| | Re: mouse hepatocytes isolation
here is my protocol. I usually get 40-60% live hepatocytes.
(I cut and pasted my WORD file, so the formating may be off)
Mouse Hepatocyte Isolation
Liver Perfusion Media Invitrogen# 17701-038 500mL. store 4oC
Liver Digest Media Invitrogen# 17703-034 500mL. store -20oC
Thaw immediately before use. Store in 35mL aliquots at -20oC (do not filter ppt before aliquoting ; more ppt will form after thawing). To remove precipitate from aliquots, filter with 0.45uM filter before use.
M199 Media Invitrogen# 11150-059 500mL. Store 4oC
Collagen Sigma# C9791-250mg (from calf skin). Store 4oC
Peristalic Pump Rainin Dynamax, model RP-1 (available in the lab)
To perfuse the mouse, portal vein perfusion will be used. Perfusion and Digest media will be pumped into the portal vein (via 27g needle), through the liver, and out the Inferior Vena Cava (by sniping this vein below the kidneys). This circuit allows direct perfusion of the liver.
Plates need to be coated with collagen; collagen coating is necessary for the cells to adhere to the plate. Typically use 6 or 12 well plates. (see collagen coating protocol below).
Practice with several mice before doing the experiment.
Preset the pump to setting 10 (~3mL per minute)
Perfuse the mouse with Liver Perfusion Medium within ~5-10 minutes of sacrificing the mouse to avoid blood clotting.
To prime the pump, fill the reservoir filled with 25mL pre-warmed (37oC) liver Perfusion Media, press the forward button, and then the prime button. Flick the full length of the tubing during priming to dislodge air bubbles. Any air entering the system and going into the mouse will disrupt the perfusion.
While perfusing the mouse, be careful during the transition from Liver Perfusion Media to Liver Digest Media. Do not let the reservoir run dry. Stop the pump before the last few milliliters of Perfusion Media are pumped out, fill reservoir with 30mL Digest Media (avoid creating bubbles) and restart the pump.
Cell Preparation Notes
1-6x10e7 live cells can be obtained from one 6-12 week old mouse (average is ~2x10e7).
Mice younger than 6 weeks are not recommended.
The best cell preps are usually with 7-9 week old mice.
10-12 week old mice can also work well.
Male mice have larger livers and may yield more cells.
Older mice (5-12 months old) might not work as well.
A good preparation contains about 50-80% live cells as determined by trypan blue staining (average is ~45-65%).
Most protocols report 90-95% viability, but Ive never found this to be the case.
Preps with ~30-40% live cells can be plated but with fewer cells adhering. To increase the number of cells on the plate, a second prep can be either combined with these or plated on top of these (after aspirating the non-adherent dead cells).
Preps with 20% or lower live cells usually yield no attached cells and should be discarded.
1-2x10e6 live cells are usually plated in one well of a 6 well collagen coated plate (for best results plate 2x10e6). One well is usually lysed with 150 uL 1x lysis buffer to obtain 100-350 ug protein. This is typically enough for several WB or for IP.
Hepatocyte Isolation Protocol
Media, Collagen Plates, and Pump preparation
Collagen-coat plates before beginning (see protocol below).
Pre-warm (37oC) the Liver Perfusion Media (this is the bottle with the clear colored media)
Thaw and Pre-warm (37oC) the Liver Digest Media (this is the bottle with the red colored media). There is usually a precipitate (ppt) in the Digest Media after thawing. Filter with 0.45uM syringe filter to remove ppt. Two 35 mL aliquots can be combined in a 60mL syringe and filtered at the same time.
Note: Alternatively, the ppt. can be pelleted at 2000 rpm for 5-10 minutes. Carefully remove the media with a 25mL pipet; leave 1-2 mLs media above the pellet as to not disturb the ppt.
(the ppt pelleting procedure saves syringes and filters).
Fill the pump reservoir with ~25mL pre-warmed (37oC) Liver Perfusion Media and prime the pump as described above in general notes.
Attach a new 27g needle to the tubing and press the forward button to prime the needle by flicking the needle (leave the protective cap on until ready to use. Use one needle per mouse to ensure a sharp needle).
Set pump to setting 10 (~3mL per minute)
Mouse Preparation and Perfusion
To anesthesize/sacrifice the mouse, put a Kimwipe into a 50mL tube and add 1-3mLs of isoflurane. Cap the tube when not in use.
Hold the mouse behind the neck and put its nose inside the isoflurane tube until it passes out, and then put the whole mouse inside the tube until it dies. This should only take 1-3 minutes; if the mouse breathes longer than that, add more isoflurane to the tube.
Work quickly from this point to perfuse the mouse with Liver Perfusion Media within 5-10 minutes after sacrificing to avoid blood clotting.
Wet the mouse completely with 70% EtOH.
Tent, snip and pull apart the skin at mid abdomen.
Use pins to hold down all four feet.
Dissect away the skin to expose the whole abdomen and chest.
Open the chest cavity by cutting along both sides of the ribcage (not up the middle along the breastbone). Be careful not to cut any major arteries, especially around the heart. Snip the diaphragm. Pin back the rib cage so it is out of the way (opening the chest cavity makes it easier to remove the liver after perfusion. This is not shown in fig 1).
Push the intestines to the side and the liver up to expose the portal vein (on the underside of the liver.) You will see that the portal vein runs from the liver into the intestines. Note the dark color of the liver before perfusion. See figure 1.
Uncap and briefly re-prime the needle by pressing the forward button and flick the needle to be sure no air is inside. Turn the pump off after priming.
Use needles or tape to secure the tubing so any slack or tension will not pull the tubing and, therefore, the needle, after inserting it into the portal vein.
Carefully insert the 27g needle (bevel up, pump off) into the portal vein going towards the liver.
Turn on the pump (setting 10, ~3mL/min). Be sure and have the pump reservoir filled with ~20-25mL pre-warmed Perfusion Media.
Immediately snip the Inferior Vena Cava (IVC) below the kidneys (the IVC branches to both kidneys, snip below that branch point, about where the IVC arrow is pointing in Figure 1).
The liver will quickly become pale as the blood is replaced with Perfusion Media. The IVC will become clear as blood and perfusion media are pumped out. See Figure 2)
Perfuse all 20-25mLs through the liver. The Perfusion Media flushes out blood, prevents blood clotting, and begins dissociating the hepatocytes.
Stop pump before the last few milliters of Perfusion Media run out. Do not allow bubbles into the tubing.
Fill reservoir with 30mL pre-warmed (37oC) liver Digest Media and restart the pump, setting 10. Put the remaining 5mL Digest Media into a p100 for later use.
The Digest Media will turn the liver slightly more pale. (sometimes you can see this happen)
To ensure the Digest Media is being pumped through the liver, clamp the IVC just above the snip with tweezers to cut off the media draining from the IVC. The liver will begin to swell, release the clamp and the liver will deflate. Repeat this several times throughout the perfusion. Do not let the liver swell maximally. The liver can swell so much it almost doubles in size. If there is too much pressure, the lower left lobes can actually bleep and rupture, short-circuiting flow through the liver. Let the liver swell to only a 30-60% increase in size. Swell and release repeatedly until all the digest media is gone. Note: Blanka does not do the swell and release technique and is still able to digest the liver. I developed this technique after failing for months to successfully digest livers and isolate cells. Blanka also utilizes perfusion through the heart (insert needle into the left atrium and snip the pulmonary vein at top left of the heart. I no longer use heart perfusion.).
Perfuse all 30 mLs through the liver. You may be able see that the liver looks internally digested by morphological changes (fluid collection in the liver and what looks like mushy cell clumps inside the liver). The liver will become very soft as it is digested from inside.
Remove Liver, Dissociate, and Count Hepatocytes
Carefully remove the gal bladder. This can usually be done without spilling the contents.
Carefully remove the liver by cutting the tissue between stomach, gut, and liver. Cut off the liver from the diaphragm. Avoid cutting or tearing open the liver, stomach, or intestines.
Take out the liver from the animal. Place in a p100 dish with 5mL liver digest media and incubate at 37oC for 5-15 minutes. Use Primary Tissue Culture (TC) incubators or heat block in the lab.
If possible, use the Primary TC hoods for the remaining steps. Otherwise work on the bench.
Place plate on ice and add ~20mLs Plating Media (DMEM low glucose, 5%FCS, P/S) to the p100 plate.
Use tweezers to gently peel off the capsule and shake the liver. Gently tease the liver and shake out the hepatocytes. Cells will release from the liver and look like dust in the media. After ~10 minutes of gently shaking and teasing, only a small amount of connective tissue should remain. The media will be cloudy and the large, heavy hepatocytes will quickly settle and look like fine sand in the dish. This procedure may be better visualized by placing the plate on a dark background.
Livers that are not digested well will fall apart in small pieces/clumps and very few individual cells will be released. The liver will be harder and after 10 minutes of teasing there will still be a lot of liver material remaining.
Use a large bore 25mL pipet and slowly (the cells are fragile. Do not use maximum pipet force) pipet the cells through a 70 micron cell strainer (available in Tissue Culture Core) into a 50mL tube on ice.
Count cells. Resuspend the cells by inverting the tube a few times and dilute an aliquot 1:1 with trypan blue (~30uL cells with ~30uL trypan blue in a 1.5 Eppendorph tube) Mix cells and trypan blue by flicking the tube, not by pipeting up and down, and count live cells on a hemacytometer. Note: For best results use the center setting light filter on the TC scopes (adjust the right-left sliding bar on the light source to the center position- the brightest setting). Live cells are bright and dead cells stain blue. The cells are large and many have double nuclei (easily seen in dead, trypan blue positive cells).
Count total live and dead cells. Cell preps with 50% or more live cells are the best. Cell preps with less than 50% live cells do not plate well because the live cells stick to dead cells, preventing the live cells from adhering to the plate.
Spin cells low speed, 50xg, for 2-5 minutes (50xg = ~450-500 rpm in the Sorval swing bucket centrifuge) 4oC. Low speed allows only the large, heavy hepatocytes to pellet and not the other smaller, lighter cells (i.e. red and white blood cells.)
Aspirate supernatant (leave ~1mL so as to not aspirate the cell pellet)
Resuspend/dislodge the pellet by gently flicking the tube
Wash cells by resuspening in 20mL Plating Media. Spin 50xg 2-5min
Repeat wash as above with 20mLs Plating Media.
Aspirate, resuspend/dislodge pellet and resuspend cells in appropriate amount of Plating Media for plating.
Plate cells. For 6 well plate, plate 1-2x10e6 live cells/1.5mls. (Note: use collagen coated plates, see collagen coating protocol below)
After 2-4 hours, the live cells will have attached, although most will not look spread out yet.
Change Media to M199 Media + or Adenovirus CRE
After 2-4 hours the live cells will be stuck to the plate. Some will look like they are beginning to spread out, but most will appear as bright cells that have not yet begun to spread out. There will be many floating dead cells. The adhered cells will look about 40-60% confluent.
Aspirate Plating Media and floating cells.
Wash once with 1mL PBS (1mL/well of 6 well plate). Add the PBS with a 10mL pipet by touching the pipet to the side of the well. Pipet slowly and do not add directly to the center on the plate. This washes away dead cells and Plating Media.
Add 1.5 mL/well of 6 well plate M199 Media + or adenovirus CRE (see formulation below). See adenovirus CRE Treatment and Multiplicity of Infection (MOI) Calculation protocols below.
The next day the cells are spread out and are no longer bright. The cells will appear pentagonal and most will have two nuclei. The bright, non-spread out cells are most likely dead cells stuck to the live cells. Depending on %dead cells, there could be a layer of these covering the live cells. See Figure 3.
Cells are OK for up to 2-5 days (I usually do experiments no later than day 3. The cells change greatly in morphology by day 3-5 and can begin to die. Andy N., in DeBoise Lab, says rat hepatocytes respond to insulin for up to 5 days). Hepatocytes do not proliferate and cannot be passed or frozen.
Adenovirus CRE treatment
Multiplicity of Infection (MOI) 15-20. Higher MOIs can be toxic; MOI 45 has proven to be toxic after 48hrs of treatment. See MOI calculation below.
Adenovirus CRE treat for 48 hours, change media to fresh M199 media (no CRE), and do experiments on the fourth day.
Adenovirus CRE treatment schedule
Day1 Day2 Day3 (48hrs) Day4
Harvest/plate x Change Media experiment
Hepatocytes. Fresh M199 (no CRE)
Add adenovirus CRE
in M199 Media.
DMEM Low Glucose (Invitrogen#11885-084 500mL)
0.4% (w/v) in PBS (200mg/50mL)
Filter with 0.22uM filter to sterilize and remove undissolved dye.
M199 media- with or without Adenovirus CRE (make fresh for each experiment)
M199 media (Invitrogen 11150-067)
100 nM T3 (Sigma T6397) Dissolved in EtOH -20oC
1 nM Insulin (Sigma I5500) Dissolved in H2O -20oC
100 nM dexamethasone (Sigma D4902) Dissolved in M199 or H2O -20oC
With or Without Adenovirus CRE
Reagent stock Final concentration uL for 10mLsT3 1000X (100uM) 100nM 10
Insulin 1000X (1uM) 1nM 10
Dexamethasone 500X (50uM) 100nM 20
Pen/Strep 100X 1X 100
Adenovirus CRE Depends on MOI, pfu/mL, and cell number
T3 (3,3′,5-Triiodo-L-thyronine sodium salt) MW 672.96, powder stored -20oC
Make 10mg/mL solution in 1M Hydrochloric Acid:EtOH (1:4)
This is a 14.8 mM solution (or 148,000X)
Serial dilute to 1000X in 1M Hydrochloric Acid:EtOH (1:4) and use this as a stock solution
Insulin MW 5733.49, powder stored -20oC
Dissolve 2mg/mL acidic H2O (adjust H2O to pH2 with HCl)
This is a 0.348mM solution or 340,000nM (348,000X)
Serial dilute to 1000X in acidic H2O and use this as a stock solution
Store aliquots -20oC. Avoid repeated freeze thaw
Dexamethasone MW 392.46, powder stored 4oC
Dissolve 1mg/mL in MeOH. This is a 2.5mM (25,000X) solution
Dilute 1:50 to 50uM in M199 media. This is a 500X solution. Use this as a stock.
Multiplicity of Infection (MOI) Calculation
1. (Desired MOI) x (Cell#)=pfu needed
2. (pfu needed) / (pfu of adenovirus CRE)=uL of Adenovirus CRE to add to cells
Example MOI 20 for 2.0x10e6 Hepatocytes
1. (20 MOI)x(2.0x10e6 cells)= 4.0x10e7 pfu needed
2. (4.0x10e7 pfu needed) / (2x10e7pfu/uL Adenovirus CRE)=2uL Adenovirus CRE for 2x10e6 Hepatocytes
Collagen coated plates
Modified from Sigma Protocol:
Collagen Type I (Product Nos. C1809, C7661, C9791, and C8919)
Optimal conditions for attachment must be determined for each cell line and application.
1. Add collagen to 0.1 M acetic acid to obtain 0.1% (w/v) (10X) collagen solution. Stir at room temperature 1-3 hours until dissolved. (i.e. add 250mg to 250mL 0.1M acetic acid.)
2. We recommend transferring the collagen solution to a glass bottle with a screw cap and carefully layering chloroform at the bottom. The amount of chloroform to use should be ~10% of the volume of collagen solution. DO NOT SHAKE OR STIR. Allow to stand overnight at 28 °C. Aseptically remove the top layer containing the collagen solution. We do not recommend sterilizing the collagen solution by membrane filtration. We have found substantial protein loss by this method.
3. Dilute desired volume (according to surface area to be treated) of sterile 10X stock solution in step 2 in sterile TC H2O to 0.01% (1X) for coating surfaces.
Dilute 50mL 10X aliquot in 500 total mLs sterile H2O (final concentration 1X). Store unused portion at 4oC.
4. Add 1.5mL to well of 6 well plate. Ensure surface area is completely covered). Allow the protein to bind for several hours (~4hrs) at room temperature or 37 °C, or overnight at 28 °C.
5. Remove excess fluid from the coated surface, and allow it to dry in the TC hood under UV light.
(NOTE: UV light is absolutely necessary since the calf solution cannot be properly sterilized and most likely contains yeast).
~30 minutes UV is enough.
6. Rinse with sterile tissue culture grade water. (NOTE: Do not skip this step. Cells will not attach if any trace acetic acid is left on the plate). Again, allow to dry.
Unused plates can be stored at room temperature.
NOTE may be better not to store 10X or 1X solution at 4oC. Yeast contamination is a problem. Make a small batch of 10X/1X and coat several plates. UV and store the plates at RT.