscle, thereby exposing the entire surface of the abdominal musculature (see Figure 3A). Irrigate the exposed muscle with room temperature 0.9% saline solution.</li>
	<li>Due to the small size of the SEA, position the animal underneath a stereomicroscope for further procedures. Lift the fat pad located at the bottom of the sternum with forceps, and make an incision through it. Continue the incision along the bottom rib, making sure not to cut too deeply into the underlying tissue. The SEA should be visible; take care to avoid damaging it. Once the incision is complete, irrigate the exposed tissue with room temperature 0.9% saline solution.</li>
	<li>After the top layer of skeletal muscle has been retracted, the SEA and underlying muscle are easily identified. Tip: Note the length of the SEA in vivo before isolating to extend endothelial tubes (which shorten along their axis upon isolation) to approximate their original length. Using forceps and scissors, carefully excise the thin muscle layer underneath the SEA.</li>
	<li>Using angled forceps, pass a length of 6-0 silk suture underneath the SEA and ligate the artery and its adjacent vein to keep it pressurized and blood retained within the vessel lumen to facilitate visualization during subsequent isolation and cleaning.</li>
	<li>Repeat the SEA ligation procedure on the contralateral side.</li>
	<li>Once both SEAs have been ligated, make an incision along the midline of the abdominal muscles to separate respective sides.</li>
	<li>Extend the incision laterally in each direction as done for the skin, then continue the incision vertically along the outer edge to completely separate the abdominal muscle from the body. Avoid damage to vasculature fed by the SEA to maintain the pressurized lumen.</li>
	<li>Cut the SEA above the ligation to maintain the seal, and place the isolated muscle and artery in a 50 ml&#160;beaker containing 10 ml&#160;of 4 &#176;C dissection buffer.</li>
	<li>Repeat the procedure for the other side of the abdomen and incubate the tissues in the dissection buffer for 10 min.</li>
	<li>Place the abdominal muscle containing the SEA in a chilled (4 &#176;C) dissection chamber containing dissection buffer (see Figure 3B). The dissection chamber consists of a Petri dish with a layer of Sylgard (polydimethylsiloxane [PDMS]) at the bottom.</li>
	<li>Using insect pins (0.15 mm), stretch the isolated SEA and abdominal muscle out to approximate in vivo lengths (noted above) and secure it to the Sylgard. Tip: Orienting the muscle such that the thin layer facing the peritoneum is on top makes the SEA readily visible using transillumination.</li>
	<li>Utilizing fine microdissection instruments and working from the upstream site of ligation towards the downstream end, clear the SEA from its paired vein and the surrounding tissue until the first major branch site (1-2 cm). Then excise the SEA by cutting it just above the branch site and just below the ligation.</li>
	<li>To remove blood retained within the vessel lumen, cannulate the SEA and flush it with ice cold dissection buffer. Using a piece of silastic tubing, attach the back end of a cannulation pipette to a 5 ml&#160;syringe containing ice cold dissection buffer and secure the micropipette in a micromanipulator to position its tip within the dissection chamber to cannulate the SEA.</li>
	<li>Once all of the erythrocytes are flushed out, remove the SEA from the cannulation pipette, and lift the pipette out of the dissection dish.</li>
	<li>Cut the SEA into smaller segments each 1-3 mm in length. These shorter segments facilitate the isolation of endothelial tubes.</li>
</ol>
4. Isolation of the Endothelial Tube
<ol>
	<li>Fill a 12 mm x 75 mm glass culture tube halfway with ice cold dissection buffer. Using angled forceps, transfer the arterial pieces into the culture tube and place on ice. At this time, combine the digestion enzymes with Dissociation buffer to a final volume of 1 ml&#160;in a separate 12 mm x 75 mm culture tube (see 1.3.2 Solutions) and preheat the solution to 37 &#176;C with a heating block.</li>
	<li>While the dissociation buffer and enzymes are warming up, carefully aspirate the dissection buffer from the culture tube leaving a small volume containing the vessel segments.</li>
	<li>Gently add room temperature dissociation buffer without enzymes to wash away remaining dissection buffer. Tip: Add the dissociation buffer slowly such that the arterial pieces remain on the bottom of the culture tube to save time waiting for them to resettle. Raise the buffer temperatures over multiple steps from ice (4 &#176;C), to room temperature (~24 &#176;C), to 37 &#176;C to minimize shock.</li>
	<li>Carefully aspirate the dissociation buffer from the culture tube, again being sure to leave a small volume containing the vessel segments.</li>
	<li>Remove the preheating dissociation buffer with enzymes from the heating block, transfer the 1 ml&#160;of enzyme solution to the culture tube containing the vessel segments, and place the culture tube back in the heating block. Incubate at 37 &#176;C for 30 min.</li>
	<li>During the incubation with enzymes, prepare the trituration pipette, backfill it with mineral oil and secure it on a microsyringe that is mounted in a micromanipulator. Then retract the microsyringe plunger to fill the pipette with 2 nl&#160;of dissociation buffer and position the pipette tip over the flow chamber.</li>
	<li>At the end of the 30 min&#160;incubation, remove the culture tube from the heating block and carefully aspirate the buffer as above.</li>
	<li>Wash the arterial segments with 4 ml&#160;of room temperature dissociation buffer. Note: It is no longer necessary to keep the vessel segments at the bottom of the culture tube; disturbing the arterial pieces actually helps to ensure that residual enzymes are effectively removed.</li>
	<li>Transfer one vessel segment to the flow chamber by gently aspirating it with a 1 ml&#160;micropipette, and place it into the flow chamber with 1 ml&#160;of the dissociation buffer.</li>
	<li>Utilizing the micromanipulator containing the microsyringe, position the tip of the trituration pipette near one end of the vessel segment.</li>
	<li>Aspirate the vessel segment (500 nl&#160;withdrawn) into the triturating pipette slowly (225 nl/sec), so as not to cause mechanical strain to the ECs, and then eject it back into the chamber. If the digestion was successful, the smooth muscle cells and adventitia will be dissociated from the endothelial tube.</li>
	<li>Repeat the trituration until all smooth muscle cells are dissociated. Be aware that excessive trituration (4x or more) can damage the ECs and render them unresponsive to agonists. Tip: using the trituration pipette, move the adventitia away from the endothelial tube as soon as possible to keep it from entangling the tube. Note: Using the trituration pipette, the isolated endothelial tube can be aspirated and transferred to a separate chamber.</li>
	<li>Position the endothelial tube in the center of the flow chamber, aligned along the direction of flow, and remove the triturating pipette.</li>
	<li>Secure pinning pipettes in micromanipulators mounted at either end of the flow chamber, and position their tips at respective ends of the endothelial tube.</li>
	<li>Lower a pinning pipette over one end of the tube to press it against the bottom of the chamber. Position the second pinning pipette in the same way at the opposite end of the tube. Note: Placement of the pinning pipettes is a trade-off between securing the tube (placing them farther from the end of the tube) and allowing for more imaging/experimental area (placing them closer to the end of the tube). Locating them ~50 &#181;m from each end of the tube works well. Just as the pinning pipette touches the tube, slowly retract it along the axis of the tube, thereby extending it to approximate in vivo length; then press the pipette against the chamber bottom to secure the tube.</li>
	<li>Begin the flow of superfusion solution, and allow the endothelial tube to rest for at least 20 min&#160;to equilibrate and adhere (see Figure 3C). Utilize a peristaltic pump to maintain constant fluid flow (superfusion) across the endothelial tube. Note: Pinning pipettes can be removed once the endothelial tube has adhered to the bottom of the flow chamber (~25 min) or left in place to ensure the endothelial tube does not become dislodged.</li>
</ol>
5. Dye Loading and Calcium Imaging
<ol>
	<li>To visualize intracellular calcium responses, load the endothelial tube with fluo-4 AM dye at room temperature (~24 &#176;C). Stop fluid flow in the bath, and add fluo-4 AM to the chamber at a final concentration of 10 &#181;M. Wait 10 min to allow the dye to enter cells, then superfuse the endothelial tube with fresh PSS for 20 min to ensure extracellular dye is removed and that intracellular dye has been fully de-esterified.</li>
	<li>Perform calcium imaging. For this work, an upright microscope (see Figure 2) equipped with a spinning disc confocal imaging system was used at ambient room temperature. Excite the fluorescent calcium dye with a 491 nm laser and acquire images (at 500-550 nm emission) using an intensified digital camera.</li>
</ol>

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Here we describe the isolation of endothelial tubes from the SEA and the use of this preparation to visualize Ca2+ signaling events within its constitutive ECs. This procedure was adapted from one originally developed to isolate endothelial tubes from arterioles of the hamster cremaster muscle8. Utilizing minor variations of the techniques presented here, we have isolated endothelial tubes from a variety of vascular beds, including: feed arteries of the hamster retractor muscle and cheek pouch arterioles10, mouse superior epigastric artery6,7,9, mouse mesenteric and cerebral arteries and lymphatic microvessels (unpublished observations; references are included for isolating mesenteric vessels12 and cerebral vessels13). Isolating endothelial tubes from new vascular beds may require modifications to the original protocol. If isolation is unsuccessful, it is best to begin by altering digestion times: Increase the digestion time if endothelial tubes are difficult to isolate from smooth muscle cells and adventitia and reduce digestion time if the endothelial tube does not remain intact. If altering digestion times is unsuccessful, adjusting the concentration of digestion enzymes or the digestion temperature should be tried. Another option is to reduce the internal diameter of the trituration pipette, which increases shear force in removing smooth muscle cells. Increasing the velocity of fluid ejection can also be tried for the same effect but is more likely to damage the preparation. It should be recognized that it may not be possible to isolate intact endothelial tubes from vessels in which endothelial cells are not well connected to each other by junctional proteins.
Dissociation of smooth muscle cells following enzymatic digestion was initially performed using a hand-style pipettor8. We have refined the trituration procedure using a microsyringe system, which has enabled consistent isolation of longer endothelial tubes (up to 3 mm)6. When using this system, the trituration pipette is backfilled with mineral oil to provide a continuous fluid column between the piston controlling fluid movement and PSS containing the vessel segment. The incompressibility of fluid along with the constant driving force of the microsyringe piston results in constant shear as the vessel is forced through the pipette tip. In contrast, hand techniques often used for dissociating cells (e.g.&#160;squeezing rubber bulb at the end of a Pasteur pipette) involves the compression of air, which introduces variability in the driving pressure and, thereby, shear exerted at the pipette tip.
There are multiple advantages to the tube preparation for studying endothelial function in microvessels. The first is that the isolation process provides distinct homogenous native cellular populations of ECs and smooth muscle cells. Since the ECs remain physically connected as tubes, they are distinct from individual smooth muscle cells, which retain a &#34;C&#34; configuration if they remain relaxed during trituration. Thus respective cell types are readily distinguished, e.g.&#160;when obtaining samples for molecular techniques such as real-time PCR and immunohistochemistry10. Since multiple tubes are isolated from a single vessel (or from bilateral vessels as done for the SEA), molecular data and functional data can be obtained from the same vessels of a given animal. Thus molecular expression can be correlated with the functional behavior of microvascular endothelium. Further, both intra- and intercellular signaling events intrinsic to microvascular endothelium can be resolved independent from the influence of hemodynamic forces (pressure, flow), vasoactive agents carried in the blood stream (e.g.&#160;hormones), or from surrounding smooth muscle cells (e.g.&#160;via myoendothelial coupling14-16), nerves17, or tissue parenchyma18.
Importantly, since the endothelium is freshly isolated from designated microvessels, there is no alteration of phenotype that is otherwise associated with culturing ECs19-21. For example, cultured ECs lose muscarinic receptor expression and thereby alter their calcium signaling profiles. Further, electrophysiological properties of ECs can change in culture22. Because individual ECs remain coupled through functional gap junction channels, the endothelial tube presents an ideal model for studying the conduction of electrical and Ca2+ signals between cells6,7. It should also be recognized that the dissociated smooth muscle cells are readily studied with patch-clamp techniques for complementary data underlying microvascular function23.
A key limitation to the endothelial tube model is the instability of the preparation with increasing temperature. While our preparations from the SEA have proven stable and healthy at ambient room temperature (~24 &#176;C) and for several hours at 32 &#176;C, morphological and functional degradation occur in less than an hour at 37 &#176;C9. A second important limitation of the endothelial tube is the loss of myoendothelial junctions and their inherent signaling domains that are integral to EC function in the intact vessel wall14-16. It should also be recognized that, while longer tubes enable intercellular signaling to be studied over relatively great distances6, preparation of tubes is complicated as they get longer because it becomes more difficult to fully dissociate surrounding smooth muscle cells and adventitia. We have found that tubes longer than 1-2 mm are also more difficult to position and secure in the flow chamber. In contrast, while shorter tubes (e.g.&#160;&#60;1 mm) enable Ca2+ and electrical signals to be studied8, they are difficult to maintain during superfusion in the flow chamber. Finally, even with optimal isolation of tubes bilaterally, there is insufficient material for traditional quantification of protein expression using Western blots, though immunolabeling provides an index of protein expression and localization. Despite such limitations, the endothelial tube represents a novel preparation for providing new insight into mechanisms of microvascular endothelial cell function in vivo.

In this protocol, we describe the isolation of endothelial cell tubes from the mouse SEA. The SEA arises from the internal thoracic artery and supplies oxygenated blood to the anterior abdominal musculature. Our working with the SEA as a model is attributable to it providing relatively long, unbranched microvessel segments that are well suited for studying intercellular signaling events underlying the role of the endothelium in governing and coordinating smooth muscle cell relaxation. For those interested in tissues other than skeletal muscle, we have found this technique to be readily adapted for obtaining endothelial tubes from microvessels of brain, gut and the lymphatic system.
The key features of this method are that intact lengths (1-3 mm) of microvascular endothelium are isolated, secured against a glass coverslip in a flow chamber in which they are superfused with PSS, and imaged for Ca2+ signaling7-9 or impaled for electrical signaling6,8. Within the flow chamber, the upper half of a tube is exposed to the superfusion solution and responds to experimental interventions, while the bottom half (in contact with the coverslip) is protected and remains quiescent. In addition to studying both intra- and intercellular Ca2+ signaling events, endothelial tubes can be used for quantitative real-time PCR to assess gene expression6,10, immunohistochemistry to investigate protein expression6,10, and electrophysiological studies to define biophysical properties of electrical conduction6,11.
There are several advantages to the endothelial tube preparation for studying endothelial function. First is that the isolation process provides a simple distinction between two cell populations: endothelial cells (which remain in tube formation) and smooth muscle cells (individually dissociated; typically &#34;C&#34; shaped when relaxed). This allows for selective sampling and study of unique properties underlying respective cells&#39; contribution to vessel function. Second, this model enables the resolution of signaling events intrinsic to the endothelium, independent from the influence of the blood flow, surrounding smooth muscle, nerves, and parenchyma. Third, the endothelium is studied while freshly isolated, thereby avoiding alterations in gene or protein expression associated with cell culture.

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			<td height="22" style="height:22px;width:159px;">Sodium Chloride</td>
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			<td style="width:159px;">C1016</td>
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			<td style="width:159px;">Sigma</td>
			<td style="width:159px;">C-8051</td>
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			<td style="width:159px;">Warner Instruments</td>
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			<td style="width:159px;"></td>
		</tr>
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			<td style="width:159px;">World Precision Instruments</td>
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			<td style="width:159px;">Updated to Nanoliter 2010 from Nanoliter 2000</td>
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			<td height="43" style="height:43px;width:159px;">Micro4 Controller</td>
			<td style="width:159px;">World Precision Instruments</td>
			<td style="width:159px;">SYS-MICRO4</td>
			<td style="width:159px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:159px;">12 mm x 75 mm Culture Tubes</td>
			<td style="width:159px;">Fisher</td>
			<td style="width:159px;">14-961-26</td>
			<td style="width:159px;"></td>
		</tr>
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			<td height="43" style="height:43px;width:159px;">3-Axis Micromanipulator</td>
			<td style="width:159px;">Siskiyou Corp</td>
			<td style="width:159px;">DT3-100</td>
			<td style="width:159px;">Holding and positioning the pinning pipettes</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:159px;">Flow Chamber</td>
			<td style="width:159px;">Warner Instruments</td>
			<td style="width:159px;">RC-27N</td>
			<td style="width:159px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:159px;">100-1,000 &#181;l Pipette</td>
			<td style="width:159px;">Eppendorf</td>
			<td>3120000062</td>
			<td style="width:159px;">For transferring digested vessel segment to the flow chamber</td>
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			<td height="64" style="height:64px;width:159px;">1 ml Pipette Tips</td>
			<td style="width:159px;">Fisher</td>
			<td style="width:159px;">02-707-405</td>
			<td style="width:159px;">For transferring digested vessel segment to the flow chamber</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:159px;">Upright Microscope</td>
			<td style="width:159px;">Olympus</td>
			<td style="width:159px;">BX51W1</td>
			<td style="width:159px;">The actual microscope is up to the user</td>
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			<td height="43" style="height:43px;width:159px;">Spinning Disc Confocal System</td>
			<td style="width:159px;">Yokogawa</td>
			<td style="width:159px;">CSU-X1</td>
			<td style="width:159px;">Confocal imaging is optional</td>
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			<td height="43" style="height:43px;width:159px;">XR/TURBO EX Camera</td>
			<td style="width:159px;">Stanford Photonics</td>
			<td style="width:159px;">XR/TURBO EX</td>
			<td style="width:159px;">Ideal for our needs; may vary with user</td>
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		<tr height="43">
			<td height="43" style="height:43px;width:159px;">Piper Control Software</td>
			<td style="width:159px;">Stanford Photonics</td>
			<td style="width:159px;"></td>
			<td style="width:159px;">Imaging software for TURBO EX camera</td>
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			<td height="22" style="height:22px;width:159px;">Stereo Microscrope</td>
			<td style="width:159px;">Leica</td>
			<td style="width:159px;">MZ8</td>
			<td style="width:159px;">may vary with user</td>
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			<td height="22" style="height:22px;width:159px;">Sylgard</td>
			<td style="width:159px;">Dow Corning</td>
			<td style="width:159px;">184</td>
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			<td style="width:159px;">15003-08</td>
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			<td style="width:159px;">Austerlitz</td>
			<td style="width:159px;">M size 0.15 mm</td>
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			<td style="width:159px;">Millipore</td>
			<td style="width:159px;">SCGPT05RE</td>
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			<td height="43" style="height:43px;width:159px;">Pipette Scoring Device</td>
			<td style="width:159px;">Austin Flameworks</td>
			<td style="width:159px;">Small Handheld Scoring Tool</td>
			<td style="width:159px;">austinflameworks.com</td>
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			<td style="width:159px;">Model 9966</td>
			<td style="width:159px;">Clean well after each use to maintain life</td>
		</tr>
	</tbody>
</table>

isolation of microvascular endothelial tubes from mouse resistance arteries

The control of blood flow by the resistance vasculature regulates the supply of oxygen and nutrients concomitant with the removal of metabolic by-products, as exemplified by exercising skeletal muscle. Endothelial cells (ECs) line the intima of all resistance vessels and serve a key role in controlling diameter (e.g.&#160;endothelium-dependent vasodilation) and, thereby, the magnitude and distribution of tissue blood flow. The regulation of vascular resistance by ECs is effected by intracellular Ca2+ signaling, which leads to production of diffusible autacoids (e.g.&#160;nitric oxide and arachidonic acid metabolites)1-3 and hyperpolarization4,5 that elicit smooth muscle cell relaxation. Thus understanding the dynamics of endothelial Ca2+ signaling is a key step towards understanding mechanisms governing blood flow control. Isolating endothelial tubes eliminates confounding variables associated with blood in the vessel lumen and with surrounding smooth muscle cells and perivascular nerves, which otherwise influence EC structure and function. Here we present the isolation of endothelial tubes from the superior epigastric artery (SEA) using a protocol optimized for this vessel.
To isolate endothelial tubes from an anesthetized mouse, the SEA is ligated in situ to maintain blood within the vessel lumen (to facilitate visualizing it during dissection), and the entire sheet of abdominal muscle is excised. The SEA is dissected free from surrounding skeletal muscle fibers and connective tissue, blood is flushed from the lumen, and mild enzymatic digestion is performed to enable removal of adventitia, nerves and smooth muscle cells using gentle trituration. These freshly-isolated preparations of intact endothelium retain their native morphology, with individual ECs remaining functionally coupled to one another, able to transfer chemical and electrical signals intercellularly through gap junctions6,7. In addition to providing new insight into calcium signaling and membrane biophysics, these preparations enable molecular studies of gene expression and protein localization within native microvascular endothelium.

Calcium responses can be initiated in freshly isolated endothelial tubes using acetylcholine (ACh). In this movie, an endothelial tube loaded with 10 &#181;M fluo-4 AM is stimulated with superfusion of 1 &#181;M ACh (Movie 1).&#160;<a href="https://www.jove.com/files/ftp_upload/50759/SochaandSegal_movie1.avi" target="_blank">Click here to view movie</a>. The dye is excited at 491 nm and emission is recorded from 500-550 nm using an intensified charge-coupled device camera. Image stacks are acquired at 120 frames/sec for a 25 sec period and can then be averaged (e.g.&#160;to 40 frames/sec). Representative fluorescence images over time are shown in Figure 4.
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/50759/50759fig1.jpg" width="500px" /> 
Figure 1. Flow chamber and pipettes used during isolation of endothelial tubes. A) The flow chamber assembled with a 24 mm x 50 mm glass coverslip as the base. B) Examples of a pulled (left) and a pulled, broken, polished and angled (right) cannulation pipette. This pipette is used to flush erythrocytes from the lumen of the artery following dissection. Scale bar = 200 &#181;m. C) Examples of a pulled (left) and a pulled, broken and polished (right) trituration pipette. The internal diameter of this pipette is determined by the exterior diameter of the artery. Scale bar = 200 &#181;m. D) Examples of a pulled (left) and a pulled, broken and polished (right) pinning pipette. The tip of this pipette is rounded off and completely sealed to secure the endothelial tube in the flow chamber. Scale bar = 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50759/50759fig1highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 2" fo:content-width="6in" src="/files/ftp_upload/50759/50759fig2.jpg" width="500px" /> 
Figure 2. Spinning disc confocal microscope for imaging of Ca2+ signals. A) Upright microscope used for calcium imaging. B) (a) Confocal spinning disc unit with an (b) intensified charge coupled device camera. C) Immersion objectives (a) 63X (NA = 1) and (b) 20X (NA = 0.5) used for imaging. <a href="https://www.jove.com/files/ftp_upload/50759/50759fig2highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/50759/50759fig3.jpg" width="500px" /> 
Figure 3. Isolation of the abdominal muscle and superior epigastric artery. A) Anesthetized mouse with abdominal muscle exposed and irrigated with room temperature 0.9% saline solution. Red arrows mark the sites under fat pads at which respective SEAs enter each rectus abdominis muscle (i.e.&#160;where vessels should be ligated). Scale bar = 1 cm. B) Isolated abdominal muscle (unilateral) and SEA pinned out in a dissection chamber for microdissection. Red arrow indicates the first major bifurcation site of the SEA (i.e. the point at which cleaning of the vessel is stopped). Scale bar = 1 cm. C) Isolated endothelial cell tube from an SEA. Differential interference contrast image of an endothelial tube following 1 hr incubation at room temperature. The endothelial tube was superfused with Superfusion buffer at 3 ml/min. Scale bar = 50 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50759/50759fig3highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/50759/50759fig4.jpg" width="500px" /> 
Figure 4. Calcium fluorescence in an endothelial cell tube. Representative fluorescent images of an endothelial tube in response to stimulation with 1 &#181;M acetylcholine. A) Fluorescence before stimulation. B) endothelial tube fluorescence at the time of acetylcholine administration, t=0 sec. C) endothelial tube fluorescence at t=5 sec. D) endothelial tube fluorescence at t=10 sec. E) endothelial tube fluorescence at t=15 sec. F) endothelial tube fluorescence at t=20 sec. Scale bars = 40 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50759/50759fig4highres.jpg" target="_blank">Click here to view larger image</a>.

Watch this Scientific Journal Video about Isolation of Microvascular Endothelial Tubes from Mouse Resistance Arteries at JoVE.com

Isolation of Microvascular Endothelial Tubes from Mouse Resistance Arteries

We present a preparation for visualizing and manipulating calcium signaling in native, intact microvascular endothelium. Endothelial tubes freshly isolated from mouse resistance arteries supplying skeletal muscle retain in vivo morphology and dynamic signaling within and between neighboring cells. Endothelial tubes can be prepared from microvessels of other tissues and organs.

The control of blood flow by the resistance vasculature regulates the supply of oxygen and nutrients concomitant with the removal of metabolic ...

isolation-microvascular-endothelial-tubes-from-mouse-resistance

Research

JoVE Journal

Biology

15.6K Views.  University of Missouri. We present a preparation for visualizing and manipulating calcium signaling in native, intact microvascular endothelium. Endothelial tubes freshly isolated from mouse resistance arteries supplying skeletal muscle retain in vivo morphology and dynamic signaling within and between neighboring cells. Endothelial tubes can be prepared from microvessels of other tissues and organs.

Video: Isolation of Microvascular Endothelial Tubes from Mouse Resistance Arteries

Isolation of Valvular Endothelial Cells

Heart valves are solely responsible for maintaining unidirectional blood flow through the cardiovascular system. These thin, fibrous tissues are subjected to significant mechanical stresses as they open and close several billion times over a lifespan. The incredible endurance of these tissues is due to the resident valvular endothelial (VEC) and interstitial cells (VIC) that constantly repair and remodel in response to local mechanical and biological signals. Only recently have we begun to understand the unique behaviors of these cells, for which in vitro experimentation has played a key role. Particularly challenging is the isolation and culture of VEC. Special care must be used from the moment the tissue is removed from the host through final plating. Here we present protocols for direct isolation, side specific isolation, culture, and verification of pure populations of VEC. We use enzymatic digestion followed by a gentle swab scraping technique to dislodge only surface cells. These cells are then collected into a tube and centrifuged into a pellet. The pellet is then resuspended and plated into culture flasks pre-coated with collagen I matrix. VEC phenotype is confirmed by contact inhibited growth and the expression of endothelial specific markers such as PECAM1 (CD31), Von Willebrand Factor (vWF), and negative expression of alpha-smooth muscle actin (&alpha;-SMA). The functional characteristics of VEC are associated with high levels of acetylated LDL. Unlike vascular endothelial cells, VEC have the unique capacity to transform into mesenchyme, which normally occurs during embryonic valve formation1. This can also occur during significantly prolonged post confluent in vitro culture, so care should be made to passage at or near confluence. After VEC isolation, pure populations of VIC can then be easily acquired.

We provide a method for isolating and culturing pure populations of heart valve endothelial cells (VEC). VEC can be isolated from either side of the cusp or leaflet and immediately following, underlying interstitial cell (VIC) isolation is straightforward.

Heart valves are solely responsible for maintaining unidirectional blood flow through the cardiovascular system.  These thin, fibrous tissues are ...

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 ...

RNA-seq Analysis of Transcriptomes in Thrombin-treated and Control Human Pulmonary Microvascular Endothelial Cells

The characterization of gene expression in cells via measurement of mRNA levels is a useful tool in determining how the transcriptional machinery of the cell is affected by external signals (e.g. drug treatment), or how cells differ between a healthy state and a diseased state. With the advent and continuous refinement of next-generation DNA sequencing technology, RNA-sequencing (RNA-seq) has become an increasingly popular method of transcriptome analysis to catalog all species of transcripts, to determine the transcriptional structure of all expressed genes and to quantify the changing expression levels of the total set of transcripts in a given cell, tissue or organism1,2 . RNA-seq is gradually replacing DNA microarrays as a preferred method for transcriptome analysis because it has the advantages of profiling a complete transcriptome, providing a digital type datum (copy number of any transcript) and not relying on any known genomic sequence3.
Here, we present a complete and detailed protocol to apply RNA-seq to profile transcriptomes in human pulmonary microvascular endothelial cells with or without thrombin treatment. This protocol is based on our recent published study entitled "RNA-seq Reveals Novel Transcriptome of Genes and Their Isoforms in Human Pulmonary Microvascular Endothelial Cells Treated with Thrombin,"4 in which we successfully performed the first complete transcriptome analysis of human pulmonary microvascular endothelial cells treated with thrombin using RNA-seq. It yielded unprecedented resources for further experimentation to gain insights into molecular mechanisms underlying thrombin-mediated endothelial dysfunction in the pathogenesis of inflammatory conditions, cancer, diabetes, and coronary heart disease, and provides potential new leads for therapeutic targets to those diseases.
The descriptive text of this protocol is divided into four parts. The first part describes the treatment of human pulmonary microvascular endothelial cells with thrombin and RNA isolation, quality analysis and quantification. The second part describes library construction and sequencing. The third part describes the data analysis. The fourth part describes an RT-PCR validation assay. Representative results of several key steps are displayed. Useful tips or precautions to boost success in key steps are provided in the Discussion section. Although this protocol uses human pulmonary microvascular endothelial cells treated with thrombin, it can be generalized to profile transcriptomes in both mammalian and non-mammalian cells and in tissues treated with different stimuli or inhibitors, or to compare transcriptomes in cells or tissues between a healthy state and a disease state.

This protocol presents a complete and detailed procedure to apply RNA-seq, a powerful next-generation DNA sequencing technology, to profile transcriptomes in human pulmonary microvascular endothelial cells with or without thrombin treatment. This protocol is generalizable to various cells or tissues affected by different reagents or disease states.

The characterization of gene expression in cells via measurement of mRNA levels is a useful tool in determining how the transcriptional machinery of the ...

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 ...

En Face Detection of Nitric Oxide and Superoxide in Endothelial Layer of Intact Arteries

Endothelium-derived nitric oxide (NO) produced from endothelial NO-synthase (eNOS) is one of the most important vasoprotective molecules in cardiovascular physiology. Dysfunctional eNOS such as uncoupling of eNOS leads to decrease in NO bioavailability and increase in superoxide anion (O2.&#8722;) production, and in turn promotes cardiovascular diseases. Therefore, appropriate measurement of NO and O2.&#8722; levels in the endothelial cells are pivotal for research on cardiovascular diseases and complications. Because of the extremely labile nature of NO and O2.&#8722;, it is difficult to measure NO and O2.&#8722; directly in a blood vessel. Numerous methods have been developed to measure NO and O2.&#8722; production. It is, however, either insensitive, or non-specific, or technically demanding and requires special equipment. Here we describe an adaption of the fluorescence dye method for en face simultaneous detection and visualization of intracellular NO and O2.&#8722; using the cell permeable diaminofluorescein-2 diacetate (DAF-2DA) and dihydroethidium (DHE), respectively, in intact aortas of an obesity mouse model induced by high-fat-diet feeding. We could demonstrate decreased intracellular NO and enhanced O2.&#8722; levels in the freshly isolated intact aortas of obesity mouse as compared to the control lean mouse. We demonstrate that this method is an easy technique for direct detection and visualization of NO and O2.&#8722; in the intact blood vessels and can be widely applied for investigation of endothelial (dys)function under (physio)pathological conditions.

En Face Detection of Nitric Oxide and Superoxide in Endothelial Layer of Intact Arteries

In this article we describe an adapted relatively easy method using the fluorescence dye diaminofluorescein-2 diacetate (DAF-2DA) and dihydroethidium (DHE) for en face simultaneous detection and visualization of intracellular nitric oxide (NO) and superoxide anion (O2.&#8722;) respectively, in freshly isolated intact aortas of an obesity mouse model.

Endothelium-derived nitric oxide (NO) produced from endothelial NO-synthase (eNOS) is one of the most important vasoprotective molecules in cardiovascular ...

Isolation of Mouse Coronary Endothelial Cells

Endothelial cells line the inner wall of blood vessels and play an important role in the regulation of vascular tone, vascular permeability, and new vascular formation. Endothelial cell dysfunction is implicated in the development and progression of many cardiovascular diseases including ischemic heart disease. To examine the function and characterization of coronary endothelial cells, cell isolation is the first step and it requires high purity and quantity to conduct subsequent experiments. This protocol describes an efficient method to isolate adult mouse coronary endothelial cells. The mouse heart is dissected and minced into small pieces. After the digestion of the heart using dispase and collagenase II, cells are washed&#160;and incubated with magnetic beads which are conjugated with anti-CD31 antibody. The beads with endothelial cells are washed several times and are ready to use in various applications, including imaging and molecular biological experiments. Efficient isolation yields approximately 104 cells per one heart with over 90% purity.

This protocol is prepared to share our method of isolating mouse coronary endothelial cells for the purpose of imaging or to conduct molecular biological experiments.

Endothelial cells line the inner wall of blood vessels and play an important role in the regulation of vascular tone, vascular permeability, and new ...

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 ...

Isolation and Characterization of Mouse Primary Liver Sinusoidal Endothelial Cells

Liver sinusoidal endothelial cells (LSECs) are specialized endothelial cells located at the interface between the circulation and the liver parenchyma. LSECs have a distinct morphology characterized by the presence of fenestrae and the absence of basement membrane. LSECs play essential roles in many pathological disorders in the liver, including metabolic dysregulation, inflammation, fibrosis, angiogenesis, and carcinogenesis. However, little has been published about the isolation and characterization of the LSECs. Here, this protocol discusses the isolation of LSEC from both healthy and nonalcoholic fatty liver disease (NAFLD) mice. The protocol is based on collagenase perfusion of the mouse liver and magnetic beads positive selection of nonparenchymal cells to purify LSECs. This study characterizes LSECs using specific markers by flow cytometry and identifies the characteristic phenotypic features by scanning electron microscopy. LSECs isolated following this protocol can be used for functional studies, including adhesion and permeability assays, as well as downstream studies for a particular pathway of interest. In addition, these LSECs can be pooled or used individually, allowing multi-omics data generation including RNA-seq bulk or single cell, proteomic or phospho-proteomics, and Assay for Transposase-Accessible Chromatin using sequencing (ATAC-seq), among others. This protocol will be useful for investigators studying LSECs&#39; communication with other liver cells in health and disease and allow an in-depth understanding of the role of LSECs in the pathogenic mechanisms of acute and chronic liver injury.

Here we outline and demonstrate a protocol for primary mouse liver sinusoidal endothelial cell (LSEC) isolation. The protocol is based on liver collagenase perfusion, nonparenchymal cell purification by low-speed centrifugation, and CD146 magnetic bead selection. We also phenotype and characterize these isolated LSECs using flow cytometry and scanning electron microscopy.

Liver sinusoidal endothelial cells (LSECs) are specialized endothelial cells located at the interface between the circulation and the liver parenchyma. ...

Isolation of Endothelial Cells from the Lumen of Mouse Carotid Arteries for Single-Cell Multi-Omics Experiments

Atherosclerosis is an inflammatory disease of the arterial regions exposed to disturbed blood flow (d-flow). D-flow regulates the expression of genes in the endothelium at the transcriptomic and epigenomic levels, resulting in proatherogenic responses. Recently, single-cell RNA sequencing (scRNAseq) and single-cell Assay for Transposase Accessible Chromatin sequencing (scATACseq) studies were performed to determine the transcriptomic and chromatin accessibility changes at a single-cell resolution using the mouse partial carotid ligation (PCL) model. As endothelial cells (ECs) represent a minor fraction of the total cell populations in the artery wall, a luminal digestion method was used to obtain EC-enriched single-cell preparations. For this study, mice were subjected to PCL surgery to induce d-flow in the left carotid artery (LCA) while using the right carotid artery (RCA) as a control. The carotid arteries were dissected out two days or two weeks post PCL surgery. The lumen of each carotid was subjected to collagenase digestion, and endothelial-enriched single cells or single nuclei were obtained. These single-cell and single-nuclei preparations were subsequently barcoded using a 10x Genomics microfluidic setup. The barcoded single-cells and single-nuclei were then utilized for RNA preparation, library generation, and sequencing on a high-throughput DNA sequencer. Post bioinformatics processing, the scRNAseq and scATACseq datasets identified various cell types from the luminal digestion, primarily consisting of ECs. Smooth muscle cells, fibroblasts, and immune cells were also present. This EC-enrichment method aided in understanding the effect of blood flow on the endothelium, which could have been difficult with the total artery digestion method. The EC-enriched single-cell preparation method can be used to perform single-cell omics studies in EC-knockouts and transgenic mice where the effect of blood flow on these genes has not been studied. Importantly, this technique can be adapted to isolate EC-enriched single cells from human artery explants to perform similar mechanistic studies.

We present a method for isolating endothelial cells and nuclei from the lumen of mouse carotid arteries exposed to stable or disturbed flow conditions to perform single-cell omics experiments.

Atherosclerosis is an inflammatory disease of the arterial regions exposed to disturbed blood flow (d-flow). D-flow regulates the expression of genes in ...

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Subtitles

Isolation of Microvascular Endothelial Tubes from Mouse Resistance Arteries