Xenografts

Overview

This protocol covers various forms of xenografts.

Protocol

Materials

  • Scissors
  • Forceps (1 curved, 1 straight)
  • Tape
  • Surgical staples, stapler, and staple remover (3M Vetbond)
  • Heating pad
  • Absorbant diapers
  • Cotton gauze bandages
  • petri dishes (10 cm)
  • PBS
  • Chlorohexadine
  • Mouse hair clipper
  • Eye drops (Refresh Liquigel 0.5 fl oz, sterile)
  • Ketoprofen (0.83 µg/µl)
  • Insulin syringes (30cc)
  • Scale for weighing mice
  • Isoflurane and isoflurane system (RAR provides, send email to Linda Key (keyx001@umn.edu) to reserve machine)
  • Lab book and mouse book, pen and marker
  • Surgery Record sheet
  • Ice bucket with ice
  • Styrofoam container with dry ice

Setup

Reserve the procedure room 1-134A and reserve the isoflurane system (email to Linda Key, keyx001@umn.edu).

Gather all materials on a cart and transport to mouse procedure room.

Turn on BSL2 hood, plug in and turn on heating pad in hood, cover heating pad with diaper.

Set up isoflurane system (see isoflurane setup and operation below).

Load stapler with staples.

Load syringe with ketoprofen

Record all animal numbers, dates, times, and cages in lab book and surgery procedure sheet.

Preparation of tumor chunks for subcutaneous (SC) xenografts

Harvest tumor from mouse or collect tumor from operating room and transport to the lab, generally in a 50 ml conical filled with DMEM or RPMI on ice. Try to minimize the time from tumor collection to implantation. We have found that once the blood supply to the tumor is severed, the tumor tissue will rapidly start to die. Ideally, we try to implant the tumor within 60 minutes of harvesting from the donor.

Work in a BSL2 level fume hood using sterile techniques

Transfer tumor chunks to a petri dish containing ice cold PBS. Separate non-tumor tissue and necrotic-looking tumor tissue from good quality tumor tissue using scissors or scalpel and forceps.

Transfer quality tumor tissue to fresh petri dish containing ice cold PBS. If blood or other contaminants are present, rinse with ice cold PBS.

Cut tumor into small pieces (2mm^3) for implantation and for saving tissue for DNA, RNA, protein, and formalin fixation. Place tissue that will not be used for xenografts into appropriately labelled tubes and store at appropriate temperatures.

Transport xenograft tissue chunks to mouse procedure room

Isoflurane system setup

Plug the air intake filter into the "inspiration" nozzle and place the air intake filter on top of the machine. Connect exit filter to "expiration" nozzle. RAR staff monitors the air intake filter and will replace when necessary.

Plug the tube from the oxygen tank into the open/shut nozzle. If oxygen tank is new, rip off the "new" portion of the card hanging from the oxygen tank, leaving the "in service" portion of the card. If you empty the oxygen tank, rip off the "in service" portion and notify RAR staff that the tank is empty.

Remove the plug from the exit hole and plug in the tube that will connect to the nose cone and the charging tank. Connect the splitter that allows you to connect to the charging tank and the nose cone. Connect other end to the charging tank in the hood, along with the nose cone.

Cut off the pinky finger of a latex glove (approximately 3 cm). Remove the O-ring on the nose cone and place the portion of latex glove over the nose cone. Make two small snips in the tip of the latex glove and then replace the O-Ring. This creates a more snug fit for the mouse's head to ensure good flow of oxygen/isoflurane.

Tape the tubing down in the hood.

Open the oxygen tank by turning the black handle 1/4 turn and twist the Open knob.

Twist the pressure regulator knob and allow oxygen to flow at maximum rate (4). Turn on the isoflurane by pressing the white lock button and twisting the top of the tank to 5%. Charge the tank for approximately 60 seconds while holding your hand over the nose cone. Reduce isoflurane content by turning tank knob to 3.5%. Turn down the flow rate to (3) using the pressure regulator knob.

Surgery

Place 5 mice in isoflurane charging tank and wait until they are anesthetized. Check depth of anesthesia by pinching toe (should not see any reflex) and by monitoring respiratory rate (ideal is 50-60 beats/minute). Monitor mice every five minute throughout the procedure. Adjust isoflurane flow rate if more or less anesthesia is required

Remove mouse from charging tank, gripping from the neck, and firmly place mouses face into the nose cone. Wipe the back of the mouse with a cotton gauze pad saturated with chlorohexidine. Clip ear of mouse using our numbering system.

Two tumor chunks will be implanted along the dorsal midline, with one chunk being placed close to the tail and the other chunk being placed closer to the ears.

Gently grasp the skin with the forceps and make a single snip along the midline with the scissors. The length of the snip should be approximately 2.5 mm, which is the width of the staple that will be used to close the incision.

After creating the incision, gently insert the scissors laterally into the incision approximately 7 cm and gently open the scissors to create a "pocket" for the xenograft. Using forceps, place one of the tumor chunks into the pocket, and gently push it so that it is as far from the incision as possible. With two tweezers pick up the skin of the mouse on either side of the incision to create a "tent". Make sure the tumor chunk is not within the "tent", but has been pushed away into the pocket. Use the stapler to close the incision.

Administer an intraperitoneal (IP) injection of ketoprofen. Remove mouse from nose cone and place eye drops on each eye. Allow mouse to recover on heating pad and then return to cage when mouse is moving. Fill out RAR procedure card and place on cage. Return cage to mouse room.

Disassemble isoflurane machine, turn off oxygen, turn off regulators, press the flush oxygen button. Return to RAR location.

Post-operative care

Monitor mouse daily for signs of pain and distress. For the next two days after surgery administer an IP injection of ketoprofen and record this on the cage card. If signs of pain and distress are evident after three days, consult with RAR staff for further options.

Fill out surgery records and entries in the lab book.

Collection of tumor tissue

When tumor meets the endpoint criteria or when the experimental endpoint has been reached, euthanize the mouse following the standard procedures below:

  • Fill out a necropsy sheet (be sure to put your name, date, mouse number, and reason for the necropsy). Follow any special instructions on the necropsy sheet.
  • Prepare storage tubes and liquid nitrogen and any storage media
  • Weigh the mouse and measure the tumor size before euthanizing
  • Euthanize the mouse following our euthanization SOP
  • Take photos of any tumors or unusual organs. Be sure to include a small piece of paper next to the sample with the mouse's name.

Collect the following samples:

  • Samples for fixing: place in 10% buffered formalin overnight shaking at 4ºC in a 15 ml conical. Make sure the volume of 10% buffered formalin is at least 10x the volume of the samples being fixed. Pour off the formalin and rinse 1x in PBS, then re-fill 15 ml conical with 70% ethanol. Can store this indefinitely.
  • Samples for DNA: Cut small chunks (5 mm x 5mm), at least three, and place in 1.5 ml cryovial and snap freeze.
  • Samples for Protein: Cut small chunks (5 mm x 5mm), at least three, and place in 1.5 ml cryovial and snap freeze.
  • Samples for RNA: Cut small chunks (5 mm x 5mm), at least three, and place in 1.5 ml cryovial. Add 1.5 ml RNALater and let sit overnight at room temperature. The next day, remove the RNALater and snap freeze the sample.
  • Samples for viable transplantation: Cut small chunks (2 mm x 2 mm) and place in freezing media (generally 95% FBS and 5% DMSO). Generally use two cryovials with enough pieces in each cryovial for 5 mice (12 pieces). Place in a freezing container and place in the -80ºC freezer overnight. The next day, transfer vials to a liquid nitrogen tank.

Collect any other samples required by the experimental protocol.

For ovarian subcutaneous xenografts see Alkema, et al., Sci Reports 2015 for a comparison of viable freezing methods