A Technique for Human Skin Grafting in a Mouse Model

Protokol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Processing of donor human skin sample

NOTE: The human skin sample used in this transplant was a large sample collected from the abdomen of a healthy patient. The sample must be at least 15 cm x 7.5 cm. Size limitations may affect the number of mice for which skin is available and the choice of graft size.

1. Maintain the skin sample at cold temperatures (on ice; 4°C) before preparation and grafting. Keep the sample moist in a closed specimen collection cup with the gauze soaked in phosphate-buffered saline (PBS).
   NOTE: Storing the skin sample at 4 °C for longer than 2 days is not recommended. However, reports exist where the skin samples are stored for a longer time.
   CAUTION: Treat all human tissue with standard biohazard precautions.
2. Prepare to dermatome the human skin sample in a sterilized negative pressure tissue culture hood on a sterilized dissection board.
3. Place the skin sample, epidermis side up, on the dissection board. Wipe the epidermis with a sterile alcohol prep pad and then with PBS.
4. Pin the closer edge of the skin in place with a 1.5-inch dissecting T-pin (see Table of Materials).
5. Dermatome the skin specimen at a 400 µm thickness, applying steady pressure while cutting forward at a 30°-45° angle. Follow all instrument-specific instructions and safety measures (see Table of Materials).
   NOTE: For details on the dermatome technique, please see a previously published report.
6. Prepare a 100 mm x 20 mm Petri dish by placing a sterile gauze soaked in sterile PBS at the bottom of the dish. Place the skin, epidermis side up, onto the wet gauze.
7. Seal and cover the plate edges with a semi-transparent sealing film (see Table of Materials) to ensure the sample is not contaminated. Store the sample at 4° C prior to grafting.

2. Pre-surgery conditioning and preparation

1. Prepare the sterile instruments and a sterile surgical station for grafting. Use autoclaved paper towels as sterile surfaces for instrument and mouse placement.
   NOTE: Mice may be grafted after weaning but are preferably grafted between 8-10 weeks of age. Mice of either sex may be grafted.
2. Perform the surgical preparation, such as hair removal, in an area physically separated from the surgical station.
3. Prepare the anti-GR1 (see Table of Materials) by diluting it to 1 mg/mL in sterile saline. Dose each mouse with 100 µg/100 µL of the anti-GR1 solution intraperitoneally following anesthesia induction.
4. Anesthetize the mice, one at a time, with isoflurane or other institutionally approved anesthetics.
   NOTE: Isoflurane needs to be given at a 3%-5% concentration during induction. Once the mouse is immobile, lower the isoflurane concentration to 1%-3% to affect the duration of the surgery.
   1. Monitor the mouse for appropriate depth of anesthesia by observing respiratory rate, absence of toe-pinch response, and appropriate pink coloration of ears and mouth.
      CAUTION: Use appropriate anesthetic machinery and scavenging methods, and avoid exposure to isoflurane vapors.
5. Transfer the mouse to a heating pad or another heat source (see Table of Materials).
6. Administer the ophthalmic ointment by dabbing a small drop of ointment on the eye with a gloved finger.
7. Administer analgesics Buprenorphine (0.08 mg/kg) and Carprofen (5 mg/kg) (see Table of Materials) subcutaneously by pinching the skin and injecting at an angle parallel to the body.   
   NOTE: Prepare the pre-treatment analgesia following institutional protocols. Follow institutional guidelines for the selection and administration of analgesics. The method of analgesia used in this study is outlined in step 2.7 and Figure 1.
8. Administer the anti-GR1 (prepared in step 2.4) intraperitoneally by slightly lifting the mouse by the tail, exposing the abdomen, and injecting at a 30° angle using an insulin 1 mL (12.7 mm) syringe.
9. Use animal-safe electric clippers (see Table of Materials) to shave the middle and upper portions of the dorsal side of the mouse.
10. Clear all the hair and apply a generous amount of hair removal ointment onto the shaved skin for 30 s to 1 min.
11. Completely wipe away hair removal ointment with a paper towel and PBS.

3. Transplantation procedure

1. Transfer the mouse to a secondary surgical location, away from the hair removal station.
2. Sterilize the surgical site with the iodine swab stick in a circular motion, starting in the middle and working out toward the edge of the depilated area.
3. Place a piece of sterile plastic wrap over the mouse and cut a window in the plastic slightly larger than the size of the area to be grafted.
4. Cut a rectangle-shaped 10 mm x 10 mm portion of donor skin to be grafted with a scalpel. Do this by firmly holding the donor skin in place with the backside of the forceps and cutting alongside the forceps with the scalpel.
5. Using the surgical scissors, snip a rectangular area of mouse skin matching the size of the donor skin piece, creating a graft bed. Use forceps to pull the skin away from the body, and cut the skin with the scissors angled away from the body to avoid cutting deeply into the facia.
6. Place the donor skin piece, epidermis side up, onto the prepared graft bed.
7. Using the back of the forceps, manipulate the skin, sliding back and forth until the donor skin lies completely flat against the graft bed.
8. Add drops of surgical glue tissue adhesive (see Table of Materials) where the donor skin meets the mouse skin and hold the mouse and donor skin together with forceps for 1-2 s so that the glue adheres to the tissues. Completely seal the edge of the graft and allow for the glue to dry fully.
9. Bandage the mice (Figure 2) following the steps below.
   1. Cut a piece of petrolatum gauze (see Table of Materials) large enough to cover the graft area completely.
   2. Cover the graft with the petrolatum gauze and lightly press the gauze against the skin using forceps.
   3. Cut a strip of a transparent film dressing lengthwise so that the width is large enough to cover the mouse's wound.
   4. Firmly press the transparent film dressing, adhesive side down, over the gauze. Quickly roll the mouse to wrap the dressing completely around the torso, ensuring it fits tightly without impeding respiration, and all limbs are free for movement.
   5. Place the mouse in a recovery cage and monitor it until it is alert and moving around. Provide a heat source on part of the cage for at least 15 min following recovery.
      NOTE: The animals are expected to recover within 1-5 min after placing them in the recovery cage.

Sonuçlar

Figure 1: Timeline of the protocol adopted for the xenograft study.

Figure 2: Method for bandaging mouse after surgery. The mouse is wrapped tightly in petrolatum gauze and transparent film dressing while under anesthesia. (A) Petrolatum gauze ...

Malzemeler

Name	Company	Catalog Number	Comments
10% Neutral Buffered Formalin	Fisher	SF100-20	Fixative for histology
3M Vetbond Tissue Adhesive	3M	1469SB	Surgical glue
Anti-GR1 clone RB6-8C5	BioXcell	BE0075	Anti-rejection
Autoclave pouches	VWR	89140-800	For autoclaving tools and paper towels
Buprenex 0.3 mg/mL	Covetrus	59122	Analgesia
Carprofen 50 mg/mL	Zoetis	NADA # 141-199	Analgesia
D42 Dermatome blade	Humeca	5.D42BL10	Dermatome (1 blade per sample)
Dermatome D42	Humeca	4.D42	Dermatome
Disposable Scalpel	Bard-Parker	371610	skin preparation
Dissecting T-Pins; 1-1/2 inch, 1000/CS 1.5	Cole-Parmer	UX-10915-03	To pin skin specimen for dermatome
Dissection scissors	medicon	02.04.10	Sample preparation and mouse dissection
DNAse	Sigma-Aldrich	DN25-1G	Skin digestion
Electric clippers	Kent	CL8787-KIT	Hair removal
Forceps	medicon	07.60.07	Sample preparation and mouse dissection
Gauze	Fisherbrand	22-362-178	Sample preparation
Heating lamp	Morganville Scientific	HL0100	Post-surgical care
Heating pads 4" x 10"	Pristech	20415	Surgical heat supply
Insulin 1cc 12.7 mm syringes	BD	329410	Drug administration
Isoflurane	United States Pharmacopeia (USP)	NDC 66794-013-25	Anesthesia
Isoflurane machine	VetEquip	911103	Anesthesia
Nair for Men	Nair	‎10022600588556	Hair removal
Neomycin and Polymyxin Bisulfates and Bacitracin Zinc Ophthalmic ointment	Dechra	NDC 17478-235-35	Eye ointment to prevent drying
NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ (NSG) mice	The Jackson Laboratory	5557	Mice
Paper towels	Kleenex	100848	May be autoclaved for sterile surfaces
Parafilm	Fisher Scientific	13-374-12	Semitransparent sealing film
Petri Dish 150 mm	Corning	430597	Sample storage
Plastic Wrap	Fisherbrand	22-305-654	Site preparation
Providone-Iodine Swab stick	PDI	S41350	Site sterilization
Soft-Feed and Oral Hydration (Napa Nectar)	Se Lab Group Inc	NC9066511	For supplementing poorly recovering mice post-surgery
Specimen Collection Cups	Fisher Scientific	22-150-266	Sample storage
Sterile alcohol prep pad	Fisherbrand	22-363-750	Skin preparation
Sterile PBS	Gibco	14190-144	Media for sample storage
Sterile saline	Hospira	NDC 0409-4888-02	For drug dilution
Tegaderm Film 4" x 43/4"	3M	1626	Transparent film wound dressing
Vaseline Petrolatum Gauze 3" x 8"	Kendall	414600	Wound dressing