Portal Vein Injection: A Method to Study Cancer Metastasis to the Liver

1. Preparation of the Surgical Area and Instruments

1. Prepare the scissors, forceps, and hemostat by autoclaving at 124 °C for 30 min, 1 – 2 days prior to the planned surgeries. Ensure access to autoclaved or sterile bedding, cages, and food for post-surgical recovery.
2. Prepare an aseptic surgical area, preferably in a laminar flow hood.
   1. Wipe down all surfaces of the surgical area with 10% bleach, including the heating pad, light source, anesthesia tubing and nose cone, and any other part of the surgical suite that will be in close proximity to the surgical procedure while it is being performed.
   2. In the aseptic surgical area, place the cleaned heating pad with sterile drape, light source, anesthesia tubing and nosecone, insulin syringes, 1 ml syringes, bupivacaine, artificial tears, sterile saline, 2 x 2" sterile gauze sponges, 4 x 4" sterile gauze, hemostatic gauze cut into 0.5 – 1 cm2 pieces, scissors, forceps, hemostat, 4-0 vicryl sutures with taper needle, and 50 ml 2% Chlorhexidine gluconate in an autoclaved container.
   3. Ensure that there is room in this space for prepared tumor cells stored on ice.
   4. On the bench adjacent to the surgical area, prepare the recovery area with a second heating pad and clean cages with sterile bedding.
      NOTE: This area can also house items such as a bead sterilizer.

2. Portal Vein Injection

1. One-hour prior to the planned injections, treat Balb/c female mice aged 8 – 15 weeks with 100 µl of 0.015 mg/ml buprenorphine, subcutaneously, for pain management.
   NOTE: This injection protocol may be applied to any strain of female or male mouse at any age, using the appropriate cell lines for changes to the strain.
2. Prepare the tumor cells for injection based on protocols for the cell line or tumor explant of choice. Test all tumor cell lines prior to administration for the presence of murine pathogens to reduce the risk of introducing such pathogens into the animal colony.
   1. For syngeneic Balb/c tumor cell lines including D2A1, D2.OR, and 4T1 tumor cells, thaw cells into a 10 cm tissue culture plate 3 days prior to injection such that the following day cells are at ~ 90 – 100% confluency.
   2. 1 day following tumor cell thaw wash cells once with 1x phosphate buffered saline (PBS) and trypsinize the confluent tumor cells using 2 ml of 0.05% trypsin at 37 °C for 5 min. Add 8 ml of complete media (DMEM high glucose, 10% fetal bovine serum, 2 mM L-glutamine, and 1x penicillin/streptomycin) and passage 1:10 into a fresh 10 cm dish with 10 ml of complete media.
   3. On the day of the injections, wash cells once with 1x PBS and trypsinize as described above.
   4. Resuspend trypsinized cells in 8 ml of complete media, spin for 5 min at 1,500 x g, remove the media and resuspend in 5 ml 1x PBS.
   5. Count cells on a hemocytometer using trypan blue exclusion for viability assessment. Resuspend cells for injection in 1x PBS at a pre-determined concentration and volume.
      NOTE: 5 – 10 μl is recommended as smaller injection volumes prevent unnecessary damage to the liver.
   6. Keep cells on ice for the duration of the injections. Following completion of injections, return a sample of cells to the laboratory and place in culture in complete media for 1 day to ensure viability.
3. Place the mouse under anesthesia with 2 – 2.5% isoflurane (2-chloro-2-(difluoromethoxy)-1,1,1-trifluoro-ethane) delivered in oxygen. Maintain body temperature using the heating pad.  Ensure complete anesthetization by assessing for a reaction to a toe pinch, and then maintain anesthesia at 2 – 2.5% isoflurane.
   NOTE: It is important to monitor the animals breathing rate and adjust the isoflurane flow-rate accordingly throughout the procedure.
4. Place a small amount of artificial tears or vet ointment over each eye to avoid excessive drying of the eyes during the surgical procedure.
5. Place the mouse in a supine position, on its back with abdomen exposed.
6. Remove hair on the ventral left side of the rodent from the second rib space down to the 4th inguinal mammary gland nipple by wiping the area with chemical depilatory. Allow the depilatory to sit for 1 – 2 min and then remove completely with gauze and H2O. This step can be done 1 – 2 days in advance to save time if numerous surgeries are planned.
7. Take one 2 x 2" sterile gauze sponge (soaked in 2% Chlorhexidine) and wipe down the mouse at the site of hair removal. Sterilize the entire surrounding area, including the tail, to minimize bacterial contamination of instruments.
8. Wipe the site of hair removal and surrounding area down with an alcohol prep pad.
9. Repeat 2% Chlorhexidine and alcohol steps once more and finish with a final Chlorhexidine wipe down for a total of three 2% Chlorhexidine and two alcohol prep pad washes. Do the final Chlorhexidine wipe such that the chemical is not dripping around the surgical site to avoid getting Chlorhexidine on internal organs.
   NOTE: Application of large amounts of chlorhexidine and alcohol to the skin and surrounding fur may result in a significant drop in body temperature. Do not wipe with excess volume during steps 2.7-2.9. Maintain body temperature with a heating pad.
10. Using sterile gloves and a sterilized scalpel with sterile blade, make a single 1-inch incision into the skin between the median and sagittal planes on the left side of the mouse, starting just below the ribs and ending just above the plane of the fourth inguinal mammary gland teat.
11. Using autoclaved or bead sterilized scissors and forceps, make a similar 1-inch incision into the peritoneum. Avoid cutting into the mammary fat pad and ensure not to cut the intestines, liver, or diaphragm.
12. Place a 4 x 4" gauze pad soaked in sterile saline on the left side of the mouse, where the incision was made, such that internal organs can be placed on the gauze and not come into contact with the surrounding skin or surgical area.
13. Prepare tumor cells by pipetting up and down several times as tumor cells will settle during preparation of the mouse. Prepare a 25 μl removable needle syringe and 32-gauge needle with tumor cells. Push on the syringe until tumor cells are at the tip of the needle and the plunger is at the appropriate volume for injection; avoid injection of air bubbles.
14. Wipe the outside of the needle with a sterile alcohol pad to remove any external tumor cells. Use caution to avoid needle sticks.
15. Hold the median side of the incision, including skin and peritoneal lining, aside with the forceps and use a sterile cotton swab to carefully pull the large and small intestines out, placing them on the sterile gauze soaked in sterile saline. Pull out large and small intestines until the portal vein is visualized.
16. Cover the internal organs in the saline soaked gauze to maintain internal moisture and sterility.
17. Have an assistant, also wearing sterile gloves, hold the intestines wrapped in the saline soaked gauze gently out of the way with a sterile cotton tipped swab to fully reveal the portal vein. Additionally, it may be necessary to use the autoclaved hemostat or forceps to hold tissue aside on the median side of the incision.
18. Insert the needle loaded with tumor cells ~ 3 – 5 mm into the portal vein ~ 10 mm below the liver at an angle < 5° to the vein, with bevel facing up. Slowly inject the full volume containing tumor cells. Allow blood to flow past the needle head for several seconds to avoid back flow of tumor cells out of the vein. Minimize moving the needle in the vein during the injection. Again, use caution to avoid needle sticks.
    NOTE: Visualization of the portal vein is done without magnification, however a stereo microscope may be used if preferred.
19. Remove the needle while simultaneously placing a sterile cotton tip applicator on the vein with pressure. With the assistant still holding the intestines aside place one piece of 0.5 – 1 cm2 hemostatic gauze over the injection site on the vein.
    NOTE: Hemostatic powder was also attempted for this step in the protocol but was not effective in stopping venous blood loss following injection.
20. Hold the hemostatic gauze at the injection site with pressure from a sterile cotton tip applicator for 5 min.
21. Assess closure of the vein by carefully lifting the hemostatic gauze, if the gauze sticks to the surrounding tissue, a small amount of sterile saline can be used to soak and lift the gauze.
22. If blood loss occurs at this time, place an additional piece of hemostatic gauze at the site with pressure for an additional 5 min. When blood flow has ceased completely, remove the gauze from the mouse.
    NOTE: Blood loss during the surgical procedure must be carefully assessed and if the total allowed blood loss volume is met or exceeded (based on regulatory standard operating procedures for the investigator's institutional review boards) the mouse must be euthanized while under anesthesia by cardiac perfusion.
23. Once the injection site is deemed intact, with no blood leaving the injection site, place the internal organs gently back into the abdominal cavity.
24. Suture the peritoneal lining and then the skin with sterile 4-0 vicryl suture and taper needle using a simple continuous or interrupted suture pattern. Typically, closing the incision requires 10-15 sutures.
25. Inject 100 µl of bupivacaine (5 mg/ml) along the incision site for local pain management using an insulin syringe. Inject 0.5 ml of sterile saline subcutaneously using a 1 ml syringe with 26-gauge needle for hydration. Surgeries take 15 – 25 min to complete.
26. To maintain sterile conditions throughout the surgery, ensure that all tools and materials coming into contact with the mouse, including gloved hands, are cleaned appropriately prior to contact. Where possible use sterile materials and gloves, or minimally utilize a 70% ethanol solution or 10% bleach solution to clean.
27. If multiple surgeries are planned for a single session remake the initial surgical area with fresh sterile drape, insulin syringes, 1 ml syringes, sterile saline, 2 x 2" sterile gauze sponges, 4 x 4" sterile gauze, hemostatic gauze cut into 0.5 – 1 cm2 pieces, 4-0 vicryl sutures with taper needle, and 2% Chlorhexidine. Bead-sterilize the scissors, forceps, and hemostat in between surgeries and allow to adequately cool prior to re-use.