*Download the PDF version of the entire protocol here
*PDF version of this protocol updated 2010-11-21; uploaded 2012-04-07. Minor differences in Troubleshooting section compared to website.
- Prepare all necessary reagents. For each mouse, you will need (see details for reagents in materials):
- 60-70mL HBSS for initial flushing/washing; 37C
- 90mL DMEM-low glucose (digestion medium); 37C
- ~120mL isolation medium; 4C
- Sufficient collagenase to make 90mL of digestion solution at 100 CDU/mL; store desiccated at 4C until ready to use
- Again, note that these quantities are based upon the C57 strain of mice weighing 20-40g.
- Preferably under the hood, dispense pre-measured collagenase aliquot into digestion medium, swirl gently, and allow to dissolve for 30-60 minutes at 37C. We have observed that the efficiency of the collagenase is more consistent (based on total cell yield) when it is allowed to dissolve for at least 30 minutes, even though it appears to go into solution instantly. Put HBSS into waterbath to warm at this time as well.
- Prepare work surface. A sterile environment is preferable, but not necessary for the initial isolation. Spray down all non-porous surfaces with 70-75% ethanol.
- Rinse out the pump/tubing by running 70-75% ethanol through and allowing it to sit for >20 minutes. Remove residual ethanol by rinsing twice with autoclaved water. Make sure the bubble trap is thoroughly cleaned and rinsed as well.
- Take two absorbent bench pads and lay one pad down on the bench; use the other pad to cover a hard board (e.g. a stiff piece of cardboard or Styrofoam). As this is a non-recirculating perfusion, all liquids will run to waste; it is advisable to fold the bench pad in such a manner that excess liquids will be trapped within, to avoid making a mess.
- If desired, you may set the board inside a shallow-wall container. This is optional for the quantity of liquid used during a mouse perfusion (the pads should provide sufficient holding capacity), but may be necessary when working with larger volumes of liquid (i.e. rat perfusions).
- You may find it more comfortable to elevate the hard board slightly, as this will aid draining of liquids, as well as bring the work surface closer to your eyes.
- Bring a 10cm culture dish (sterile) to the bench. This will be used to hold the digested liver once the perfusion is complete.
- If using a reusable Dual Mfg 75 micron filter, make sure that it is cleaned/sterilized beforehand 
- Put the following on ice:
- Bottle containing isolation medium (~120mL)
- 50mL sterile conical tube
- Ensure the following are warm:
- Digestion medium w/collagenase
- Attach your cannula to the free end of the tubing. You may find it preferable to cut the "wings" off of the butterfly cannula, as it reduces the effort needed to hold the needle tip, as well as offering better handling.
- Immediately before knocking out your mouse, pour warmed HBSS into the reservoir and prime the pump, making sure that the bubble trap
contains sufficient HBSS (at least 1/4, but no more than 1/3 full) and that there are no air bubbles in the system, particularly past the
bubble trap. If bubbles are present in the tubing past the bubble trap, turn on the pump and elevate the affected section of tubing to drive
the bubbles out. Do not be concerned about losing a few mL of HBSS.
- After priming the pump and ensuring that there are no visible air bubbles, turn the flow rate to a slow drip (1-2mL/min) and shut the pump off.
Cannulation, perfusion, and digestion
- Anesthetize the mouse. Isofluorane is recommended, as it has minimal impact on liver metabolism.
- Secure the mouse, ventral side up, to the working platform; tape or pin down all four limbs. Tape is preferred to minimize distress, should the animal wake from anesthesia.
- Thoroughly clean the abdomen/chest region, using EtOH + detergent, and/or iodine. Work quickly but thoroughly at this step, as flora from the fur is the first potential source of contamination. If the animal defecates, be sure to clean it up and disinfect the area.
- Pick up a pair of scissors and your straight forceps. Make an incision in the lower abdomen, cutting through the fur and muscle layer; try to puncture through the muscle layer as soon as you make your incision (this is why sharp-tipped scissors are recommended). Do NOT nick any internal organs, particularly the intestines. Pull up on the fur while cutting to minimize this risk.
- Continue to cut vertically until the liver, portal vein (PV), and inferior vena cava (IVC) are sufficiently exposed. It is easiest to make alternating cuts on either side of the skin while pulling up. Make a final incision on the left (your left) side of the abdomen, near the navel region, so that blood can readily drain once you cannulate and perfuse.
- Start the pump (which should be set at a low flow rate already) and wait until HBSS begins to flow from the tip of the cannula; with HBSS flowing, swiftly, and in one motion, insert the cannula into the PV.
- If performed properly, the liver should instantly begin to blanch. Once you have confirmed that the cannulation is successful, quickly cut the IVC to relieve pressure in the system and allow perfusate to drain to waste (having an assistant perform this task is highly recommended). Once the IVC has been cut, the liver should finish blanching, and become pale in color. Blanching is easiest to monitor under bright, directed lighting.
- If the liver does NOT blanch instantly, particularly after the IVC has been cut, there are two likely sources of error: occult air bubbles blocking microcapillaries, or user error (i.e. missing the portal vein and cannulating a pancreatic duct instead). The former is more common, which is why it is essential that HBSS is allowed to drip from the cannula tip during perfusion (the continued presence of liquid displaces any unseen air in the bevel/tip region of the cannula.
- Alternatively, you may perform a retrograde perfusion by cannulating the IVC and cutting the PV for drainage. Klaunig et al. (1980) found that PV cannulation resulted in higher yields than IVC cannulation, although the difference was not dramatic, nor were viability estimates significantly different. Due to the significantly larger size of the IVC, you may find it easier to work with; our experience, however, is all based on PV cannulation with IVC drainage.
- Once the IVC has been cut, increase the flow rate to 7-9mL/minute. Allow the entire volume of HBSS to perfuse through the liver.
The flow may be increased in increments of 1-2mL/second (have your assistant monitor and adjust the flow).
- A quick test for successful cannulation can be performed by applying light pressure on the IVC; at >8mL/minute, all lobes of the liver should instantly begin to swell.
- When the reservoir (NOT the bubble trap) is just about to run out of HBSS, pour in 70mL of the digestion medium
(fill the reservoir to the top). It is not necessary, nor is it recommended to stop the pump. There will obviously be some
dilution (by HBSS) of the initial few milliliters of digestion medium, but this is inconsequential. Pour the remaining digestion
solution (~20mL) into the 10cm plate. Close the plate quickly (again, these are tasks that are greatly facilitated by a second pair of hands).
- One technique which may increase yield and reduce total digestion time is to periodically (5-10 times during digestion) apply pressure to the IVC for 5-second intervals (as mentioned in step 7a). This will cause the liver to swell, and the increased pressure during the clamping aids in dissociation, and therefore, final yield. We strongly recommend this be performed as standard operating procedure.
- As digestion progresses, you should see the liver begin to swell. This presumably occurs as a result of collagenase breaking down the
elastic structure of the liver. When this swelling starts, you should be very close to the end of digestion.
- If you performed step 8a, you cannot use this as an indicator, as the liver will swell and contract as you apply and release pressure. Therefore, to monitor completeness of digestion, note that as digestion continues, the liver will contract less with each IVC clamping cycle, due to progressive loss of elasticity.
- There is no definite rule as to when digestion is complete. Generally, once the liver has begun to swell, 1-3 minutes of additional digestion should suffice. You may or may not see small clear/transparent sections on the lower lobes; furthermore, the liver will take on the texture of a wet piece of cloth, and appear almost soggy.
- The extent to which you allow the liver to digest is almost entirely dependent upon how skillful you are in excising the organ; one or two minutes of prolonged digestion will not affect viability if a sufficiently gentle (i.e. low tryptic activity) collagenase is used.
- When you are satisfied with the digestion, turn off the pump and carefully remove the cannula.
Extraction and purification
- Using your straight-tipped forceps, gently expose the central region the liver by carefully working through the lobes. Locate the bundle of fibers connecting the lobes. At this point, switch to your curved, fine-tipped forceps and securely grip the center of this bundle. Do not rush this step, as it is essential that you have a solid hold on the liver for a complete excision.
- Gripping the liver, pull the organ away and towards you (try to get behind the liver, and free the lobes from the chest cavity), exposing the chest cavity and the various connective tissues holding the liver in place. Carefully cut these connections, slowly pulling the liver forward and out as you cut. You may wish to switch to a fresh pair of scissors for this step, as your previous pair may be covered in blood and fur. Take care to avoid nicking any internal organs.
- Once a sufficient number of connections have been severed, you should, with moderate effort, be able to pull the liver free. Do NOT tear the gall bladder; if you wish to do so, remove the gall bladder at this step.
- Immediately place the liver into the 10cm dish containing digestion medium, and cover the dish (assistant).
- [Preferably in the hood] Using two pairs of forceps (either another sterile set, or the same set, cleaned with EtOH and/or flamed),
tear apart the lobes of the liver; if still present, avoid disrupting the gall bladder.
- If the liver has been properly digested, you should see the medium turn cloudy as you tear, and the liver should mostly dissolve into the medium. Ideally, all that will be left is a stem of connective tissue, but often, you will have some residual lobe matter as well. If, upon tearing, you end up with chunks of liver and little to no clouding of the media, something went wrong during the digestion. See troubleshooting section.
- Once torn apart, grab the remaining section of the liver and shake gently to free residual cells. Discard any solid
particles that remain, including the gall bladder (if you chose not to remove it in Step 3, the gall bladder should remain intact during this process; if it burst,
you either used too much force, or accidentally tore it open with your forceps).
- A torn gall bladder should not have any adverse effect on cell viability or function, but this is not something we have tested. In the interest of batch-to-batch consistency, try to minimize gall bladder bursting.
- Reminder: all steps involving handling of hepatocytes must be performed with care, using sterile equipment. The cells are very fragile at this point and are susceptible to shearing damage.
- In the same 10cm dish, triturate the suspension three times using a 25mL serological pipette.
- If you are using a Dual Mfg stainless steel filter, make sure you pre-wet the bottom of the filter at this time, or even before you start tearing the liver. Simply pipette 2-3 mL of isolation medium onto the bottom of the filter, swirl until the entire surface area is wet, and discard the remaining medium.
- Filter the suspension through a 70-75-micron membrane. If you are using the Dual Mfg filter this step should be very fast; with BD
disposable falcon tube nylon filters, this will take some time, and possibly a few filters (a sterile eyedropper pipette might be easier
for falcon tube strainers).
- If using the Dual Mfg filter, your cells will be in a 95mm dish; transfer them into a sterile, clean 50mL conical tube. For the falcon tube strainers, your cells will already be inside a 50mL conical tube and ready to use.
- Spin at 4C for two minutes at 50 xg in a swinging-arm centrifuge. The centrifuge does not have to be refrigerated, although this is preferred.
- (in the hood) Aspirate the supernatant (should be cloudy/opaque) using a sterile glass Pasteur pipette and add 25mL of cold isolation medium. Triturate gently a few times to break up the cell mass at the bottom and resuspend.
- Repeat steps 9 and 10 two more times, for a total of three washes. The medium should be almost completely clear by the second wash, and clear by the third.
- After the final spin, aspirate the medium and add 25-45mL of cold isolation medium. You want your cells to be concentrated enough so that you do not have to re-spin them prior to plating, but not so concentrated that they will be difficult to count. Resuspend cells gently with a 25mL serological pipette. Remove an 80uL aliquot and transfer it into a microfuge tube for counting/trypan blue staining.
Staining, quantitation, and plating
- Add 20uL of 0.4% trypan blue to your 80uL cell suspension aliquot. Pipette up and down several times to mix.
- Allow to stain for ~1 minute at room temperature.
- Pipette up and down again, and take a 10uL aliquot and dispense onto a hemacytometer.
- Count all non-blue, non-blebby cells. Note that trypan blue OVERESTIMATES viability. Cells which take up
the dye are certainly non-viable; however, cells which do not stain may nonetheless be damaged and useless. In
our experience, viable, healthy hepatocytes have a bright, clear, smooth appearance and are relatively small and
rounded up. Dead/damaged cells generally are swollen and appear rough and granular. Viability MUST be over 85% at
minimum to ensure batch-to-batch consistency.
- From a typical 8-12 week-old C57 mouse liver, you can expect total yields on the order of 30-50 million cells. The average yield should be ~40 million, and yields below 30 million indicate that something was wrong with the procedure (most likely attributable to digestion/cannulation).
- After counting cells (make sure you correct for the 25% dilution factor due to trypan blue), make a working dilution for plating. A general estimate for dilution is to add three million (viable) cells per plate, at 10mL total volume per plate, or 300,000 cells/mL.
- As a general guideline, we recommend:
- 8mL/plate for 10cm plates (7-9mL OK)
- 1.7mL/well for 6-well plates (1.5-2.0mL OK)
- 800uL/well for 12-well plates (700-820uL OK)
- 360uL/well for 24-well plates (340-450uL OK)
These volumes have been found to give the most consistent plating results, as they minimize the tendency for cells to cluster in the middle or on the edges. See Troubleshooting #13 for a special note about 24-well or smaller plates.
- Before plating all cells, you should plate a single well and double-check under the microscope that the actual density is close to what you assume it to be, based on your counting. Make sure to shake the plate thoroughly (in a back-and-forth, and NEVER in a circular fashion) and allow the cells to settle for ~30 seconds before looking at them. In a typical prep of >90% viable cells, you should aim for approximately 60-70% confluence at this time. This level of confluence allows for cell-cell contact, while maintaining sufficient space for the hepatocytes to grow to their full size, and will yield a final confluence of 90-95%.
- After confirming that the cell density is OK and making any necessary adjustments, plate cells down. Make sure you resuspend cells after every minute or two, as hepatocytes are quite dense and settle rapidly. Generally, you should resuspend cells after every two 12-well plates, or every (one) 24-well plate.
- Thoroughly shake your plates in a linear fashion before placing them in the incubator; this step is critical, as the cells tend to sink straight down and attach, and failure to spread them out in a homogenous monolayer at this time will result in a patchy, inconsistent lawn.
- Allow cells to attach for 45-60 minutes @37C after plating.
Post-plating and overnight culture
- After cells have attached, wash once with DMEM-low and add back culture media (with 10% FBS) for 3-4 hours.
- At this time, if your cells are healthy, nuclei should start to become visible under low power. Cells will still be rounded, but somewhat larger; note that clusters of cells tend to recover more rapidly than single cells.
- Use caution when changing the media at this time, as the hepatocytes are still relatively fragile and can be easily damaged or disrupted by direct contact; pipette only down the side of the well, and never directly on top of the cells.
- After the first 4-5 hours of plating, we suggest that cells be kept in serum-free medium. This helps to maintain their morphology, and there are no adverse effects to serum-free medium. The most critical time during which serum must be present is the first hour of plating, where is has been suggested that the various proteins found in serum aid in attachment.