a technique for human skin grafting in a mouse model

A Technique for Human Skin Grafting in a Mouse Model

After transferring the mouse to the surgical station, sterilize the surgical area and place a sterile plastic wrap over the mouse. Cut a window in the plastic slightly larger than the size of the area to be grafted.
Hold the donor skin firmly in place with the forceps&#39; backside, and cut a rectangle-shaped 10-by-10 millimeter portion of donor skin to be grafted alongside the forceps with a scalpel. Pull the skin away from the body to avoid cutting deeply into the fascia. To create a graft bed, snip a rectangular area of mouse skin, matching the size of the donor skin piece.
Then, place the donor skin piece with epidermis-side up onto the prepared graft bed. Using the back of the forceps, manipulate the skin, sliding back and forth until the donor skin lies completely flat against the graft bed.
Add surgical glue tissue adhesive drops where the donor skin meets the mouse skin and hold the mouse and donor skin together with forceps for one to two seconds so that the glue adheres to the tissues. Completely seal the graft&#39;s edge and allow the glue to dry thoroughly.
Next, cut petrolatum gauze large enough to cover the graft area completely. Cover the graft with the petrolatum gauze and lightly press the gauze against the skin using forceps. Cut a strip of a transparent film dressing lengthwise so that the width is large enough to cover the mouse&#39;s wound.
Firmly press the transparent film dressing with the 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.
After placing the mouse in a recovery cage, monitor it until it is alert and moving around. Provide a heat source on the part of the cage for at least 15 minutes following recovery.

a-technique-for-human-skin-grafting-in-a-mouse-model

Research

JoVE Encyclopedia of Experiments

Immunology

Immunotherapy

Model Development and Disease Study

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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.
<ol>
	<li>Maintain the skin sample at cold temperatures (on ice; 4&#176;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 &#176;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.</li>
	<li>Prepare to dermatome the human skin sample in a sterilized negative pressure tissue culture hood on a sterilized dissection board.</li>
	<li>Place the skin sample, epidermis side up, on the dissection board. Wipe the epidermis with a sterile alcohol prep pad and then with PBS.</li>
	<li>Pin the closer edge of the skin in place with a 1.5-inch dissecting T-pin (see&#160;Table of Materials).</li>
	<li>Dermatome the skin specimen at a 400 &#181;m thickness, applying steady pressure while cutting forward at a 30&#176;-45&#176; angle. Follow all instrument-specific instructions and safety measures (see&#160;Table of Materials). 
	NOTE: For details on the dermatome technique, please see a previously published report.</li>
	<li>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.</li>
	<li>Seal and cover the plate edges with a semi-transparent sealing film (see&#160;Table of Materials) to ensure the sample is not contaminated. Store the sample at 4&#176; C prior to grafting.</li>
</ol>
2. Pre-surgery conditioning and preparation
<ol>
	<li>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.</li>
	<li>Perform the surgical preparation, such as hair removal, in an area physically separated from the surgical station.</li>
	<li>Prepare the anti-GR1 (see&#160;Table of Materials) by diluting it to 1 mg/mL in sterile saline. Dose each mouse with 100 &#181;g/100 &#181;L of the anti-GR1 solution intraperitoneally following anesthesia induction.</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Transfer the mouse to a heating pad or another heat source (see&#160;Table of Materials).</li>
	<li>Administer the ophthalmic ointment by dabbing a small drop of ointment on the eye with a gloved finger.</li>
	<li>Administer analgesics Buprenorphine (0.08 mg/kg) and Carprofen (5 mg/kg) (see&#160;Table of Materials) subcutaneously by pinching the skin and injecting at an angle parallel to the body.&#160;&#160;&#160; 
	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&#160;Figure 1.</li>
	<li>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&#176; angle using an insulin 1 mL (12.7 mm) syringe.</li>
	<li>Use animal-safe electric clippers (see&#160;Table of Materials) to shave the middle and upper portions of the dorsal side of the mouse.</li>
	<li>Clear all the hair and apply a generous amount of hair removal ointment onto the shaved skin for 30 s to 1 min.</li>
	<li>Completely wipe away hair removal ointment with a paper towel and PBS.</li>
</ol>
3. Transplantation procedure
<ol>
	<li>Transfer the mouse to a secondary surgical location, away from the hair removal station.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Using the surgical scissors, snip a rectangular area of mouse skin matching the size of the donor skin piece, creating a graft bed. Use&#160;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.</li>
	<li>Place the donor skin piece, epidermis side up, onto the prepared graft bed.</li>
	<li>Using the back of the forceps, manipulate the skin, sliding back and forth until the donor skin lies completely flat against the graft bed.</li>
	<li>Add drops of surgical glue tissue adhesive (see&#160;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.</li>
	<li>Bandage the mice (Figure 2) following the steps below.
	<ol>
		<li>Cut a piece of petrolatum gauze (see&#160;Table of Materials) large enough to cover the graft area completely.</li>
		<li>Cover the graft with the petrolatum gauze and lightly press the gauze against the skin using forceps.</li>
		<li>Cut a strip of a transparent film dressing lengthwise so that the width is large enough to cover the mouse&#39;s wound.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10% Neutral Buffered Formalin</td>
			<td>Fisher</td>
			<td>SF100-20</td>
			<td>Fixative for histology</td>
		</tr>
		<tr>
			<td>3M Vetbond Tissue Adhesive</td>
			<td>3M</td>
			<td>1469SB</td>
			<td>Surgical glue</td>
		</tr>
		<tr>
			<td>Anti-GR1 clone RB6-8C5</td>
			<td>BioXcell</td>
			<td>BE0075</td>
			<td>Anti-rejection</td>
		</tr>
		<tr>
			<td>Autoclave pouches</td>
			<td>VWR&#160;</td>
			<td>89140-800</td>
			<td>For autoclaving tools and paper towels</td>
		</tr>
		<tr>
			<td>Buprenex 0.3 mg/mL</td>
			<td>Covetrus</td>
			<td>59122</td>
			<td>Analgesia</td>
		</tr>
		<tr>
			<td>Carprofen 50 mg/mL</td>
			<td>Zoetis</td>
			<td>NADA # 141-199</td>
			<td>Analgesia</td>
		</tr>
		<tr>
			<td>D42 Dermatome blade</td>
			<td>Humeca</td>
			<td>5.D42BL10</td>
			<td>Dermatome (1 blade per sample)</td>
		</tr>
		<tr>
			<td>Dermatome D42</td>
			<td>Humeca</td>
			<td>4.D42</td>
			<td>Dermatome</td>
		</tr>
		<tr>
			<td>Disposable Scalpel</td>
			<td>Bard-Parker</td>
			<td>371610</td>
			<td>skin preparation</td>
		</tr>
		<tr>
			<td>Dissecting T-Pins; 1-1/2 inch, 1000/CS 1.5</td>
			<td>Cole-Parmer</td>
			<td>UX-10915-03</td>
			<td>To pin skin specimen for dermatome</td>
		</tr>
		<tr>
			<td>Dissection scissors</td>
			<td>medicon</td>
			<td>02.04.10</td>
			<td>Sample preparation and mouse dissection</td>
		</tr>
		<tr>
			<td>DNAse</td>
			<td>Sigma-Aldrich</td>
			<td>DN25-1G</td>
			<td>Skin digestion</td>
		</tr>
		<tr>
			<td>Electric clippers</td>
			<td>Kent</td>
			<td>CL8787-KIT</td>
			<td>Hair removal</td>
		</tr>
		<tr>
			<td>Forceps&#160;</td>
			<td>medicon</td>
			<td>07.60.07</td>
			<td>Sample preparation and mouse dissection</td>
		</tr>
		<tr>
			<td>Gauze</td>
			<td>Fisherbrand</td>
			<td>22-362-178</td>
			<td>Sample preparation</td>
		</tr>
		<tr>
			<td>Heating lamp</td>
			<td>Morganville Scientific</td>
			<td>HL0100</td>
			<td>Post-surgical care</td>
		</tr>
		<tr>
			<td>Heating pads 4&#34; x 10&#34;</td>
			<td>Pristech</td>
			<td>20415</td>
			<td>Surgical heat supply</td>
		</tr>
		<tr>
			<td>Insulin 1cc 12.7 mm syringes</td>
			<td>BD</td>
			<td>329410</td>
			<td>Drug administration</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>United States Pharmacopeia (USP)&#160;</td>
			<td>NDC 66794-013-25</td>
			<td>Anesthesia&#160;</td>
		</tr>
		<tr>
			<td>Isoflurane machine</td>
			<td>VetEquip</td>
			<td>911103</td>
			<td>Anesthesia</td>
		</tr>
		<tr>
			<td>Nair for Men</td>
			<td>Nair</td>
			<td>&#8206;10022600588556</td>
			<td>Hair removal</td>
		</tr>
		<tr>
			<td>Neomycin and Polymyxin Bisulfates and Bacitracin Zinc Ophthalmic ointment</td>
			<td>Dechra</td>
			<td>&#160;NDC 17478-235-35</td>
			<td>Eye ointment to prevent drying</td>
		</tr>
		<tr>
			<td>NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ (NSG) mice</td>
			<td>The Jackson Laboratory</td>
			<td>5557</td>
			<td>Mice</td>
		</tr>
		<tr>
			<td>Paper towels</td>
			<td>Kleenex</td>
			<td>100848</td>
			<td>May be autoclaved for sterile surfaces</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Fisher Scientific</td>
			<td>13-374-12</td>
			<td>Semitransparent sealing film</td>
		</tr>
		<tr>
			<td>Petri Dish 150 mm</td>
			<td>Corning</td>
			<td>430597</td>
			<td>Sample storage</td>
		</tr>
		<tr>
			<td>Plastic Wrap</td>
			<td>Fisherbrand</td>
			<td>22-305-654</td>
			<td>Site preparation</td>
		</tr>
		<tr>
			<td>Providone-Iodine Swab stick</td>
			<td>PDI</td>
			<td>S41350</td>
			<td>Site sterilization</td>
		</tr>
		<tr>
			<td>Soft-Feed and Oral Hydration (Napa Nectar)</td>
			<td>Se Lab Group Inc</td>
			<td>NC9066511&#160;</td>
			<td>For supplementing poorly recovering mice post-surgery</td>
		</tr>
		<tr>
			<td>Specimen Collection Cups</td>
			<td>Fisher Scientific</td>
			<td>22-150-266</td>
			<td>Sample storage</td>
		</tr>
		<tr>
			<td>Sterile alcohol prep pad</td>
			<td>Fisherbrand</td>
			<td>22-363-750</td>
			<td>Skin preparation</td>
		</tr>
		<tr>
			<td>Sterile PBS</td>
			<td>Gibco</td>
			<td>14190-144</td>
			<td>Media for sample storage</td>
		</tr>
		<tr>
			<td>Sterile saline</td>
			<td>Hospira</td>
			<td>NDC 0409-4888-02</td>
			<td>For drug dilution</td>
		</tr>
		<tr>
			<td>Tegaderm Film 4&#34; x 43/4&#34;&#160;</td>
			<td>3M</td>
			<td>1626</td>
			<td>Transparent film wound dressing</td>
		</tr>
		<tr>
			<td>Vaseline Petrolatum Gauze 3&#34; x 8&#34;&#160;</td>
			<td>Kendall</td>
			<td>414600</td>
			<td>Wound dressing</td>
		</tr>
	</tbody>
</table>

Source: Moss, M. I. et al., <a href="https://app.jove.com/v/64040">Xenograft Skin Model to Manipulate Human Immune Responses&#160;In Vivo.</a>&#160;J. Vis. Exp. (2022)
This video demonstrates a method for grafting human skin onto an immunodeficient mouse with a suppressed neutrophil response. The method involves applying human split-thickness skin to the mouse&#39;s graft bed, sealing it, and covering it for recovery. Eventually, the human skin adheres to the mouse&#39;s skin, establishing vascularization and circulation, resulting in a successful skin xenograft.

<img alt="Figure 1" src="/files/ftp_upload/22230/22230_Figure1.jpg" /> 
Figure 1: Timeline of the protocol adopted for the xenograft study.
<img alt="Figure 2" src="/files/ftp_upload/22230/22230_Figure2.jpg" /> 
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 ...

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Begin with an anesthetized, non-obese diabetic, and immunodeficient mouse that lacks various immune cells.
The mouse is pre-treated with antibodies to block the neutrophil surface receptors and suppress the immune response.
Excise the skin from the mouse&#39;s back without damaging the connective tissue and create a graft bed.
Place a human split-thickness skin, comprising the epidermis and a portion of the dermis, on the graft bed, with the epidermis facing up.
Secure it with glue and bandage it to minimize infections and restrain the graft tissue.
During recovery, the graft tissue receives oxygen and nutrients from the mouse skin, facilitating its attachment.
Over time, blood vessels develop between the human skin and the mouse skin.
This helps to establish circulation and infiltration of human immune cells, achieving a successful human skin grafting in the mouse model.

59 Aufrufe. Eine Technik zur Transplantation menschlicher Haut in einem Mausmodell

Serie: Eine Technik zur Transplantation menschlicher Haut in einem Mausmodell - Experiment

A Melanoma Patient-Derived Xenograft Model

Accumulating evidence suggests that molecular and biological properties differ in melanoma cells grown in traditional two-dimensional tissue culture vessels versus in vivo in human patients. This is due to the bottleneck selection of clonal populations of melanoma cells that can robustly grow in vitro in the absence of physiological conditions. Further, responses to therapy in two-dimensional tissue cultures overall do not faithfully reflect responses to therapy in melanoma patients, with the majority of clinical trials failing to show the efficacy of therapeutic combinations shown to be effective in vitro. Although xenografting of melanoma cells into mice provides the physiological in vivo context absent from two-dimensional tissue culture assays, the melanoma cells used for engraftment have already undergone bottleneck selection for cells that could grow under two-dimensional conditions when the cell line was established. The irreversible alterations that occur as a consequence of the bottleneck include changes in growth and invasion properties, as well as the loss of specific subpopulations. Therefore, models that better recapitulate the human condition in vivo may better predict therapeutic strategies that effectively increase the overall survival of patients with metastatic melanoma. The patient-derived xenograft (PDX) technique involves the direct implantation of tumor cells from the human patient to a mouse recipient. In this manner, tumor cells are consistently grown under physiological stresses in vivo and never undergo the two-dimensional bottleneck, which preserves the molecular and biological properties present when the tumor was in the human patient. Notable, PDX models derived from organ sites of metastases (i.e., brain) display similar metastatic capacity, while PDX models derived from therapy naive patients and patients with acquired resistance to therapy (i.e., BRAF/MEK inhibitor therapy) display similar sensitivity to therapy.

Patient-derived xenograft (PDX) models more robustly recapitulate melanoma molecular and biological features and are more predictive of therapy response compared to traditional plastic tissue culture-based assays. Here we describe our standard operating protocol for the establishment of new PDX models and the characterization/experimentation of existing PDX models.

Ein melanompatientenabgeleitetes Xenograft-Modell

Accumulating evidence suggests that molecular and biological properties differ in melanoma cells grown in traditional two-dimensional tissue culture ...

Forschung

JoVE Journal

Krebsforschung

Performing Human Skeletal Muscle Xenografts in Immunodeficient Mice

Treatment effects observed in animal studies often fail to be recapitulated in clinical trials. While this problem is multifaceted, one reason for this failure is the use of inadequate laboratory models. It is challenging to model complex human diseases in traditional laboratory organisms, but this issue can be circumvented through the study of human xenografts. The surgical method we describe here allows for the creation of human skeletal muscle xenografts, which can be used to model muscle disease and to carry out preclinical therapeutic testing. Under an Institutional Review Board (IRB)-approved protocol, skeletal muscle specimens are acquired from patients and then transplanted into NOD-Rag1null IL2r&gamma;null (NRG) host mice. These mice are ideal hosts for transplantation studies due to their inability to make mature lymphocytes and are thus unable to develop cell-mediated and humoral adaptive immune responses. Host mice are anaesthetized with isoflurane, and the mouse tibialis anterior and extensor digitorum longus muscles are removed. A piece of human muscle is then placed in the empty tibial compartment and sutured to the proximal and distal tendons of the peroneus longus muscle. The xenografted muscle is spontaneously vascularized and innervated by the mouse host, resulting in robustly regenerated human muscle that can serve as a model for preclinical studies.

Complex human diseases can be challenging to model in traditional laboratory model systems. Here, we describe a surgical approach to model human muscle disease through the transplantation of human skeletal muscle biopsies into immunodeficient mice.

Durchführung von Human Skeletal Muscle Xenografts bei immundefizienten Mäusen

Treatment effects observed in animal studies often fail to be recapitulated in clinical trials. While this problem is multifaceted, one reason for this ...

Entwicklungsbiologie

A Murine Model of a Burn Wound Reconstructed with an Allogeneic Skin Graft

Trivial superficial wounds heal without complications by primary intention. Deep wounds, such as full thickness burns, heal by secondary intention and require surgical debridement and skin grafting. Successful integration of the donor graft into a recipient wound bed depends on timely recruitment of immune cells, robust angiogenic response and new extracellular matrix formation. The development of novel therapeutic agents, which target some key processes involved in wound healing, are hindered by the lack of reliable preclinical models with optimized objective assessment of wound closure. Here, we describe an inexpensive and reproducible model of experimental full thickness burn wound reconstructed with an allogeneic skin graft. The wound is induced on the dorsum surface of anaesthetized inbred wild type mice from the BALB/C and SKH1-Hrhr backgrounds. The burn is produced using a brass template measuring 10 mm in diameter, which is preheated to 80 &#176;C and delivered at a constant pressure for 20 s. Burn eschar is excised 24 hours after the injury and replaced with a full thickness graft harvested from the tail of a genetically similar donor mouse. No specialized equipment is required for the procedure and surgical techniques are straightforward to follow. The method may be effortlessly implemented and reproduced in most research settings. Certain limitations are associated with the model. Due to technical difficulties, the harvest of thinner split thickness skin grafts is not possible. The surgical method we describe here allows for the reconstruction of burn wounds using full thickness skin grafts. It may be used to carry out preclinical therapeutic testing.

The aim of this study was to develop a murine model of burn wound healing. A thermal burn was induced on the dorsal skin of mice using a preheated brass template. Burned tissue was debrided and overlaid with a skin graft harvested from the tail of a genetically similar donor mouse.

Ein murines Modell einer Verbranntenwunde, rekonstruiert mit einem allogenen Hautgraft

Trivial superficial wounds heal without complications by primary intention. Deep wounds, such as full thickness burns, heal by secondary intention and ...

A Technique for Human Skin Grafting in a Mouse Model

Transkript

A Technique for Human Skin Grafting in a Mouse Model