This work was supported by the National Institute of Diabetes and Digestive and Kidney Diseases of the NIH (1RO1DK122948 to SHI) and the NIH Silvio O. Conte Digestive Diseases Research Core Centers P30 grant mechanism (DK084567). Support was also provided to KF by the Japan Society for the Promotion of Science (JSPS) Overseas Research Fellowships. We would also like to acknowledge Dr. Gregory J. Gores and Steven Bronk for their original design and optimization of the collagenase perfusion apparatus.

Animal protocols were conducted as approved by the Institutional Animal Care and Use Committee (IACUC) of Mayo Clinic. Eight-week-old C57BL/6J male mice were purchased from Jackson Laboratory. Mice were housed in a temperature-controlled 12:12-h light-dark cycle facility with free access to diet.
1. Preparation of collagen-coated culture dish or plate
<ol>
	<li>To make 50 mL of 0.02 mol/L acetic acid, add 0.6 mL of glacial acetic acid to 49.4 mL of H2O.</li>
	<li>Make 50 &#181;g/mL collagen type I in 0.02 mol/L acetic acid. Dilution depends on the concentration of the lot.</li>
	<li>Coat 10 cm culture dishes with 3 mL of collagen solution. Incubate at room temperature (RT) for 1 h. 
	NOTE: When using a different dish or plate for culture, the volume of coating solution needs to be adjusted based on culture area, generally use 6-10 &#181;g/cm2.</li>
	<li>Remove excess fluid from the coated surface, and wash with Phosphate-buffered saline (PBS) 3 times. Allow the dishes to air dry.</li>
	<li>If the collagen solution is not sterile, sterilize the collagen-coated dishes by exposure to ultraviolet (UV) light for 10 min in a sterile tissue culture hood.</li>
</ol>
2. Equipment setup
<ol>
	<li>Set up the heated and humidified recirculating perfusion apparatus as shown in Figure 1B.</li>
	<li>Rinse the perfusion system using 10% bleach for 5 min followed by sterile water for another 10 min.</li>
	<li>Drain the rinsing liquid as much as possible before perfusing the collagenase solution.</li>
	<li>Infuse collagenase solution through the perfusion system (as shown in Figure 1B) to prewarm it at 37 &#176;C. Set the pump rate at speed 1 as shown on the speed control dial (equal to 40 mL/min).</li>
	<li>Keep this speed the same throughout the whole procedure. Reuse the collagenase solution run in a closed circuit during the day of the LSEC isolation. 
	&#8203;NOTE: The pump speed is fixed at 1 to avoid mechanical pressure that could perturb LSECs biological condition.</li>
</ol>
3. Surgical procedure
<ol>
	<li>Weigh the mouse.</li>
	<li>Inject 90 mg/kg body weight of ketamine and 10 mg/kg bodyweight of xylazine intraperitoneally (IP).</li>
	<li>Once the mouse is nonreactive to painful stimuli, secure the mouse to the surgical surface, as shown in Figure 1B.</li>
	<li>Spray the mouse abdomen with 70% ethanol. Using surgical scissors, make an incision of ~5 cm long starting from the lower part of the abdomen up to the xiphoid process.</li>
	<li>Next, make two lateral cuts using small iris scissors on each side of the abdomen to fully expose the abdominal organs.</li>
	<li>Place the sheath of the intravenous (IV) catheter under the animal&#39;s back to lift and level the abdomen.</li>
	<li>Using a regular curved dressing forceps, gently pull the intestines and the stomach off to the left of the animal.</li>
	<li>Place a 5-0 surgical suture under the inferior vena cava (IVC) just below the exposed left kidney. Tie a loose hitch in the suture.</li>
	<li>Place another 5-0 surgical suture around the hepatic portal vein (PV) just above the splenic vein branching point off the hepatic PV. Tie a loose hitch in the suture.</li>
	<li>Using the PV suture as tension, insert the 20 G IV catheter in the hepatic PV 1 cm below where it branches to the right and left hepatic PV.</li>
	<li>Slide the catheter up the vein but keep it below the branching area. Allow the blood to travel down the catheter until it begins to drip out.</li>
	<li>With some of the Buffer A (Table 1) in an intravenous (IV) bottle positioned above the surgical area, use an IV line, and attach it to the catheter. Flush the liver with this solution while avoiding air entry into the system.</li>
	<li>Tie off the suture around the inferior vena cava below the kidney. This will allow the liver to retro perfuse.</li>
	<li>Cut the IVC below the suture to allow the animal to bleed out. 
	NOTE: This step must be performed quickly to avoid congestion of the liver.</li>
	<li>Secure the IV catheter with the PV suture.</li>
	<li>Once perfusing, cut away the stomach, intestines, spleen, and other entrails attached to the liver.</li>
	<li>Cut away the diaphragm and the major vessels from the thoracic cavity. Remove the liver from the animal and place it on the perfusion tray.</li>
	<li>Carefully remove the IV line and hook up the collagenase solution in the recirculating chamber. 
	NOTE: Steps 3.16-3.18&#160;need to be done within 5 min, so the liver will not be perfused too long with Buffer A. Be very careful not to allow any air bubbles into the liver.</li>
	<li>Allow the liver to perfuse until the capsule becomes mottled and appears mushy (10-15 min or more,&#160;period varies depending on the different lots of collagenase).</li>
	<li>While liver is in perfusion, make sure the animal is deceased before disposing it in a necropsy bag.</li>
	<li>Once digested, remove the liver from the chamber and place it in a 10 cm Petri dish with about 20 mL of serum-free Dulbecco&#39;s Modified Eagle Medium (DMEM).</li>
	<li>Gently pick apart the liver with a couple of pipette tips and discard the biliary tree.</li>
	<li>Filter the liver suspension through a 70 &#181;m cell strainer into a 50 mL conical tube.</li>
	<li>Centrifuge the cell suspension at 50 x g for 2 min at RT. Collect the supernatant containing nonparenchymal hepatic cells. 
	&#8203;NOTE: Hepatocytes are precipitated in the pellet, which can be further purified using gradient centrifugation as previously described15.</li>
</ol>
4. Separation of nonparenchymal hepatic cells and LSECs purification
NOTE: Purify the CD146+ LSECs using the immunomagnetic beads, following the manufacture&#39;s instructions.
<ol>
	<li>Centrifuge the supernatant at 300 x g for 5 min at 4 &#176;C.</li>
	<li>Collect the cell pellet and resuspend in 1 mL of isolation buffer (Table 1), determine the cell number using an automatic cell counter following the manufacturer&#39;s instructions.</li>
	<li>Centrifuge the cell suspension at 300 x g for 10 min at 4 &#176;C, aspirate the supernatant completely.</li>
	<li>Resuspend the pellet with 90 &#181;L of isolation buffer per 107 total cells.</li>
	<li>Add 10 &#181;L of CD146 magnetic beads per 107 total cells. Mix well and incubate for 15 min at 4 &#176;C.</li>
	<li>To wash the cells, add 1-2 mL of isolation buffer per 107 cells, centrifuge at 300 x g for 10 min, and then aspirate the supernatant completely.</li>
	<li>Resuspend up to 109 cells in 500 &#181;L of isolation buffer (Table 1). 
	NOTE: For higher cell numbers, scale up buffer volume accordingly.</li>
	<li>Prepare the separation column, rinse it with 3 mL of isolation buffer.</li>
	<li>Apply the cell suspension onto the column stacked with 70 &#181;m pre-separation filters.</li>
	<li>Wash the column with the 3 mL of isolation buffer 3 times. 
	NOTE: When washing the column, the isolation buffer should be added as soon as the column reservoir is empty.</li>
	<li>Remove the column from the separator and place it on a 15 mL centrifuge tube. Pipette 5 mL of isolation buffer onto the column.</li>
	<li>To recover the magnetic beads labeled cells, firmly push the plunger into the column to flush out the cells.</li>
	<li>Centrifuge at 300 x g for 5 min at 4 &#176;C. LSECs are ready for microscopic examination and downstream analysis.</li>
</ol>
5. LSECs immunophenotyping and purity assessment by flow cytometry
<ol>
	<li>Determine the cell numbers of isolated LSECs using an automatic cell counter following the manufacturer&#39;s instructions.</li>
	<li>Centrifuge the cells at 300 x g for 5 min, aspirate the supernatant completely.</li>
	<li>Take 1 x 106 cells/tube and resuspend with 90 &#181;L of staining buffer (Table 1).</li>
	<li>Add 10 &#181;L/tube of mouse FcR block and 1 &#181;L of viability dye, incubate at 4 &#176;C for 10 min.</li>
	<li>Stain the cells with a combination of CD45, CD146, and stabilin-2 antibodies diluted at 1:50 with staining buffer. Incubate at 4 &#176;C for 20 min.</li>
	<li>Wash the cells with 5 mL of staining buffer and centrifuge at 300 x g for 10 min, then aspirate the supernatant completely.</li>
	<li>Resuspend the cells with 300 &#181;L of staining buffer and run through the flow cytometer.</li>
</ol>
6. LSECs culture and examination
<ol>
	<li>Seed 1 x 106 cells/well in a 6-well plate and culture it with endothelial cells growth medium consisting of 5% fetal bovine serum (FBS), 1% endothelial cells growth supplement, and 1% primocin solution.</li>
	<li>Examine the isolated LSECs by light microscopy. Acquire bright-field images with a 10x magnification.</li>
</ol>
7. LSECs morphology and fenestrae examination by scanning electron microscopy
<ol>
	<li>Pre-coat the cell culture inserts (3 &#181;m pore size) with collagen solution.</li>
	<li>Culture isolated LSECs (120,000 cells) on the insert with endothelial cells growth medium in a 24-well plate at 37 &#176;C warm and humidified atmosphere. Allow cells to settle down and adhere for 2 h.</li>
	<li>For cell fixation, add an equivalent volume of prewarmed at 37 &#176;C Trump&#39;s fixative to the cell culture media.</li>
	<li>After incubation for 10 min, replace 50% diluted Trump&#39;s fixative with undiluted Trump&#39;s fixative.</li>
	<li>Fix the cells in Trump fixative for 2 h, then incubate for 1 h in 1% osmium tetroxides.</li>
	<li>Proceed with samples dehydration, drying in a critical point drying device, mounting, sputter coating, and examination using a scanning electron microscope.</li>
</ol>

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All authors have no conflicts to disclose.

In the current manuscript, we describe a protocol for LSEC isolation from mouse liver consisting of two-step collagenase perfusion and subsequent magnetic-activated cell sorting (MACS). This protocol consists of the following three steps: (1) Perfusion through the PV with a calcium-free buffer followed by a collagenase-containing buffer to achieve liver cell dispersion; (2) Exclusion of hepatocytes with low-speed centrifugation; and (3) MACS-based positive selection of LSECs from nonparenchymal cells (NPCs) using anti-CD...

Liver sinusoidal endothelial cells (LSECs) line the hepatic sinusoid walls and are the most abundant nonparenchymal cells in the liver1. LSECs are distinguished from other capillary endothelial cells elsewhere in the body by the presence of fenestrae and the lack of a classical basement membrane or a diaphragm2,3. Hence, the LSECs possess distinctive phenotypic and structural characteristics that enhance their permeability and endocytic capacity to eliminate a variety of circulating macromolecules, including lipids and lipoproteins. LSECs play a pivotal role in the crosstalk between parenchymal and nonparenchymal cells, such as stellate cells and immune cells. LSECs are key in maintaining liver homeostasis by keeping the stellate cells and Kupffer cells in a quiescent status4. LSECs modulate the composition of hepatic immune cells populations by mediating adhesion and trans-endothelial migration of circulating leukocytes5,6. During acute and chronic liver injury7, including ischemia-reperfusion injury (IRI)8, nonalcoholic steatohepatitis (NASH)9, and hepatocellular carcinoma (HCC), LSECs undergo phenotypic changes known as capillarization characterized by defenestration and formation of basement membrane10. These phenotypic changes in LSECs are associated with LSECs dysfunction and the acquisition of prothrombotic, proinflammatory, and profibrogenic properties.
Several methods for the isolation of LSECs from mouse liver have been developed11. Some techniques depend on separating nonparenchymal and parenchymal cells followed by density gradient centrifugation to purify the LSECs from nonparenchymal fractions. The limitation of this method is the presence of contaminating macrophages in the final steps of LSECs isolation, which could affect the purity of the isolated LSECs12. This protocol is based on collagenase perfusion of the mouse liver and CD146+ magnetic beads positive selection of nonparenchymal cells to purify LSECs. LSECs isolated using this method show high purity and preserved morphology and viability. These LSECs are optimum for functional studies, including permeability and adhesion assay, as well as downstream studies for pathways of interest. Moreover, with the growing interest in generating big datasets in both clinical research and discovery science, these high-quality LSECs isolated from both healthy and diseased livers with nonalcoholic steatohepatitis (NASH) or other conditions can be pooled or used individually, allowing multi-omics data generation and comparison between health and disease13,14. In addition, the isolated LSECs can be employed to develop two-dimensional as well as three-dimensional in vitro models like organoids to decipher the activated signaling pathway in LSECs and their intercellular communication with other liver cells under different noxious stimuli and in response to various therapeutic interventions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2.0-inch 20 G Intra Venous (IV) catheter</td>
			<td>Terumo, SOmerset, NJ, USA</td>
			<td>SR-OX2051CA</td>
			<td></td>
		</tr>
		<tr>
			<td>2&#8211;3-inch perfuion tray with a hole in the center</td>
			<td></td>
			<td></td>
			<td>customized; made in house</td>
		</tr>
		<tr>
			<td>405/520 viability dye</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-110-205</td>
			<td></td>
		</tr>
		<tr>
			<td>4-inch regular curved dressing forceps</td>
			<td>Fisher Brand</td>
			<td>FS16-100-110</td>
			<td></td>
		</tr>
		<tr>
			<td>5-0 Perma-Hand silk suture</td>
			<td>Ethicon, Raritan, NJ, USA</td>
			<td>A182H</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-stabilin-2 (Mouse) mAb-Alexa Fluor&#195; 488</td>
			<td>MBL International, Woburn, MA, USA</td>
			<td>D317-A48</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA stock</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-091-376</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-CD146 (LSEC)-PE, anti-mouse</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-118-407</td>
			<td></td>
		</tr>
		<tr>
			<td>CD146 (LSEC) MicroBeads, mouse</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-092-007</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-CD45-Viogreen, anti-mouse</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-110-803</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagen type I</td>
			<td>Corning, Corning, NY, USA</td>
			<td>354236</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase II</td>
			<td>Gibco, Waltham, MA, USA</td>
			<td>17101-015</td>
			<td></td>
		</tr>
		<tr>
			<td>Endothelial cells growth medium</td>
			<td>ScienCell Research Laboratories, Carlsbad, CA, USA</td>
			<td>211-500</td>
			<td></td>
		</tr>
		<tr>
			<td>FcR blocking reagent, mouse</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-092-575</td>
			<td></td>
		</tr>
		<tr>
			<td>FlowJo software, version 10.6</td>
			<td>Becton, Dickinson and Company</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hardened Fine scissors</td>
			<td>F.S.T, Foster city, CA, USA</td>
			<td>14091-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Heated (37 &#176;C) and humidified recirculating perfusion apparatus equipped with Oxygen injection at a rate of 10psi.</td>
			<td></td>
			<td></td>
			<td>customized; made in house</td>
		</tr>
		<tr>
			<td>Hitachi S 4700 scanning electron microscope</td>
			<td>Hitachi Inc, Pleasanton, CA, USA</td>
			<td>SEM096</td>
			<td></td>
		</tr>
		<tr>
			<td>LS columns</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-042-401</td>
			<td></td>
		</tr>
		<tr>
			<td>MACS pre-separation filters (70 &#956;m)</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-095-823</td>
			<td></td>
		</tr>
		<tr>
			<td>MACS rinsing buffer</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-091-222</td>
			<td></td>
		</tr>
		<tr>
			<td>MACS Smart Strainer (70 &#956;m)</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td>130-098-462</td>
			<td></td>
		</tr>
		<tr>
			<td>MACSQunt flow cytometer</td>
			<td>Miltenyi, Bergisch Gladbach, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Millicell Cell Culture Insert</td>
			<td>Millipore Sigma, Burlington, MA, USA</td>
			<td>PITP01250</td>
			<td></td>
		</tr>
		<tr>
			<td>Nexcelom cell counter</td>
			<td>Nexcelom bioscience, Lawrence, MA, USA</td>
			<td>Cellometer Auto T4 Plus</td>
			<td></td>
		</tr>
		<tr>
			<td>Percoll</td>
			<td>GE Healthcare, Chicago, IL, USA</td>
			<td>17-0891-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical scissors</td>
			<td>F.S.T, Foster city, CA, USA</td>
			<td>14001-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Very small curved dressing forceps</td>
			<td>F.S.T, Foster city, CA, USA</td>
			<td>11063-07</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Experimental schematics and equipment set up: 
In this protocol, mouse liver was digested using a closed perfusion circuit, then nonparenchymal cells and hepatocytes were separated by low-speed centrifugation at 50 x g for 2 min. Primary LSECs were isolated using CD146 magnetic beads selection from the nonparenchymal fraction. The experimental schematics are shown in Figure 1A. The cannula was placed through the PV while the inferior vena cava was tied up to en...

Watch this Scientific Journal Video about Isolation and Characterization of Mouse Primary Liver Sinusoidal Endothelial Cells at JoVE.com

Isolation and Characterization of Mouse Primary Liver Sinusoidal Endothelial Cells

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. ...

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6.1K Views.  Mayo Clinic. 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. 

Video: Isolation and Characterization of Mouse Primary Liver Sinusoidal Endothelial Cells

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 of Primary Mouse Trophoblast Cells and Trophoblast Invasion Assay

The placenta is responsible for the transport of nutrients, gasses and growth factors to the fetus, as well as the elimination of wastes. 
Thus, defects in placental development have important consequences for the fetus and mother, and are a major cause of embryonic lethality. The major cell type of the 
fetal portion of the placenta is the trophoblast. Primary mouse placental trophoblast cells are a useful tool for studying normal and abnormal placental development, 
and unlike cell lines, may be isolated and used to study trophoblast at specific stages of pregnancy. In addition, primary cultures of trophoblast from transgenic
mice may be used to study the role of particular genes in placental cells. The protocol presented here is based on the description by Thordarson et al.1, in which a 
percoll gradient is used to obtain a relatively pure trophoblast cell population from isolated mouse placentas. It is similar to the more widely used methods for 
human trophoblast cell isolation2-3. Purity may be assessed by immunocytochemical staining of the isolated cells for cytokeratin 74. Here, the isolated cells are 
then analyzed using a matrigel invasion assay to assess trophoblast invasiveness in vitro5-6. The invaded cells are analyzed by immunocytochemistry and stained 
for counting.

In this protocol, we describe the dissection of placentae from the mouse on pregnancy d10.5, followed by isolation of trophoblast cells using a Percoll gradient. We then demonstrate use of the isolated cells in a matrigel invasion assay.

The placenta is responsible for the transport of nutrients, gasses and growth factors to the fetus, as well as the elimination of wastes.  
Thus, defects ...

Isolation and Culture of Mouse Primary Pancreatic Acinar Cells

This protocol permits rapid isolation (in less than 1 hr) of murine pancreatic acini, making it possible to maintain them in culture for more than one week. More than 20 x 106 acinar cells can be obtained from a single murine pancreas. This protocol offers the possibility to independently process as many as 10 pancreases in parallel. Because it preserves acinar architecture, this model is well suited for studying the physiology of the exocrine pancreas in vitro in contrast to cell lines established from pancreatic tumors, which display many genetic alterations resulting in partial or total loss of their acinar differentiation.

In this publication, we describe a rapid and convenient procedure for isolating and culturing primary pancreatic acinar cells from the murine pancreas. This method constitutes a valuable approach to study the physiology of fresh primary normal/untransformed exocrine pancreatic cells.

This protocol permits rapid isolation (in less than 1 hr) of murine pancreatic acini, making it possible to maintain them in culture for more than one ...

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 ...

A Standardized Method for the Analysis of Liver Sinusoidal Endothelial Cells and Their Fenestrations by Scanning Electron Microscopy

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved substances from the blood into the liver parenchyma for metabolism and processing. Fenestrations are dynamic structures - both their size and/or number can be altered in response to various physiological states, drugs, and disease, making them an important target for modulation. An understanding of how LSEC morphology is influenced by various disease, toxic, and physiological states and how these changes impact on liver function requires accurate measurement of the size and number of fenestrations. In this paper, we describe scanning electron microscopy fixation and processing techniques used in our laboratory to ensure reproducible specimen preparation and accurate interpretation. The methods include perfusion fixation, secondary fixation and dehydration, preparation for the scanning electron microscope and analysis. Finally, we provide a step by step method for standardized image analysis which will benefit all researchers in the field.

The fenestrated liver sinusoidal endothelial cell is a biologically important filter system that is highly influenced by various diseases, toxins, and physiological states. These changes significantly impact on liver function. We describe methods for the standardisation of the measurement of the size and number of fenestrations in these cells.

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved ...

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 ...

Protocol for Isolation of Primary Human Hepatocytes and Corresponding Major Populations of Non-parenchymal Liver Cells

Beside parenchymal hepatocytes, the liver consists of non-parenchymal cells (NPC) namely Kupffer cells (KC), liver endothelial cells (LEC) and hepatic Stellate cells (HSC). Two-dimensional (2D) culture of primary human hepatocyte (PHH) is still considered as the &#34;gold standard&#34; for in vitro testing of drug metabolism and hepatotoxicity. It is well-known that the 2D monoculture of PHH suffers from dedifferentiation and loss of function. Recently it was shown that hepatic NPC play a central role in liver (patho-) physiology and the maintenance of PHH functions. Current research focuses on the reconstruction of in vivo tissue architecture by 3D- and co-culture models to overcome the limitations of 2D monocultures. Previously we published a method to isolate human liver cells and investigated the suitability of these cells for their use in cell cultures in Experimental Biology and Medicine1. Based on the broad interest in this technique the aim of this article was to provide a more detailed protocol for the liver cell isolation process including a video, which will allow an easy reproduction of this technique.
Human liver cells were isolated from human liver tissue samples of surgical interventions by a two-step EGTA/collagenase P perfusion technique. PHH were separated from the NPC by an initial centrifugation at 50 x g. Density gradient centrifugation steps were used for removal of dead cells. Individual liver cell populations were isolated from the enriched NPC fraction using specific cell properties and cell sorting procedures. Beside the PHH isolation we were able to separate KC, LEC and HSC for further cultivation.
Taken together, the presented protocol allows the isolation of PHH and NPC in high quality and quantity from one donor tissue sample. The access to purified liver cell populations could allow the creation of in vivo like human liver models.

A technique to isolate human hepatocytes and non-parenchymal liver cells from the same donor is described. The different liver cell types build the basis for functional liver models and tissue engineering. This new method aims to isolate liver cells in a high yield and viability.

Beside parenchymal hepatocytes, the liver consists of non-parenchymal cells (NPC) namely Kupffer cells (KC), liver endothelial cells (LEC) and hepatic ...

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, Characterization, And High Throughput Extracellular Flux Analysis of Mouse Primary Renal Tubular Epithelial Cells

Mitochondrial dysfunction in the renal tubular epithelial cells (TECs) can lead to renal fibrosis, a major cause of&#160;chronic kidney disease (CKD). Therefore, assessing mitochondrial function in primary TECs may provide valuable insight into the bioenergetic status of the cells, providing insight into the pathophysiology of CKD. While there are a number of complex protocols available for the isolation and purification of proximal tubules in different species, the field lacks a cost-effective method optimized for&#160;tubular cell isolation without the need for purification. Here, we provide an isolation protocol that allows for studies focusing on both primary mouse proximal and distal renal TECs. In addition to cost-effective reagents and minimal animal procedures required in this protocol, the isolated cells maintain high energy levels after isolation and can be sub-cultured up to four&#160;passages, allowing for continuous studies. Furthermore, using a high throughput extracellular flux analyzer, we assess the mitochondrial respiration directly in the isolated TECs in a 96-well plate for which we provide recommendations for the optimization of cell density and compound concentration. These observations suggest that this protocol can be used for renal tubular ex vivo studies with a consistent, well-standardized production of renal TECs. This protocol may have broader future applications to study mitochondrial dysfunction associated with renal disorders for drug discovery or drug characterization purposes.

This protocol provides a cost-effective approach to isolate and characterize mouse primary renal tubular cells that can subsequently be sub-cultured to assess renal biological functions ex vivo, including mitochondrial bioenergetics.

Mitochondrial dysfunction in the renal tubular epithelial cells (TECs) can lead to renal fibrosis, a major cause of&#160;chronic kidney disease (CKD). ...

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 ...