Generation of axolotl hematopoietic chimeras

Background

Materials

Animals

Reagents

• Chlorhexidine
• Sodium chloride (NaCl) (Fisher Scientific, # S640-50)
• Potassium chloride (KCl) (Fisher Scientific, # P217-3)
• Calcium chloride (CaCl₂) (Fisher Scientific, # C79-3kg)
• Magnesium sulfate (MgSO₄) (Fisher Scientific, # M63-3)
• NovAqua Plus water conditioner (Aquatic Eco-Systems, # NA64P)
• AmQuel Plus ammonia detoxifier (Aquatic Eco-Systems, # AM64P)
• L15 Media (Cellgro, # 10-045-CV)
• Distilled water
• FBS – Advantage (Atlanta Biologicals, # S11050)
• Penicillin-streptomycin solution (Invitrogen, # 15140122)
• Insulin-Transferrin-Selenium (Life Technologies, # 51300-044)
• Tricaine (Ethyl 3-aminobenzoate methanesulfonate salt) (SIGMA-ALDRICH, # A5040-250)
• Sodium bicarbonate (SIGMA-ALDRICH, # S5761-1)
• PBS (Fisher Scientific, # MT21040CMRF)
• 70% ethanol (Diluted with dH₂O) (Decon Labs, # 64-17-5)
• Sulfamerazine (SIGMA-ALDRICH, # S8876-250)
• Blue food dye (McCormick, UPC # 050428138861)
• Calcium chloride dihydrate (CaCl·2H₂O) (Fisher Scientific, # C79-500)
• HEPES (Fisher Scientific, BP410-500)
• Sodium hydroxide (NaOH) (Fisher Scientific, # BP359-500)
• G418 (Cellgro, # 61-24-RG)

Equipment

• Gloves (Fisher Scientific, # 191301597D)
• Plastic transfer pipettes (Fisher Scientific, # 13-711-20)
• 35 mm cell culture dishes (Fisher Scientific, # 08-757-11YZ)
• Agarose (SIGMA-ALDRICH, # A9045-10)
• Watch maker forceps (Roboz, RS-5065)
• Blunt tipped Forceps (Roboz, RS-5130)
• Microsurgical scissors (Roboz, # RS-5603)
• Standard surgical scissors (Roboz, # RS-5910)
• Irradiator (137 Cesium source irradiator)
• Microinjector (Leica mechanical micromanipulator)
• Kwik-Fil Borosilicate Glass Capillaries (1 mm outer diameter, no filament; World Precision Instruments, # MTW100F-4)
• Needle puller (Sutter Instrument Co, # P-97)
• Dissecting stereo microscope (Leica, # MZ6)
• Scalpel or razor (Personna, # 94-120-71)
• Insulin syringe (Excel International, # 26027)
• 6-0 nonabsorbable monofilament polypropylene sutures (Ethicon, # 8706)
• 4-0 nonabsorbable monofilament polypropylene sutures (Ethicon, # 8831H)
• Fully frosted microscope slides (Fisher Scientific, # 12-544-5CY)
• 70 µm nylon cell strainer (Fisher Scientific, # 22-363-548)
• Bench top centrifuge (Sorvall, # RT7 Plus)
• 50 ml conical centrifuge tubes (Homecare Supplies, # 352070)
• Non-toxic reusable adhesive Handi-tak (Super Glue Corporation, # HT2)
• Hemocytometer (Reichert Bright-Line, # 1483)
• Vacuum driven filtration system (Fisher Scientific, # SCGPU05RE)
• Parafilm (Fisher Scientific, # 1337412)
• Hot bead sterilizer (Fine science tools, # 18000-45)

Recipes

• 40% Holtfreter’s solution (167 L/44 gal)
  • 5 g/1 teaspoon KCl
  • 12.5 g/2.5 teaspoons CaCl₂
  • 30 g/2 tablespoons MgSO₄.7H₂O
  • 237 g/1 cup NaCl
• Steinberg’s solution (1 L)
  • 3.4 g NaCl
  • 0.05 g KCl
  • 0.03 g CaCl·2H₂O
  • 1.1 g HEPES
  • 1x antibiotic-antimycotic
  • 25 mg G418
  • pH to 7.4 with NaOH
  • Filter sterilize
• APBS
  • Dilute PBS to 70% with dH₂O
• Axolotl fibroblast media
  • 60% L-15 media with distilled water
  • 5% FBS
  • 1x penicillin-streptomycin
  • 1x insulin-transferrin
• Tricaine methanesulfonate solution (0.1%)
  • 1.5 L dH₂O
  • 1.5 g Tricaine methanesulfonate
  • pH to 7.4 with sodium bicarbonate

Procedure

Ablative hematopoietic cell transplantation

1.Irradiation of recipient axolotls should be done 1-2 days prior to HCT:

1.1.Anesthetize white adult recipient axolotl by submersion in 0.1% tricaine solution until unresponsive (10-20 min depending on size).

1.2.Place the axolotl in a container suitable for the irradiator with its tank water (40% Holtfreter’s solution) and protect the animal’s head and gills with lead shielding (Fig. 1A).

1.3.Irradiate axolotl with 950 rads and return the animal to its housing tank for recovery.

2.Harvesting of hematopoietic organs (liver and spleen) with non-survival surgery of donor to maximize number of hematopoietic cells. (Refer to step 3 for survival surgery procedure):

2.1.Sterilize blunt-nosed forceps, surgical scissors, and any other surgical tools in 70% ethanol, hot bead sterilizer, or autoclave.

2.2.Anesthetize GFP+/nucCherryRed+ larval donor axolotl by submerging in 0.1% tricaine solution for 5-10 min.

2.3.Remove axolotl from the tricaine solution and place on dissecting tray or moist paper towel.

2.4.With a scalpel/razor or surgical scissors decapitate the anesthetized animal.

2.5.With surgical scissors cut the ventral skin and muscle tissue starting from the neck opening towards the tail to expose the ventral cavity.

2.6.Remove the entire liver (large light pink-yellow or grey organ at the top of the ventral cavity) and spleen (dark red organ attached to the stomach) by gently lifting organs with blunt-nosed forceps and cut away connective tissue with surgical scissors (Fig. 1B).

2.7.Locate and remove the gall bladder from the liver.

2.8.Place the liver and spleen in ice cold APBS.

3.Harvesting of hematopoietic organs (liver and spleen) with survival surgery of donor for further studies:

3.1.Sterilize watchmaker forceps, microsurgical scissors, and any other surgical tools in 70% ethanol, hot bead sterilizer, or autoclave.

3.2.Anesthetize GFP+/nucCherryRed+ larval donor axolotl by submerging in 0.1% tricaine solution for 3-5 min or about 2 additional min after the animal is completely unresponsive.

3.3.Transfer anesthetized axolotl to 40% Holtfreter’s solution with 0.5% sulfamerazine for 30 s.

3.4.Place animal on dissecting tray or moist paper towel under a dissecting microscope and make sure that the animal’s skin does not dry out throughout the procedure by dripping water on it periodically.

3.5.With a scalpel/razor make two small incisions in the skin directly over the liver and spleen just enough to puncture the skin. This will be visible under the dissecting microscope (Fig. 1C).

3.6.With microsurgical scissors increase the size of the incisions to the minimum required length for access to the liver and spleen.

3.7.Using watchmaker forceps pull the caudal tip of the spleen through the incision in the skin and cut approximately half of the spleen with microsurgical scissors.

3.8.Using watchmaker forceps pull one of the left or right edges of the liver through the incision in the skin and cut off approximately 1/3 of the liver.

3.9.Place the pieces of liver and spleen in ice cold APBS.

3.10.Make sure the liver and spleen of the animal are not jutting out from the skin.

3.11.Place animal back in 40% Holtfreter’s solution with 0.5% sulfamerazine.

4.Grind the liver and spleen separately in to two 35 mm petri dishes containing 1 ml of APBS using two fully frosted glass slides.

5.Keep liver and spleen cell solutions separate and pass them through 70 µm nylon cell strainers directly into two 50 ml conical centrifuge tubes to make single cell solutions.

6.Count cells with a hemocytometer.

7.Spin cells down in a centrifuge at approximately 244g for 5 min.

8.Remove supernatant and resuspend cells at least at 5× 10⁵ cells per 150 µl of APBS for each organ then place on ice.

9.Anesthetize white adult recipient axolotl by submersion in 0.1% tricaine solution until unresponsive (10-20 min depending on size)

10.Dilute blue food dye with APBS and add one drop of dilute solution to liver and spleen cell solutions.

11.Load at least 175 µl of each cell solution into one insulin syringe (minimum total of 350 µl).

12.Locate the hind leg vein on the recipient axolotl and inject cell solution intravenously near the point where the leg joins the body avoiding the injection of any air (Fig. 1).

13.As soon as the syringe is removed, swing the leg back towards the tail to block the injection site with the tail and apply light pressure for 5-10 s (prevents leakage of cell solution from the leg).

14.Return recipient axolotl to housing tank with clean 40% Holtfreter’s solution.

15.Engraftment will be evident by observing donor-derived blood cells in the skin or circulation within 1 month. If an adult donor was used, signs of GVHD will be noticeable in the recipient between 6-8 weeks.

Microinjection hematopoietic cell transplant (non-ablative)

16.Make glass pulled needles using borosilicate glass capillaries and needle puller.

17.Break the tip of the glass pulled needles using watchmaker forceps so that they can be loaded.

18.Prepare a 35 mm petri dish with reusable adhesive and make a depression in which the embryo or larvae will fit as seen in Figure 2.

19.Mix fresh Steinberg’s solution with fresh tricaine solution at a ratio of 3:1 and fill the petri dish with this solution. Cover it when not in use.

20.Prepare glass pulled needles and microinjection apparatus.

21.Harvesting of hematopoietic organs (liver and spleen) with survival surgery of donor for further studies:

21.1.Anesthetize GFP+/nucCherryRed+ adult donor axolotl by submerging in 0.1% tricaine solution for 10-20 min or for an additional 2-5 min after the animal is completely unresponsive.

21.2.Transfer anesthetized axolotl to 40% Holtfreter’s solution with 0.5% sulfamerazine for 30 s.

21.3.Place animal on dissecting tray with a moist paper towel and make sure that the animal’s skin does not dry out throughout the procedure by dripping water on it periodically.

21.4.Apply 0.75% chlorhexidine solution with sterile gauze to the areas of the skin over the spleen and liver.

21.5.With a scalpel/razor make two small incisions in the skin directly over the liver and caudal half of the spleen just enough to puncture the skin and muscle where the internal organs will be visible (Fig. 1B-C).

21.6.Tip: the outline of the spleen will be lightly visible due to its dark red color on the left side of the axolotl’s body. Make the incisions parallel to the direction of the body to allow adjustments in size to be made if necessary.

21.7.With surgical scissors increase the size of the incisions to the minimum required length for access to the liver and spleen (less than 1 inch in length for an adult axolotl).

21.8.Using blunt-nosed forceps pull the caudal tip of the spleen through the incision in the skin and cut approximately 1/4 - 1/2 of the spleen with surgical scissors.

21.9.Place the piece of spleen in ice cold APBS.

21.10.Push the spleen back under the skin and use a simple interrupted suture pattern on the skin with a 4-0 or 6-0 nylon monofilament.

21.11.Using blunt-nosed forceps pull one of the left or right edges of the liver through the incision in the skin and cut off approximately 1/4 - 1/3 of the liver.

21.12.Place the piece of liver in ice cold APBS.

21.13.Push the liver back under the skin and use a simple interrupted suture pattern on the skin with a 4-0 or 6-0 nylon monofilament.

21.14.Place animal back in clean 40% Holtfreter’s solution with 0.5% sulfamerazine. Keep animals in clean water and with the sulfamerazine for about 3 weeks.

22.Grind the liver and spleen separately in to two 35-60 mm petri dishes containing 1-5 ml of APBS using two fully frosted glass slides.

23.Keep liver and spleen cell solutions separate and pass them through 70 µm nylon cell strainers directly into two 50 ml conical centrifuge tubes to make single cell solutions.

24.Spin cells down in a centrifuge at approximately 244 g for 5 min.

25.Remove most of the supernatant and resuspend spleen cells in leftover supernatant.

26.Remove supernatant in liver cells and aspirate off white adispose cells that overlay hematopoietic cells. Then resuspend liver cells in a small amount of APBS to maintain a highly concentrated but fluid cell solution.

27.Cell solutions can be mixed or kept separate but should be maintained on ice. A small amount of food dye can be added to help visualize the infusion of the cell solution into the heart.

28.Place embryo or larva into the 35 mm petri dish with Steinberg’s and tricaine solution.

29.With a Pasteur pipette mix cell solution and dispense one drop onto clean parafilm, petri dish, or plastic lid.

30.Load about 1/3 of the glass microinjection needle with cell solution.

31.Ensure that the embryo or larvae is anesthetized and position its back into the depression made in the adhesive (ventral side up) as seen in Figure 2.

32.Position the glass needle at an approximately 60° angle over the heart and lower it into the water.

33.Under a dissecting microscope begin to expel some of the cells to remove the Steinberg’s and tricaine solution that moved into the glass needle.

34.As cells begin to flow out of the needle, lower it into the larva’s or embryo’s heart and make sure it punctures it but does not go through it.

35.Inject the cell solution slowly until it is clearly all throughout the body or leaking into the ventral cavity.

36.Remove the needle from the heart while holding the animal gently down with forceps.

37.Transfer the animal into freshly made Steinberg’s solution. Keep animals in this solution for 24 h and then transfer them into clean 40% Holtfreter’s solution.

38.If engraftment of long-term HSCs is unsuccessful, donor cells will disappear within 6-10 weeks.

Embryo fusions

39.Prepare 35 mm petri dishes by pouring autoclaved 2% agarose to approximately 1/3 full and allow to solidify.

40.Sterilize watchmaker forceps and microsurgical scissors in 70% ethanol or hot bead sterilizer.

41.Under a dissecting microscope, identify and collect embryos at stages 14–20.

42.Under a dissecting microscope, manually de-jelly the embryos using two watchmaker forceps carefully so as not to damage the embryo.

43.Transfer embryos using appropriately sized Pasteur pipettes into fresh 100% Steinberg’s solution for 5 min as a first wash. Wash two more times in separate containers for at least 5 min each time.

44.Fill agarose lined dishes with 100% Steinberg’s solution and transfer in two same staged embryos of different colors.

45.Under the dissecting microscope, make two small depressions in the agarose using watchmaker forceps that will precisely fit each embryo (Fig. 3).

46.Transversely cut each embryo in half with microsurgical scissors ensuring that the cut is at the same location on each embryo (Fig. 3B-C).

47.Remove clear embryo membrane carefully with forceps.

48.Pair the anterior end of one embryo with the posterior end of the other (Fig. 3B-C).

49.Move paired halves into depressions made in agar with neural folds touching and cover dishes with lids. Intimate opposition of the neural folds is essential for proper healing and further development (Fig. 3B-C).

50.Move dishes very carefully to the side where they need to remain undisturbed for about 96 h. Embryos will fuse entirely within 24-48 h at 20°C (Fig. 3C).

51.Transfer embryos into fresh 100% Steinberg’s solution for another 7 days in covered petri dishes.

52.Finally, transfer embryos to 40% Holtfreter’s solution. Many embryos will have defects due to improper healing and will die.

Anticipated results

Ablative HCT

Microinjection HCT

Embryo fusions

Figure 1. Three main steps in irradiative HCT: irradiation setup, anatomy for cell harvesting, and recipient injection site. A. Lead block keeps the axolotl’s head and gills protected from irradiation that will come from above and below the animal. Any other lead shielding that will protect the head and gills but leave the liver and spleen exposed is acceptable. B. Relevant axolotl anatomy of an adult and location where incisions should be made on the donor animal. Incisions are longer than necessary in this picture and should be made only as long as is necessary to harvest desired tissues if performing the survival surgery. C. A juvenile axolotl with black lines indicating where incisions should be made. In young animals like this one the organs are visible through the skin facilitating this procedure. D. HCT injection site in the recipient’s leg vein. Syringe should follow the path of the arrow and only needs to puncture the vein with the tip.

Figure 2. Microinjection setup. White arrow indicates the heart.

Figure 3. Embryo fusion procedure in agarose lined dishes as seen under a dissecting microscope. A. Dissecting microscope setup. B. Side view diagram of the embryo surgical procedure. C. Microscope top-down view of the embryo procedure and healing process up to 24 hrs following the procedure.

Figure 4. White adult axolotl that underwent ablative HCT from spleen and liver (imaged 4 months after HCT). A. Head. B. Eye. C. Liver. D. Lung. E. Heart. F. Brain. G. Intestine. H. Spleen. I. The liver, gastrointestinal tract, and iris of the eye are highly autofluorescent tissues in the axolotl making it difficult to discern between individual donor and host cells as seen in (B), (C), and (G). A GFP+ liver is placed next to a white liver to show that autofluorescence of non-GFP tissues is usually minimal in comparison to GFP+ tissues when seen from afar. J. Abdominal area of a microinjected animal demonstrating high autofluorescence. The white arrow points to the stomach with autofluorescent food. The difference in color through the GFP channel, GFP and RFP dual channel, and RFP channel alone can be useful to distinguish between true GFP fluorescence (green and will only fluoresce in the GFP channel) and autofluorescence (yellowish but will fluoresce in multiple channels). This same technique is applicable to nucCherryRed+ cells and tissues.

Figure 5. White larval axolotl that underwent non-ablative microinjection HCT.

Troubleshooting

Ablative HCT

Figure 6. Ventral side of a larval axolotl made through the fusion of GFP+ and nucCherryRed+ embryos. A. Green channel. B. Red channel. C. Green and red dual channel.

Figure 7. Another larval axolotl derived through embryonic fusion. A. Tail seen in the green channel. B. Tail seen in the green and red dual channel. C. Hand seen in the green channel. D. Tail fin seen in the green and red dual channel.

Figure 8. In vivo GFP+ immune cell recruitment to the site of injury in the ablative HCT axolotl model. A. Circular cut made with a 2 mm biopsy punch razor on the tail fin. Skin was cut but left in place. B. The same cut at 24 h demonstrating immune cell participation in wound healing.

Microinjection HCT

Embryo fusions

Limitations

Ablative HCT

Microinjection HCT

Embryo fusions

Acknowledgements

References