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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</ol>

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-CD146 magnetic beads. The whole procedure could be completed within 3 h. Furthermore, the cost for this procedure, including all the consumable supplies, is around 150 USD per mouse, suggesting that this method of LSEC isolation is overall efficient and cost-effective. We use steps (1) and (2) for isolation of primary mouse hepatocytes as well, which is beyond the scope of the current manuscript.
There are a few critical steps in portal cannulation and liver perfusion: (i)&#160;Catheter positioning: the catheter tip should be placed no less than 3 mm distal to the PV bifurcation of an adult mouse, which is usually identified in the hilum of the liver. Deep placement of the catheter tip in the PV will compromise the perfusion of some lobes of the liver. Poor perfusion of a lobe manifests as an absence of a color change of the lobe and can be rectified if noticed immediately by a slight repositioning of the catheter into the distal PV. This maneuver could possibly optimize lobe perfusion. (ii) Air bubbles: the infusion route between the pump and the PV should be kept completely free from air bubbles. Minute air entry into the PV can cause an air embolism, which results in incomplete liver perfusion. To eliminate air in the catheter after removing the inner needle, Buffer A is injected with a syringe proximal to the bubble to expel any air before connecting the infusion line to the catheter. (iii) Collagenase strength: To maintain collagenase activity during the perfusion, it is essential to assure that Buffer B infused into the PV has the precise concentration of collagenase and pH and is kept at approximately 37 &#176;C. As shown in Figure 1, a customized chamber is used to keep the whole perfusion system in a 37 &#176;C warm and humidified atmosphere; in addition, the oxygen supply is maintained during collagenase perfusion to achieve complete digestion. An alternative option includes a water bath to pre-warm the collagenase solution20, and an open collagenase perfusion system in situ where the collagenase solution drains out of the liver through the cut IVC21.
Although the protocol uses relatively bold catheters (20 G) to secure an adequate perfusion flow, a thinner catheter will also work well based on previous reports22,23,24. However, it is sometimes difficult to successfully cannulate the PV especially when mice are used for specific disease models or if they have PV anomalies. If PV cannulation fails, mechanical and enzymatic liver digestion using an enzyme kit and gentle tissue dissociator is an alternative method for steps (1) and (2) to obtain nonparenchymal cell (NPC) suspension. We have confirmed that the cell yield and viability in this alternative method of dissociation without liver perfusion was comparable to the original method (data not shown). Portal vein perfusion for liver digestion has been employed by others and us13,16,20. Furthermore, this method avoids the theoretical risk of LSECs phenotypic or functional alterations induced by mechanical force during liver digestion. Therefore, the use of the PV perfusion-based method depicted in this protocol is so far highly recommended.
Here, the optimum LSEC yield and purity when employing the current protocol are shown. We obtained approximately 2-5 x 106 cells per mouse from both healthy mice and mice with diet-induced NASH using the current protocol. As for purity, the positivity of the same surface marker used for immunomagnetic separation (CD146) supports the accuracy of our isolation technique. In addition, there are several well recognized LSEC surface markers, such as lymphatic vessel endothelial hyaluronic acid receptor 1 (LYVE-1), CD32b, and stabilin-2. Controversies still exist around these markers; for example, i) LYVE-1 is present also in lymphatic endothelial cells, and ii) LSECs in a periportal area lack CD32b expression18. Thus, to examine the purity of these isolated LSECs with flow cytometry, stabilin-2 was employed as an established specific LSEC marker in addition to CD146.&#160;Using this methodology, we confirmed that the isolated LSECs had &#62;94% viability and &#62;90% purity (Figure 2). The LSEC-specific phenotype of the isolated cells was further confirmed using SEM, where most of the cells have fenestrae (Figure 3B), the characteristic morphological feature of LSECs.
Among the LSECs isolation methodologies, immunomagnetic beads selection is the technique used the most to isolate the LSECs from the NPC fraction24,25,26,27. In contrast, other methods include centrifugal elutriation and selective adhesion-based separation22,28,29. The non-immunomagnetic methods have been shown to deliver high yields of LSECs (approximately 9 x 106 cells per mouse)11. Hence, a relatively lower yield of isolated LSECs could be a limitation of the current protocol. On the other hand, the non-immunomagnetic selection-based methods require high technical expertise and sometimes show inconsistent results on cell yields and purities11, while the immunomagnetic selection-based method holds the advantage of consistent yield and purity over other existing methods. Moreover, for immunomagnetic beads selection-based cell isolation, the specificity of cell surface markers used is always a matter of concern. For example, CD31, which has been used previously as a marker for immunomagnetic LSEC separation, is dominantly expressed on the PV endothelium and capillarized LSECs3. Although CD146 has been reported to be expressed also on natural killer cells and hepatic stellate cells30,31,the high purity of LSECs isolated using this protocol mitigates concerns regarding the specificity of the endothelial marker CD146. Comparison of the pros and cons of various methodologies for LSEC isolation is discussed elsewhere11,18,32.Comparison of cell yields and purities between other methods and this method is beyond the scope of the current manuscript and warrants future investigations.
In conclusion, we present a primary mouse LSEC isolation protocol based on one of the most widely used and accepted approaches. Isolated LSECs display high purity and preserved function. Moreover, this protocol is efficient and applicable to the healthy mouse liver as well as the liver from different disease mouse models. High-quality LSECs from these mice can be employed for multiomics data set generation13,14. Thus, this approach facilitates the identification of molecular mechanisms regulating the function of LSEC and improves our understanding of their roles in intercellular communication in the liver in health and disease.

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><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&amp;ndash;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&amp;Atilde; 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 &amp;deg;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 &amp;mu;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 &amp;mu;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 ensure unidirectional perfusion of collagenase through the liver (Figure 1B). The in-house perfusion chamber is equipped with a heating and humidification system to ensure a 37-40 &#176;C warm and humidified apparatus air. The perfused collagenase circulates through a closed system and can be reused for two or three mice during the isolation day to make the isolation more cost-effective.
Assessment of isolated LSECs purity and surface markers by flow cytometry: 
The purity of the isolated LSECs was assessed by employing well-characterized specific LSEC surface markers CD146 and stabilin-216,17,18. Stained cells were analyzed on a flow cytometer; data were analyzed using FlowJo software.
LSEC population was gated (Figure 2A) and analyzed singlets for the following gating strategy; viable cells were defined using viability dye staining before labeling cells with specific markers (Figure 2B). Then CD45- population was gated to exclude any immune cells contamination. The isolated LSECs reached 94.8% viability (Figure 2C). The percentage of CD45- cells was 89.7% (Figure 2D), and 92.3% of the cells were CD146 and stabilin-2 double-positive (CD45-CD146+stabilin-2+) LSECs (Figure 2E).
Morphology of the isolated LSECs: 
The isolated LSECs were seeded in a 6-well plate at 1 x 106 cells/well, cultured in a complete growth medium, and examined by light microscopy after 6 h of culture (Figure 3A). LSECs were plated on a 3 &#181;m pore-sized culture insert to visualize the LSECs fenestrae by scanning electron microscope (SEM). After fixation and processing, the fenestrae were identified, as shown in Figure 3B. As previously reported, LSECs lose their fenestrae in culture overnight19; hence, it is recommended to process the cells as soon as possible after the isolation for downstream studies to avoid dedifferentiation in vitro.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63062/63062fig01.jpg" /> 
Figure 1: Experimental schematics and equipment set up. (A) Experimental schematics were designed using Bio render. (B) Equipment setup and process. The liver was harvested and perfused in a closed collagenase perfusion apparatus. The liver was then transferred into a dish and dissociated. The liver suspension was collected. The nonparenchymal cell fraction was separated by centrifugation at 50 x g for 2 min. LSECs were purified using magnetic beads. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63062/63062fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63062/63062fig02.jpg" /> 
Figure 2: Assessment of isolated LSECs purity and surface markers by flow cytometry. Data analysis and gating strategy as shown, (A) gated LSECs and (B) singlets based on forward scatter (FSC)/side scatter (SSC). Isolated LSECs were stained with a viability dye and a combination of CD45, CD146, and stabilin-2 antibodies. Singlets were gated and analyzed, (C) viable LSECs were gated on (D) CD45 population and analyzed for (E) CD146 and stabilin-2 expression. All gates were determined by fluorescence minus one (FMO). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63062/63062fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63062/63062fig03.jpg" /> 
Figure 3: Morphology of isolated LSECs. (A) Bright field picture of cultured LSECs. Isolated LSECs were cultured in the complete growth medium for 6 h and examined by light microscopy (Scale bar 100 &#181;m). (B) LSEC was examined by scanning electron microscope (SEM). White arrows indicate LSEC fenestrae (Scale bar, left panel 5 &#181;m, right panel 1 &#181;m). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63062/63062fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">KRH stock solution (10x)</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td>Concentration</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>1.15 M</td>
		</tr>
		<tr>
			<td>HEPES (free acid)</td>
			<td>0.2 M</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>0.05 M</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>0.01 M</td>
		</tr>
		<tr>
			<td colspan="2">Buffer A</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td>Concentration</td>
		</tr>
		<tr>
			<td>KRH solution, PH=7.4</td>
			<td>1x</td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>0.5 mM</td>
		</tr>
		<tr>
			<td colspan="2">Adjust PH to 7.4, filter through a 0.2 &#956;m filter before use.</td>
		</tr>
		<tr>
			<td colspan="2">Stored at room temperature for up to 6 months.</td>
		</tr>
		<tr>
			<td colspan="2">Buffer B</td>
		</tr>
		<tr>
			<td>KRH solution, PH=7.4</td>
			<td>1x</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td colspan="2">Adjust PH to 7.4, filter through a 0.2 &#956;m filter before use.</td>
		</tr>
		<tr>
			<td colspan="2">Stored at room temperature for up to 6 months.</td>
		</tr>
		<tr>
			<td colspan="2">Collagenase solution</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td>Concentration</td>
		</tr>
		<tr>
			<td>Buffer B, PH=7.4</td>
			<td>125 mL</td>
		</tr>
		<tr>
			<td>Collagenase</td>
			<td>35&#8211;40 mg, add BSA up to 100 mg</td>
		</tr>
		<tr>
			<td colspan="2">Percoll solution</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td>Concentration</td>
		</tr>
		<tr>
			<td>Percoll</td>
			<td>22.5 mL</td>
		</tr>
		<tr>
			<td>PBS (10x), PH=7.4</td>
			<td>2.5 mL</td>
		</tr>
		<tr>
			<td colspan="2">Isoaltion/Staining Buffer</td>
		</tr>
		<tr>
			<td>Reagents</td>
			<td>Concentration</td>
		</tr>
		<tr>
			<td>MACS rinsing buffer</td>
			<td>1250 mL</td>
		</tr>
		<tr>
			<td>BSA stock</td>
			<td>125 mL</td>
		</tr>
	</tbody>
</table>
Table 1: Buffers recipe. The table comprises the composition of the buffers and solutions used in this study.

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. ...

isolation-characterization-mouse-primary-liver-sinusoidal-endothelial

Nanyang Technological University

Research

JoVE Journal

Biology

6.4K 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

Purification of Hepatocytes and Sinusoidal Endothelial Cells from Mouse Liver Perfusion

This protocol demonstrates a method for obtaining high yield and viability for mouse hepatocytes and sinusoidal endothelial cells (SECs) suitable for culturing or for obtaining cell lysates. In this protocol, the portal vein is used as the site for catheterization, rather than the vena cava, as this limits contamination of other possible cell types in the final liver preparation. No special instrumentation is required throughout the procedure. A water bath is used as a source of heat to maintain the temperature of all the buffers and solutions. A standard peristaltic pump is used to drive the fluid, and a refrigerated table-top centrifuge is required for the centrifugation procedures. The only limitation of this technique is the placement of the catheter within the portal vein, which is challenging on some of the mice in the 18 - 25 g size range. An advantage of this technique is that only one vein is utilized for the perfusion and the access to the vein is quick, which minimizes ischemia and reperfusion of the liver that reduces hepatic cell viability. Another advantage to this protocol is that it is easy to distinguish live from dead hepatocytes by eyesight due to the difference in cellular density during the centrifugation steps. Cells from this protocol may be used in cell culture for any downstream application as well as processed for any biochemical assessment.

The goal of this protocol is to obtain high viability and high yield of hepatocytes and sinusoidal endothelial cells from liver. This is accomplished by perfusing the liver with a type IV collagenase solution via the portal vein, followed by differential centrifugation to obtain hepatocytes and sinusoidal endothelial cells.

This protocol demonstrates a method for obtaining high yield and viability for mouse hepatocytes and sinusoidal endothelial cells (SECs) suitable for ...

Biochemistry

Isolation of Primary Murine Brain Microvascular Endothelial Cells

The blood-brain-barrier is ultrastructurally assembled by a monolayer of brain microvascular endothelial cells (BMEC) interconnected by a junctional complex of tight and adherens junctions. Together with other cell-types such as astrocytes or pericytes, they form the neurovascular unit (NVU), which specifically regulates the interchange of fluids, molecules and cells between the peripheral blood and the CNS. Through this complex and dynamic system BMECs are involved in various processes maintaining the homeostasis of the CNS. A dysfunction of the BBB is observed as an essential step in the pathogenesis of many severe CNS diseases. However, specific and targeted therapies are very limited, as the underlying mechanisms are still far from being understood. 
Animal and in vitro models have been extensively used to gain in-depth understanding of complex physiological and pathophysiological processes. By reduction and simplification it is possible to focus the investigation on the subject of interest and to exclude a variety of confounding factors. However, comparability and transferability are also reduced in model systems, which have to be taken into account for evaluation. The most common animal models are based on mice, among other reasons, mainly due to the constantly increasing possibilities of methodology. In vitro studies of isolated murine BMECs might enable an in-depth analysis of their properties and of the blood-brain-barrier under physiological and pathophysiological conditions. Further insights into the complex mechanisms at the BBB potentially provide the basis for new therapeutic strategies.
This protocol describes a method to isolate primary murine microvascular endothelial cells by a sequence of physical and chemical purification steps. Special considerations for purity and cultivation of MBMECs as well as quality control, potential applications and limitations are discussed.

Brain microvascular endothelial cells (BMEC) are interconnected by specific junctional proteins forming a highly regulated barrier separating blood and the central nervous system (CNS), the so-called blood-brain-barrier (BBB). The isolation of primary murine brain microvascular endothelial cells, as discussed in this protocol, enables detailed in vitro studies of the BBB.

The blood-brain-barrier is ultrastructurally assembled by a monolayer of brain microvascular endothelial cells (BMEC) interconnected by a junctional ...

Neuroscience

Isolation of Primary Murine Skeletal Muscle Microvascular Endothelial Cells

The endothelial cells of skeletal muscle capillaries (muscle microvascular endothelial cells, MMEC) build up the barrier between blood stream and skeletal muscles regulating the exchange of fluids and nutrients as well as the immune response against infectious agents by controlling immune cell migration. For these functions, MMEC form a functional &#34;myovascular unit&#34; (MVU), with further cell types, such as fibroblasts, pericytes and skeletal muscle cells. Consequently, a dysfunction of MMEC and therefore the MVU contributes to a vast variety of myopathies. However, regulatory mechanisms of MMEC in health and disease remain insufficiently understood and their elucidation precedes more specific treatments for myopathies. The isolation and in-depth investigation of primary MMEC functions in the context of the MVU might facilitate a better understanding of these processes.
This article provides a protocol to isolate primary murine MMEC of the skeletal muscle by mechanical and enzymatic dissociation including purification and culture maintenance steps.

Microvascular endothelial cells of skeletal muscles (MMEC) shape the inner wall of muscle capillaries and regulate both, exchange of fluids/molecules and migration of (immune) cells between muscle tissue and blood. Isolation of primary murine MMEC, as described here, enables comprehensive in vitro investigations of the &#34;myovascular unit&#34;.

The endothelial cells of skeletal muscle capillaries (muscle microvascular endothelial cells, MMEC) build up the barrier between blood stream and skeletal ...

Isolation of Primary Mouse Hepatocytes for Nascent Protein Synthesis Analysis by Non-radioactive L-azidohomoalanine Labeling Method

Hepatocytes are parenchymal cells of the liver and engage multiple metabolic functions, including synthesis and secretion of proteins essential for systemic energy homeostasis. Primary hepatocytes isolated from the murine liver constitute a valuable biological tool to understand the functional properties or alterations occurring in the liver. Herein we describe a method for the isolation and culture of primary mouse hepatocytes by performing a two-step collagenase perfusion technique and discuss their utilization for investigating protein metabolism. The liver of an adult mouse is sequentially perfused with ethylene glycol-bis tetraacetic acid (EGTA) and collagenase, followed by the isolation of hepatocytes with the density gradient buffer. These isolated hepatocytes are viable on culture plates and maintain the majority of endowed characteristics of hepatocytes. These hepatocytes can be used for assessments of protein metabolism including nascent protein synthesis with non-radioactive reagents. We show that the isolated hepatocytes are readily controlled and comprise a higher quality and volume stability of protein synthesis linked to energy metabolism by utilizing the chemo-selective ligation reaction with a Tetramethylrhodamine (TAMRA) protein detection method and western blotting analyses. Therefore, this method is valuable for investigating hepatic nascent protein synthesis linked to energy homeostasis. The following protocol outlines the materials and methods for the isolation of high-quality primary mouse hepatocytes and detection of nascent protein synthesis.

Here, we present a protocol for the isolation of healthy and functional primary mouse hepatocytes. Instructions for detecting hepatic nascent protein synthesis by non-radioactive labeling substrate were provided to help understand the mechanisms underlying protein synthesis in the context of energy-metabolism homeostasis in the liver.

Hepatocytes are parenchymal cells of the liver and engage multiple metabolic functions, including synthesis and secretion of proteins essential for ...

Medicine

Isolation of Primary Mouse Lung Endothelial Cells

Endothelial cells play critical roles in the regulation of vascular tone, immunity, coagulation, and permeability. Endothelial dysfunction occurs in medical conditions including diabetes, atherosclerosis, sepsis, and acute lung injury. A reliable and reproducible method to isolate pure endothelial cells from mice is needed to investigate the role of endothelial cells in the pathogenesis of these and other conditions. In this protocol, lung microvascular endothelial cells were prepared from 5-7 day old neonatal mouse pups. Lungs are harvested, minced, and enzymatically digested with collagenase I, and released cells are cultured overnight. Endothelial cells are then selected using anti-PECAM1 (CD-31) IgG conjugated to magnetic beads, and cells again are cultured to confluence. A secondary cell selection then occurs with anti-ICAM2 (CD-102) IgG conjugated to magnetic beads to increase the purity of the endothelial cells, and the cells again are cultured to confluence. The entire process takes approximately 7-10 days before the cells can be used for experimentation. This simple protocol yields highly pure (purity &#62;92%) endothelial cells that can be immediately used for in vitro studies, including the studies focused on endothelial cytokine and chemokine production, leukocyte-endothelial interactions, endothelial coagulation pathways, and endothelial permeability. With many knockouts and transgenic mouse lines available, this procedure also lends itself to understanding the function of specific genes expressed by endothelial cells in healthy and pathologic responses to injury, infection, and inflammation.

In this article, primary lung endothelial cells were isolated and cultured from neonatal mice.

Endothelial cells play critical roles in the regulation of vascular tone, immunity, coagulation, and permeability. Endothelial dysfunction occurs in ...

Immunology and Infection

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 and 3D Collagen Sandwich Culture of Primary Mouse Hepatocytes to Study the Role of Cytoskeleton in Bile Canalicular Formation In Vitro

Hepatocytes are the central cells of the liver responsible for its metabolic function. As such, they form a uniquely polarized epithelium, in which two or more hepatocytes contribute apical membranes to form a bile canalicular network through which bile is secreted. Hepatocyte polarization is essential for correct canalicular formation and depends on interactions between the hepatocyte cytoskeleton, cell-cell contacts, and the extracellular matrix. In vitro studies of hepatocyte cytoskeleton involvement in canaliculi formation and its response to pathological situations are handicapped by the lack of cell culture, which would closely resemble the canaliculi network structure in vivo. Described here is a protocol for the isolation of mouse hepatocytes from the adult mouse liver using a modified collagenase perfusion technique. Also described is the production of culture in a 3D collagen sandwich setting, which is used for immunolabeling of cytoskeletal components to study bile canalicular formation and its response to treatments in vitro. It is shown that hepatocyte 3D collagen sandwich cultures respond to treatments with toxins (ethanol) or actin cytoskeleton altering drugs (e.g., blebbistatin) and serve as a valuable tool for in vitro studies of bile canaliculi formation and function.

Presented here is a protocol for the isolation of mouse hepatocytes from adult mouse livers using a modified collagenase perfusion technique. Also described is the long-term culture of hepatocytes in a 3D collagen sandwich setting as well as immunolabeling of the cytoskeletal components to study bile canalicular formation and its response to treatment.

Hepatocytes are the central cells of the liver responsible for its metabolic function. As such, they form a uniquely polarized epithelium, in which two or ...

An Improved Time- and Labor- Efficient Protocol for Mouse Primary Hepatocyte Isolation

Primary hepatocytes are used extensively in liver in vitro research, especially in glucose metabolism studies. A base technique has been adapted based on different needs, like time, labor, cost, and primary hepatocyte usage, resulting in various primary hepatocyte isolation protocols. However, the numerous steps and time-consuming reagent preparations in primary hepatocyte isolation are major drawbacks for efficiency. After comparing different protocols for their pros and cons, the advantages of each were combined, and a rapid and efficient primary hepatocyte isolation protocol was formulated. Within only ~35 min, this protocol could yield as much, if not more, healthy primary hepatocytes as other protocols. Further, glucose metabolism experiments performed using the isolated primary hepatocytes validated the usefulness of this protocol in in vitro liver metabolism studies. We also extensively reviewed and analyzed the significance and purpose of each step in this study so that future researchers can further optimize this protocol based on needs.

Primary hepatocytes are a valuable tool to study liver response and metabolism in vitro. Utilizing commercially available reagents, an improved time- and labor-efficient protocol for mouse primary hepatocyte isolation was developed.

Primary hepatocytes are used extensively in liver in vitro research, especially in glucose metabolism studies. A base technique has been adapted based on ...