Purification of Hepatocytes and Sinusoidal Endothelial Cells from Mouse Liver Perfusion

All animal procedures outlined in this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Lincoln – Nebraska under protocol #1435.

1. Preparation

1. Preparation of solutions and culture media
   1. Prepare all culture media and buffer according to the Table of Materials.
2. Preparation of instruments
   1. Set up a water bath (>10 L) at 42 °C, a peristaltic pump with variable speed, and clamps for holding flasks. Prepare curved scissors and forceps (Figure 1).
   2. On or adjacent to the water bath, set up a baking sheet (about 38 x 26 cm) with a Styrofoam pad (50 mL conical rack), overlaid with an absorbent underpad cut into a 20 x 20 cm piece (Figure 1).
   3. Place a piece of masking tape (5 – 8 cm) nearby. Place a 20 cm polyester sewing thread nearby that is cut and ready to use. Obtain a plastic catheter with a retractable needle (7 mm x 19 mm, Figure 1).
3. Preparation of glassware
   1. Using clamps, place and immobilize a 1 L flask in the water bath containing 200 mL of 1x PBS with a stopper that has two 1 mL pipettes going into the solution. The rubber cork is notched to allow recirculation of fluid in a closed circuit.
   2. With another clamp, place a 125 mL flask containing 45 mL of Buffer 2 along with a stopper that has two 1 mL pipettes, one of which is in the liquid near the bottom of the flask and the other which stays above the liquid and blows oxygen into the flask.
      NOTE: Each pipette in the stoppers will have a quick disconnect adaptor attached to the end for easy switching of the tubing. Oxygen will be gently flowing into both flasks via the pipette in the stopper that is not immersed in the liquid.
   3. Set aside two sterile crystallizing dishes.

2. Animal Procedure

1. Warm up the PBS and Buffer 2 solutions to 42 °C in the water bath and circulate PBS for at least 20 mL/min in the tubing in a closed circuit to keep the liquid lines warm. Immerse any excess tubing in the water bath to remain as close as possible to 42 °C. Drain the male end of the tubing into the 1 L flask containing PBS.
2. Place a cotton ball on the absorbent underpad and add about 2 mL of 30% isoflurane (made up in polyethylene glycol 200 which decreases the evaporation rate of isoflurane)  on the cotton ball using a transfer pipette.
3. Pick up the mouse by the tail, place it in one of the crystallizing dishes, and then quickly overturn the dish onto the cotton ball so that the mouse has a small space in which to inhale the anesthesia. Observe the breathing rate of the mouse and ensure the mouse is effectively under anesthesia.
   NOTE: The breathing rate should be slower and deeper, the tail flaccid, and the paws nonresponsive to the pinch test under anesthesia; see IACUC guidelines for additional details.
4. While the mouse is going under anesthesia (this takes 1-2 min), prepare the barrel of a 10 mL syringe by pulling out the plunger and inserting a small cotton ball. Add 1–2 mL of 30% isoflurane with a transfer pipette to the cotton ball inside the barrel and place the barrel open-end down on a table while waiting for the mouse to become unconscious.
5. Quickly take off the crystallizing dish, flip the mouse on its back and place the syringe barrel over its nose. Double check the breathing rate, toe pinch, and flaccid tail. Any responses to the toe pinch and flaccid tail indicates that the mouse is not fully under anesthesia. Keep the syringe barrel over the snout to maintain unconsciousness.
6. Place thumb tacks through the paws of the mouse, with the limbs outstretched in supine position.Wet the abdomen and rib cage with 70% isopropanol or ethanol.
7. With the straight forceps in one hand, lift up the skin near the base of the abdomen. With scissors in the other hand, cut the tented skin and peritoneum. The incision should be horizontal or across the base of the tented skin. Be sure to cut through all layers of skin and peritoneum in order to gain access to the gut. Cut laterally around the abdomen up to the rib cage on both sides without nicking any of the organs.
8. Cut off the flap of skin by cutting across the lower rib cage. Move the intestines to the right with the back of the forceps to expose the portal vein.
   NOTE: Bleeding from the animal should be minimal; the animal should still be breathing and under deep anesthesia.
9. Place the closed curved forceps underneath the portal vein between the liver and superior pancreaticoduodenal vein (Figure 3 arrow).
10. Open the forceps while underneath the portal vein. Grasp the thread and carefully pull it through so that it is centered underneath the portal vein. Tie an overhand knot around the portal vein without cinching it down.
11. Place the curved forceps under the portal vein and gently pull it towards the tail of the mouse to straighten out the vein (Figure 4A).
12. With the catheter in the other hand, place the bevel of the needle face-up and parallel with the lower part of the portal vein near the forceps (Figure 4B).
13. Gently puncture the vein with the needle (Figure 4C). Ensure the bevel of the needle is in the lumen of the vein. Retract the spring-loaded needle and continue to push the polymer catheter through the vein until the bevel is near the venous branched area. This is within the liver. If the catheter is placed correctly, a back-flow of blood will be visible. (Figure 4D).
14. Tighten the overhand knot and pull it down on the catheter to help stabilize it. Immediately turn down the pump rate from 20 mL/min to 4 mL/min and cut another major blood vessel for drainage. Cut the aorta abdominalis for optimal results.
15. Place the male end of the tubing to the female end of the catheter (Figure 5A). Ensure that there are no air bubbles in the line. Be careful that the catheter is not either pushed into the liver or out of the vein. Placement of the thread is not essential, though it helps prevent back-flow out of the vein and helps keep the catheter in the correct position (Figure 5B, C).
16. Use masking tape to immobilize the tubing onto the underpad. Use the straight forceps to squeeze the effluent blood vessel to inflate the liver a few times to ensure all the blood has drained out (Figure 5D).
    NOTE: Standard laboratory tape may not work; however, masking tape will remain stuck to the underpad, even in wet conditions.

3. Liver Perfusion

1. While the liver is flushing with PBS, measure out about 24 mg of Collagenase Type IV and place it in the flask containing 45 mL of Buffer 2 in the water bath. Be sure to swirl the liquid in the flask so that the collagenase is fully dissolved and place the flask back in the clamp so that the liquid containing portion of the flask is fully submerged in the water.
2. Observe the change in color of the liver as it is flushed with PBS. Change the outflow tubing from the 1 L flask to the 125 mL flask. Do not allow air bubbles to flow into the liver.
3. Once the perfusion buffer reaches the liver, briefly squeeze tight the effluent blood vessel to build up some pressure within the vessel and allow this liquid to fill all lobes of the liver. Be sure not to cut off drainage too long, as that may burst the thin connective tissue surrounding the liver (Glisson's capsule) and destroy the perfusion or flow of liquid within the capillary bed of the liver.
4. Allow Buffer 2 with collagenase to perfuse through the liver until all 45 mL have flowed through the tissue.
   NOTE: If successful, the Glisson's capsule should be separated from the parenchyma or the liver tissue and the liver itself should appear amorphous.
5. Add about 10 mL of Buffer 1 to the other crystallizing dish and place it next to the mouse.
6. Remove the catheter and turn the pump off.
7. With the straight forceps and scissors, cut the liver from the mouse. If the collagenase digestion was very efficient, it may be necessary to have a clean, sterile spoon on hand to scoop the liver from the mouse and place it in Buffer 1.

4. Hepatocyte Purification

NOTE: Manipulation of the cells are performed in a sterile tissue culture hood to limit contamination if the cells are to be cultured in all subsequent steps.

1. Using sterile technique, grab the liver with forceps and gently shake the cells from the liver. Tear apart or pull back the Glisson's capsule: the buffer will become opaque as cells are shaken from the liver.
2. Pour off the cell solution into a 50 mL conical with a 100 µm filter placed over the top.
3. Add more Buffer 1 to the liver remnants and continue to shake out cells. Continue until the liver appears devoid of cells or when the undigested portions of the liver do not yield anymore cells.
4. Pour the cellular liquid into another 50 mL conical containing a 40 µm filter.
5. Centrifuge the conical in a swinging bucket rotor at 100 x g for 3 min.
   ​NOTE: Do not use the maximum brake as this may dislodge the cell pellet. Use the brake at 80% which is sufficient for all remaining centrifugation steps at 4 °C
6. Pour off the supernatant (which contains NPCs and dead hepatocytes) into a clean conical and place it on ice. Ensure this is done in one motion to observe pellet adherence.
7. Resuspend the pellet in 40 mL of Buffer 1. Repeat steps 4.5 and 4.6. Resuspend the pellet in 40 mL of Buffer 3. Repeat steps 4.5 and 4.6 twice.
8. Resuspend the purified hepatocytes in warm DMEM + 10% FBS + 2x Penicillin-Streptomycin (Pen/Strep).
9. If the cells are cultured on collagen coated plates, allow them to adhere for 1 h and replace the media at least once as this will prevent any nascent bacterial contaminants.
   1. Make the collagen coated plates by incubating them with 0.05% collagen in 0.001% acetic acid for at least 1 h at 37 °C, and then washing with 1x PBS.
10. If the hepatocyte viability is low and there is a desire to obtain purified high-viable hepatocytes then use the following protocol adapted from Kreamer et al¹⁰.
    1. Mix 9 volumes of PVP solution (Percoll, a 23% w/w solution of water and 15 – 30 nm colloidal silica particles coated in polyvinylpyrrolidone for density centrifugation) and 1 volume of 10x HBSS to make an iso-osmotic PVP solution (SIP).
    2. Add 24 mL of SIP to a sterile 50 mL conical which may be stored in these conditions for up to 2 months at 4 °C.
    3. Adjust the hepatocyte concentration to 5 – 10 × 10⁶ cells/mL with culture medium such as the one noted in step 4.8.
    4. Add 25 mL of cell suspension to each 24 mL SIP containing conical and mix by gentle inversion. The density of this solution is 1.06 g/mL.
    5. Centrifuge the conical at 50 x g for 10 min at 4 °C.
    6. Aspirate the SIP and flocculant, and resuspend the cells in HBSS.
    7. Wash the cells by centrifugation at 50 x g for 10 min at 4 °C.
    8. Repeat step 4.10.6 once more and then resuspend cells in growth medium.

5. SEC Purification

1. Pellet the cells in the supernatants from step 4.6 by spinning the tubes at 163 x g for 10 min. Discard the supernatants from the tubes and resuspend all of cell pellets in 5 mL of RPMI without serum and pool them together in one tube.
2. Once all pellets have been pooled, add RPMI to final volume of 35 mL.
3. Centrifuge the pooled conical at 25 x g for 3 min. Carefully aspirate the top 25 mL of RPMI with a 25 mL pipette and place this media containing NPCs in a clean 50 mL conical on ice.
4. Add 25 mL of fresh RPMI back to the tube and resuspend the pellet.
5. Repeat step 5.3 for a second wash and pool the supernatant. Discard the remaining 10 mL of media and pellet (the pellet consists mostly of dead hepatocytes). Centrifuge the pooled supernatant at 163 x g for 10 min.
6. During the centrifugation, prepare the PVP solution gradients in a 50 mL conical.
7. Add 15 mL of 50% PVP solution to the 50 mL conical followed by 20 mL of 25% Percoll overlaid with a pipettor set to the slowest ejection speed. Be sure to observe a refraction line at the 15 mL mark that demarcates the two layers, as this is where the SECs and KCs will aggregate after the centrifugation.
8. Resuspend the cells previously pelleted in step 5.5 in 10 mL of cold RPMI and overlay them onto the PVP solution gradient. Be sure to maintain a clear demarcation between the two layers. Centrifuge the gradient at 805 x g for 20 min with a brake set at 50%.
9. Aspirate from the top down direction to the 20 mL mark on the tube and discard this material. With a 5 mL transfer pipette, collect the SECs and KCs that are located at the 25/50% interface. Discard any brown clumps of cells as these are dead hepatocytes.
10. Place the cells in a 50 mL conical and add RPMI up to the 50 mL mark to dilute out the PVP solution. Centrifuge at 200 x g for 10 min, with 80% brake.
11. Aspirate and discard the supernatant and resuspend the pellet while washing off the cone of the conical in 12 mL of warm RPMI.
12. Separate the SECs from KCs by placing the supernatant in a polystyrene Petri dish and incubate in a humidified tissue culture incubator for 8 min at room temperature.
13. Aspirate the media with a 25 mL pipette and rinse the plate with the same media, with the pipettor set at the lowest speed to collect the remaining SECs while the KCs adhere to the plate.
14. Centrifuge the SECs at 200 x g for 10 min and resuspend in RPMI + 5% FBS + Pen/Strep. Then titer the cells, and place in collagen-coated culture dishes.