This work was supported by the Office of the Assistant Secretary of Defense for Health Affairs through the&#160;Congressionally Directed Medical Research Program under Award No. W81XWH-13-2-0057. Opinions, interpretations, conclusions, and recommendations are those of the authors and are not necessarily endorsed by the Department of Defense.

All animal surgeries were performed in accordance with protocols approved by the University of Louisville&#39;s Institutional Animal Care and Use Committee (IACUC-approved protocol 18198) and the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals10. Four-month-old male Brown-Norway (RT1.An) and 4-month-old male Lewis (RT1.Al) rats were used as VCA donor and recipients, respectively.
1. Donor Allograft Harvest
<ol>
	<li>Sedate the donor animal using vaporized isoflurane applied through a chamber.</li>
	<li>Shave the graft donor area (hindlimb), as well as the groin and abdomen areas. Following that, treat with depilatory cream in order to reduce the amount of fuzz left by the clippers.</li>
	<li>Deeply anesthetize donor animals using intraperitoneal (IP) ketamine (60 mg/kg)/xylazine (15 mg/kg)/acepromazine (2 mg/kg). Administer an initial dose of 0.2 mL/100 g of body weight and additional doses of 0.2 mL every h.&#160;For convenience, it is optional to perform this step before step 2.</li>
	<li>Continuously monitor animals while under anesthesia for respiration, body temperature, and depth of anesthesia, using the toe pinch withdrawal reflex test.</li>
	<li>Administer 30 U of heparin solution subcutaneously (SC) in the scruff area prior to the surgery to prevent clotting.</li>
	<li>Wear a mask, a head cover, a disposable isolation gown, and disposable gloves.</li>
	<li>Place the donor animal supine on a heating pad. Produce a sterile surgical field by prepping, scrubbing, and draping the surgical area&#160;including the both ventral and dorsal aspects of the leg. Don sterile gloves.</li>
	<li>Make a 3 cm skin incision in the groin concavity using scalpel blade #15 and reflect&#160;the inguinal fat pad laterally using iris scissors.</li>
	<li>Expose the common femoral vessels and place a wire hook with an elastic band to retract the abdominal muscles.</li>
	<li>Using a dissecting microscope (40x), dissect the pedicle proximally from the emergence of the common femoral vessels under the inguinal ligament and distally to the confluence of popliteal vessels into the graft.</li>
	<li>Using microclips and bipolar jewelers&#8217; forceps, ligate and divide the large arterial and venous branches, such as lateral circumflex femoral vessels, superficial caudal epigastric vessels, the saphenous artery, and proximal caudal femoral vessels, to mobilize the main femoral vessels. Cauterize any small branches using fine bipolar forceps.</li>
	<li>Make a skin incision from the center of the previous skin cut along the ventral side of the hindlimb, to the ankle area, using iris scissors.</li>
	<li>Cut the gracilis muscle, as well as the other adductor muscles underneath it, in a vertical fashion to expose and ligate the medial proximal genicular vessels, deep-branching small vessels, and the sciatic nerve. 
	NOTE: At this point, on a separate surgical table, the other surgeon should intubate and anesthetize (2.5%&#8211;3% isoflurane) the recipient animal; this allows the surgeons to prepare the recipient surgical site in time for graft placement and minimize the graft&#39;s ischemic time.</li>
	<li>On the donor animal, make circumferential skin incisions at the level of the knee and ankle. Disarticulate the knee and ankle, remove extraneous muscle and tissue, and make a vertical skin incision on the dorsal side of the hindlimb to free the graft. At this point, the graft (composed of fibula and tibia, covered with related muscles and skin island nourished by its perforators) is connected only by the pedicle.</li>
	<li>Place small clamps as proximally as possible on the femoral artery and vein, and cut the pedicle as proximally as possible, near the inguinal ligament.</li>
	<li>To flush the graft of blood, inject heparinized saline (30 U/mL) into the femoral artery using a 27 G flushing blunt cannula. 
	NOTE: Dilating the artery prior to the heparin flush allows easy access for insertion of the cannula. During the flush, closely monitor the outflow from the femoral vein. Once clear fluid exits the femoral vein, stop the flush.</li>
	<li>Wrap the isolated graft in warm saline-soaked gauze and transport it immediately to the recipient animal&#8217;s table. At this time, the recipient surgical site should already be prepared for vascular anastomosis.</li>
	<li>After the graft harvest, immediately euthanize the donor rat via pneumothorax.</li>
</ol>
2. Recipient Transplantation Surgery
<ol>
	<li>Following the sedation induction using vaporized isoflurane applied through a chamber, deeply anesthetize the recipient animal via a ventilator-controlled endotracheal tube and 2.5%&#8211;3% isoflurane. 
	NOTE: At this stage, the donor rat is still anesthetized.</li>
	<li>Continuously monitor the heart rate, respiratory rate, body temperature, and depth of anesthesia of the recipient animal, using the toe pinch withdrawal reflex test.</li>
	<li>In order to prevent dehydration and hypoglycemia, inject 2 mL of lactated Ringer&#8217;s solution and 2.5% dextrose subcutaneously at the beginning and another 2 mL at the end of the surgery.</li>
	<li>Shave the groin area, then treat with depilatory cream in order to reduce the amount of fuzz left by the clippers.</li>
	<li>Wear a mask, a head cover, a disposable isolation gown, and sterile gloves.</li>
	<li>Place the animal supine on a heating pad. Apply ophthalmic ointment to prevent corneal abrasions during anesthesia. Produce a sterile surgical field by prepping, scrubbing, and draping the surgical area.</li>
	<li>Make a 3 cm skin incision in the groin concavity using scalpel blade #15 and reflect the inguinal fat pad laterally using iris scissors.</li>
	<li>Expose the common femoral vessels and place a wire hook with an elastic band to retract the abdominal muscles.</li>
	<li>Ligate and divide Murphy branches.</li>
	<li>Using 10-0 nylon interrupted sutures, anastomose donor vessels to recipient vessels via venous end-to-side technique and arterial end-to-end technique. Gradually release the clamps from the artery and then the vein. Monitor the anastomotic sites for bleeding and add additional sutures if needed.</li>
	<li>Visually assess the vascular anastomosis in order to assure effective graft reperfusion.</li>
	<li>Inset the graft into the inguinal pocket and orient it upside down, with the ankle joint superior and knee joint inferior.</li>
	<li>Using tucking sutures, secure the graft to adjacent muscles. Close the skin via interrupted horizontal mattress skin absorbable 4-0 sutures.</li>
	<li>Remove the recipient animal from anesthesia and wean it off the ventilator. Place the animal on a heating pad for thermal support. 
	NOTE: The overall operation time is between 3 to 4 h, depending on the surgeon&#8217;s experience and acquaintance with the surgical procedure.</li>
	<li>Administer meloxicam (1 mg/kg) subcutaneously&#160;for pain suppression and monitor until the animal is fully recovered and mobile.</li>
</ol>
3. VCA recipient Monitoring
<ol>
	<li>House the recipient rats singly and monitor them daily for clinical signs of pain, dehydration, weight loss, and decreased activity in addition to surgical failure (for the first 48&#8211;72 h) or rejection. Administer meloxicam subcutaneously (1 mg/kg) daily for the first 3 days for pain suppression.</li>
	<li>Based on the research endpoint, choose an immunosuppressant drug to be administered.</li>
</ol>
4. Histology
<ol>
	<li>Under inhaled isoflurane anesthesia (2.5%&#8211;3%), obtain the serial skin and underlying muscle biopsies from the donor graft at desired time points.&#160;The skin should be scrubbed and draped prior to obtaining a biopsy, and a sterile field and technique should performed.</li>
	<li>Close the wound with one to two stitches, using absorbable 4-0 sutures. Return the animal to its cage and allow it to recover from the anesthesia.</li>
	<li>Fix the biopsied tissues in separate tubes in 10% formalin.</li>
	<li>At the terminal time point and under inhaled isoflurane anesthesia (2.5%&#8211;3%), take a larger skin biopsy that spans the donor/recipient border. Carefully locate the vessel leash pair at the site of anastomoses; the proper site will be apparent due to the sutures. Take the desired vessel samples from the artery and/or vein. Fix all samples separately in 10% formalin.&#160;After collection of tissue samples, and while the animal is still under isoflurane anesthesia, immediately euthanize the animal via pneumothorax.</li>
	<li>Using a tissue processor (or other preferred embedding technique), paraffin-embed each biopsy into its own block. For skin samples, orient the tissue so that all epidermal and dermal layers may be seen in a single slice. For vessel samples, orient the vessels so that cross sections may be obtained.</li>
	<li>Using a microtome, cut 6 &#181;m-thick sections and apply them to slides for hematoxylin and eosin (H&#38;E) staining.</li>
	<li>Stain for H&#38;E using a standard protocol.</li>
	<li>Obtain representative images of all desired tissue samples using brightfield microscopy techniques.</li>
</ol>

<ol>
	<li>Gilbert Fernandez, J. J., Febres-Cordero, R. G., Simpson, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Untold+Story+of+the+First+Hand+Transplant:+Dedicated+to+the+Memory+of+one+of+the+Great+Minds+of+the+Ecuadorian+Medical+Community+and+the+World.">The Untold Story of the First Hand Transplant: Dedicated to the Memory of one of the Great Minds of the Ecuadorian Medical Community and the World.</a> Journal of Reconstructive Microsurgery. , (2018).</li><li>Dubernard, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+hand+allograft:+report+on+first+6+months.">Human hand allograft: report on first 6 months.</a> Lancet. 353 (9161), 1315-1320 (1999).</li><li>Jones, J. W., Gruber, S. A., Barker, J. H., Breidenbach, W. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Successful+hand+transplantation.+One-year+follow-up.+Louisville+Hand+Transplant+Team.">Successful hand transplantation. One-year follow-up. Louisville Hand Transplant Team.</a> The New England Journal of Medicine. 343 (7), 468-473 (2000).</li><li>Devauchelle, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=First+human+face+allograft:+early+report.">First human face allograft: early report.</a> Lancet. 368 (9531), 203-209 (2006).</li><li>Broyles, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+abdominal+wall+reconstruction+using+an+innervated+abdominal+wall+vascularized+composite+tissue+allograft:+a+cadaveric+study+and+review+of+the+literature.">Functional abdominal wall reconstruction using an innervated abdominal wall vascularized composite tissue allograft: a cadaveric study and review of the literature.</a> Journal of Reconstructive Microsurgery. 31 (1), 39-44 (2015).</li><li>Kollar, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Innovations+in+reconstructive+microsurgery:+Reconstructive+transplantation.">Innovations in reconstructive microsurgery: Reconstructive transplantation.</a> Journal of Surgical Oncology. 118 (5), 800-806 (2018).</li><li>Kaufman, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Graft+vasculopathy+in+clinical+hand+transplantation.">Graft vasculopathy in clinical hand transplantation.</a> American Journal of Transplantation. 12 (4), 1004-1016 (2012).</li><li>Brandacher, G., Grahammer, J., Sucher, R., Lee, W. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+for+basic+and+translational+research+in+reconstructive+transplantation.">Animal models for basic and translational research in reconstructive transplantation.</a> Birth Defects Research Part C: Embryo Today. 96 (1), 39-50 (2012).</li><li>Kaufman, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunobiology+in+VCA.">Immunobiology in VCA.</a> Transplantation International. 29 (6), 644-654 (2016).</li><li>Committee for the Update of the Guide for the Care and Use of Laboratory Animals, Institute for Laboratory Animal Research, Division on Earth and Life Studies, National Research Council of the National Academies. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Guide for the Care and Use of Laboratory Animals, 8th edition. , (2011).</li><li>Nazzal, J. A., Johnson, T. S., Gordon, C. R., Randolph, M. A., Lee, W. 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The authors have nothing to disclose.

In developing this model of VCA, several key issues were considered. First, it was important to include intact bone (tibia and fibula), bone marrow, and skin in the graft. While clinical hand transplants from adult donors do not transfer significant amounts of actively hematopoietic marrow, studies of the role of the bone marrow niche are better mirrored using an intact, vascularized bone rather than a cut long bone, which results in fibrosis of the exposed marrow. Moreover, the closed bone osteomyocutaneous flap design ...

Reconstructive surgery for the catastrophic tissue loss from amputation, blast injuries, malignancies, and congenital defects are limited by the availability of tissue from the patient and the additional morbidity caused at the donor site. In some cases, such as burn victims or quadrilateral amputees, viable tissue for reconstruction is not available from the patient. In 1964, the first modern hand transplant was performed in Ecuador. While this was a technical success, immunosuppression available at the time was insufficient to prevent rejection, and the graft was lost in less than 3 weeks1. In 1998 and 1999, the first hand transplants in the modern era of immunosuppression were performed in Lyon, France2 and Louisville, Kentucky, USA3. For the first time, reconstructive surgeons could replace like with like. Face transplantation was first performed in 20054, and a number of other VCA grafts are now routinely being performed, such as abdominal wall5, uterine, and urogenital transplants6.
Unlike solid organ transplantation, most VCA techniques involve the presence of the highly antigenic donor skin. Clinical experience has determined that the acute rejection of the skin is relatively easy to control but may contribute to the chronic rejection of the underlying tissues and vessels, which do not respond well to treatment7. The vascular dysfunction associated with an alloimmune response is a more ominous obstacle for the field of VCA7. Macrovasculopathies lead to perfusion deficits, delayed healing, and proinflammatory conditions. Both confluent aggressive large-vessel vasculopathy and focal intimal hyperplasia occur in hand transplant recipients7. Additionally, microvasculopathies likely contribute to VCA complications as well and may even lead to rejection events. While both immune and nonimmune factors likely play a role in the vasculopathy of hand transplant recipients, the specific mechanisms promoting distal vessel dysfunction in VCA are not known, particularly in the context of low-grade, chronic rejection. These unanswered questions necessitate the development of an animal VCA model that will allow for the serial assessment of the graft during the clinical course of VCA rejection/maintenance and vasculopathy. Such a model will offer insights into the rejection and vasculopathy in the face of immunosuppression, infectious challenge, and/or other postoperative traumatic injury8,9.
Presented here is an allogeneic rat VCA heterotopic hindlimb osteomyocutaneous flap model. Based on previously published VCA models, this procedure is technically easy to perform, reproducible in a large number, and exhibits minimal morbidity and discomfort to the recipient animal. This model was designed to allow clinical and histopathological assessments of VCA acceptance vs. rejection, and provides an opportunity to evaluate underlying immune and nonimmune mechanisms involved in rejection.

<table border="0" cellpadding="0" cellspacing="0" style="width:697px;" width="698">
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		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:269px;">Acepromazine</td>
			<td style="width:143px;">Henry Schein</td>
			<td style="width:119px;">5700850</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Adventitia Scissors</td>
			<td>ASSI&#160;</td>
			<td>SAS15R8</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Approximator Clamp (Double)</td>
			<td>ASSI</td>
			<td>ABB2V, ABB22V</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Approximator Clamp (single)</td>
			<td>FST</td>
			<td>00398-02</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Clamp Applying Forceps</td>
			<td>ASSI&#160;</td>
			<td>CAF4</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dissecting Scissors</td>
			<td>ASSI</td>
			<td>SDS18R8</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Flushing blunt needle 27G</td>
			<td>SAI</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Heparin Sodium</td>
			<td>Sagent</td>
			<td>25021-400-30</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isoflurane</td>
			<td>Patterson Veterinary</td>
			<td>14043-704-06</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Jewelers Bipolar</td>
			<td>ASSI</td>
			<td>103000BPS03</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Jewelers forceps #3</td>
			<td>FST</td>
			<td>11231-30</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ketamine HCl 100 mg/ml</td>
			<td>Zoetis</td>
			<td>043-304</td>
			<td>DEA License required</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lactated Ringer Solution</td>
			<td>Hospira</td>
			<td>0409-7953-03</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lactated Ringer Solution + 5% Dextrose</td>
			<td>Hospira</td>
			<td>0409-7953-09</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Meloxicam</td>
			<td>Henry Schein</td>
			<td>11695-6925-2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Micro forceps</td>
			<td>ASSI&#160;</td>
			<td>JFAL3</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Micro needle holder</td>
			<td>ASSI</td>
			<td>B138</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Prograf (Tacrolimus) 5 mg/ml</td>
			<td>Astellas</td>
			<td>0469-3016-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Suture, 10-0 Prolene</td>
			<td>Ethicon</td>
			<td>W2790</td>
			<td>or 10-0 Ethilon (2830)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Suture, 4-0 Coated Vicryl</td>
			<td>Ethicon</td>
			<td>J714D</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vessel Dilator Forceps</td>
			<td>ASSI</td>
			<td>D5AZ</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Xylazine</td>
			<td>VetOne</td>
			<td>13985-612-50</td>
			<td></td>
		</tr>
	</tbody>
</table>

modified heterotopic hindlimb osteomyocutaneous flap model in the rat for translational vascularized composite allotransplantation research

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand and face transplants and, more recently, abdominal wall, uterus, and urogenital transplants. Functional outcomes have exceeded initial expectations, and most recipients enjoy an improved quality of life. However, as clinical experience accumulates, chronic rejection and complications from the immunosuppression must be addressed. In many cases where grafts have failed, the causative pathology has been ischemic vasculopathy. <span style="font-size: 11pt; line-height: 115%; font-family: Arial, sans-serif; color: rgb(41, 43, 49); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">The biological mechanisms of the acute and chronic rejection associated with VCA, especially ischemic vasculopathy, are important areas of research. However, due to the very small number of VCA patients, the evaluation of proposed mechanisms is better addressed in an experimental model. Multiple groups have used animal models to address some of the relevant unsolved questions in VCA rejection and vasculopathy. Several model designs involving a variety of species are described in the literature. Here we present a reproducible model of VCA heterotopic hindlimb osteomyocutaneous flap in the rat that can be utilized for translational VCA research. This model allows for the serial evaluation of the graft, including biopsies and different imaging modalities, while maintaining a low level of morbidity.

The rat VCA heterotopic hindlimb osteomyocutaneous flap model allows for long-term allograft survival under immunosuppression. The model is reliable, reproducible, and simple to perform. The flap is well hidden in the groin area and constitutes minimal morbidity and discomfort to the animal. The skin presentation is a clinical manifestation of the allograft&#8217;s survival and rejection (Figure 1). The flap design allows for gross clinical monitoring and creates an opportunity for various i...

Watch this Scientific Journal Video about Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research at JoVE.com

Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research

Vascularized composite allograft offers life-altering benefits to transplant recipients, but the biological causes of graft rejection and vasculopathy remain poorly understood. The rodent surgical model presented here offers a reproducible, clinically relevant model of transplantation, allowing researchers to evaluate rejection events and potential therapeutic strategies to prevent their occurrence.

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand ...

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5.7K Views.  Christine M. Kleinert Institute for Hand and Microsurgery. Vascularized composite allograft offers life-altering benefits to transplant recipients, but the biological causes of graft rejection and vasculopathy remain poorly understood. The rodent surgical model presented here offers a reproducible, clinically relevant model of transplantation, allowing researchers to evaluate rejection events and potential therapeutic strategies to prevent their occurrence.

Video: Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research

A Modified Heterotopic Swine Hind Limb Transplant Model for Translational Vascularized Composite Allotransplantation (VCA) Research

Vascularized Composite Allotransplantation (VCA) such as hand and face transplants represent a viable treatment option for complex musculoskeletal trauma and devastating tissue loss. Despite favorable and highly encouraging early and intermediate functional outcomes, rejection of the highly immunogenic skin component of a VCA and potential adverse effects of chronic multi-drug immunosuppression continue to hamper widespread clinical application of VCA. Therefore, research in this novel field needs to focus on translational studies related to unique immunologic features of VCA and to develop novel immunomodulatory strategies for immunomodulation and tolerance induction following VCA without the need for long term immunosuppression.
This article describes a reliable and reproducible translational large animal model of VCA that is comprised of an osteomyocutaneous flap in a MHC-defined swine heterotopic hind limb allotransplantation. Briefly, a well-vascularized skin paddle is identified in the anteromedial thigh region using near infrared laser angiography. The underlying muscles, knee joint, distal femur, and proximal tibia are harvested on a femoral vascular pedicle. This allograft can be considered both a VCA and a vascularized bone marrow transplant with its unique immune privileged features. The graft is transplanted to a subcutaneous abdominal pocket in the recipient animal with a skin component exteriorized to the dorsolateral region for immune monitoring.
Three surgical teams work simultaneously in a well-coordinated manner to reduce anesthesia and ischemia times, thereby improving efficiency of this model and reducing potential confounders in experimental protocols. This model serves as the groundwork for future therapeutic strategies aimed at reducing and potentially eliminating the need for chronic multi-drug immunosuppression in VCA.

A Modified Heterotopic Swine Hind Limb Transplant Model for Translational Vascularized Composite Allotransplantation VCA Research

Vascularized Composite Allotransplantations (VCA) have become a clinical reality. However, broad clinical application of VCA is limited by chronic multi-drug immunosuppression. The authors present a reliable and reproducible large animal model to translate novel immunomodulatory strategies that can minimize or potentially eliminate the need of immunosuppression in VCA.

Vascularized Composite Allotransplantation (VCA) such as hand and face transplants represent a viable treatment option for complex musculoskeletal trauma ...

A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression must be pursued. Employing vascularized bone marrow transplantation and co-transplantation of the thymus have shown promise in this regard in various animal models.1-11 Vascularized bone marrow transplantation allows for the uninterrupted transfer of donor bone marrow cells within the preserved donor microenvironment, and the incorporation of thymus tissue with vascularized bone marrow transplantation has shown to increase T-cell chimerism ultimately playing a supportive role in the induction of immune regulation. The combination of solid organ and vascularized composite allotransplantation can uniquely combine these strategies in the form of a novel transplant model. Murine models serve as an excellent paradigm to explore the mechanisms of acute and chronic rejection, chimerism, and tolerance induction, thus providing the foundation to propagate superior allograft survival strategies for larger animal models and future clinical application. Herein, we developed a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique. The experience in syngeneic and allogeneic transplant settings is described for future broader immunological investigations via an instructional manuscript and video supplement.

To study combined solid organ and vascularized composite allotransplantation, we describe a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique.

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression ...

Orthotopic Hind Limb Transplantation in the Mouse

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and translational research in the field of transplantation medicine. A vast array of reagents is available exclusively in this setting, including mono- and polyclonal antibodies for both diagnostic and interventional applications. In addition, a vast number of genotyped, inbred, transgenic, and knock out strains allow detailed investigation of the individual contributions of humoral and cellular components to the complex interplay of an immune response and make the mouse the gold standard for immunological research. 
Vascularized Composite Allotransplantation (VCA) delineates a novel field of transplantation using allografts to replace "like with like" in patients suffering traumatic or congenital tissue loss. This surgical methodological protocol shows the use of a non-suture cuff technique for super-microvascular anastomosis in an orthotopic mouse hind limb transplantation model. The model specifically allows for comparison between established paradigms in solid organ transplantation with a novel form of transplants consisting of various different tissue components. Uniquely, this model allows for the transplantation of a viable vascularized bone marrow compartment and niche that have the potential to exert a beneficial effect on the balance of immune acceptance and rejection. This technique provides a tool to investigate alloantigen recognition and allograft rejection and acceptance, as well as enables the pursuit of functional nerve regeneration studies to further advance this novel field of transplantation.

This novel model for orthotopic hind limb transplantation in the mouse, applying a non-suture cuff technique for super-microvascular anastomosis, provides a powerful tool for in&#160;vivo mechanistic immunological research related to vascularized composite allotransplantation (VCA).

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and ...

Mouse Model for Pancreas Transplantation Using a Modified Cuff Technique

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the availability of the widest range of molecular probes and reagents to perform in vivo as well as in vitro studies. Based on our experience with various murine transplantation models, we developed a heterotopic pancreas transplantation model in mice with the intent to analyze mechanisms underlying severe ischemia reperfusion injury-associated early graft damage. In contrast to previously described techniques using suture techniques, herein we describe a new procedure using a non-suture cuff technique. 
In recent years, we have performed more than 300 pancreas transplantations in mice with an overall success rate of &gt;90%, a success rate never described before in mouse pancreas transplantation. The backbone of this non-suture cuff technique for graft revascularization consists of two major steps: (I) pulling the recipient vessel over a polyethylene/polyamide cuff and fixing it with a circumferential ligature, and (II) placing the donor vessel over the everted recipient vessel and fixing it with a second circumferential ligature. The resultant continuity of the endothelial layer results in less thrombogenic lesions with high patency rates and, finally, high success rates.
In this model, arterial anastomosis is achieved by pulling the abdominal aorta of the donor graft over the everted common carotid artery of the recipient animal. Venous drainage of the graft is achieved by pulling the portal vein of the graft over the everted external jugular vein of the recipient. This manuscript provides details and crucial steps of the organ recovery and organ implantation procedures, which will allow researchers with microsurgical skills to perform the transplantation successfully in their laboratories.

Among abdominal solid organ transplantation, pancreatic grafts are prone to develop severe ischemia reperfusion injury-associated graft damage, leading eventually to early graft loss. This protocol describes a model of murine pancreas transplantation using a non-suture cuff technique, ideally suited for analyzing these early, deleterious damages.

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the ...

Surgical Angiogenesis in Porcine Tibial Allotransplantation: A New Large Animal Bone Vascularized Composite Allotransplantation Model

Segmental bone loss resulting from trauma, infection malignancy and congenital anomaly remains a major reconstructive challenge. Current therapeutic options have significant risk of failure and substantial morbidity.
Use of bone vascularized composite allotransplantation (VCA) would offer both a close match of resected bone size and shape and the healing and remodeling potential of living bone. At present, life-long drug immunosuppression (IS) is required. Organ toxicity, opportunistic infection and neoplasm risks are of concern to treat such non-lethal indications.
We have previously demonstrated that bone and joint VCA viability may be maintained in rats and rabbits without the need of long-term-immunosuppression by implantation of recipient derived vessels within the VCA. It generates an autogenous, neoangiogenic circulation with measurable flow and active bone remodeling, requiring only 2 weeks of IS. As small animals differ from man substantially in anatomy, bone physiology and immunology, we have developed a porcine bone VCA model to evaluate this technique before clinical application is undertaken. Miniature swine are currently widely used for allotransplantation research, given their immunologic, anatomic, physiologic and size similarities to man. Here, we describe a new porcine orthotopic tibial bone VCA model to test the role of autogenous surgical angiogenesis to maintain VCA viability.
The model reconstructs segmental tibial bone defects using size- and shape-matched allogeneic tibial bone segments, transplanted across a major swine leukocyte antigen (SLA) mismatch in Yucatan miniature swine. Nutrient vessel repair and implantation of recipient derived autogenous vessels into the medullary canal of allogeneic tibial bone segments is performed in combination with simultaneous short-term IS. This permits a neoangiogenic autogenous circulation to develop from the implanted tissue, maintaining flow through the allogeneic nutrient vessels for a short time. Once established, the new autogenous circulation maintains bone viability following cessation of drug therapy and subsequent nutrient vessel thrombosis.

Currently any kind of vascularized composite allotransplantation depends on long-term-immunosuppression, difficult to support for non-life-critical indications. We present a new porcine tibial VCA model that can be used to study bone VCA and demonstrate the use of surgical angiogenesis to maintain bone viability without the need of long-term immune-modulation.

Segmental bone loss resulting from trauma, infection malignancy and congenital anomaly remains a major reconstructive challenge. Current therapeutic ...

A Model of Free Tissue Transfer: The Rat Epigastric Free Flap

Free tissue transfer has been increasingly used in clinical practice since the 1970s, allowing reconstruction of complex and otherwise untreatable defects resulting from tumor extirpation, trauma, infections, malformations or burns. Free flaps are particularly useful for reconstructing highly complex anatomical regions, like those of the head and neck, the hand, the foot and the perineum. Moreover, basic and translational research in the area of free tissue transfer is of great clinical potential. Notwithstanding, surgical trainees and researchers are frequently deterred from using microsurgical models of tissue transfer, due to lack of information regarding the technical aspects involved in the operative procedures. The aim of this paper is to present the steps required to transfer a fasciocutaneous epigastric free flap to the neck in the rat.
This flap is based on the superficial epigastric artery and vein, which originates from and drain into the femoral artery and vein, respectively. On average the caliber of the superficial epigastric vein is 0.6 to 0.8 mm, contrasting with the 0.3 to 0.5 mm of the superficial epigastric artery. Histologically, the flap is a composite block of tissues, containing skin (epidermis and dermis), a layer of fat tissue (panniculus adiposus), a layer of striated muscle (panniculus carnosus), and a layer of loose areolar tissue.
Succinctly, the epigastric flap is raised on its pedicle vessels that are then anastomosed to the external jugular vein and to the carotid artery on the ventral surface of the rat's neck. According to our experience, this model guarantees the complete survival of approximately 70 to 80% of epigastric flaps transferred to the neck region. The flap can be evaluated whenever needed by visual inspection. Hence, the authors believe this is a good experimental model for microsurgical research and training.

This paper describes the steps required to raise a fasciocutaneous epigastric free flap and transfer it to the neck in the rat.

Free tissue transfer has been increasingly used in clinical practice since the 1970s, allowing reconstruction of complex and otherwise untreatable defects ...

Development of a Novel Task-oriented Rehabilitation Program using a Bimanual Exoskeleton Robotic Hand

A robot-assisted hand is used for the rehabilitation of patients with impaired upper limb function, particularly for stroke patients with a loss of motor control. However, it is unclear how conventional occupational training strategies can be applied to the use of rehabilitation robots. Novel robotic technologies and occupational therapy concepts are used to develop a protocol that allows patients with impaired upper limb function to grasp objects using their affected hand through a variety of pinching and grasping functions. To conduct this appropriately, we used five types of objects: a peg, a rectangular cube, a cube, a ball, and a cylindrical bar. We also equipped the patients with a robotic hand, the Mirror Hand, an exoskeleton&#160;hand that is fitted to the subject&#8217;s affected hand and follows the movement of the sensor glove fitted to their unaffected hand (bimanual movement training (BMT)). This study had two stages. Three healthy subjects were first recruited to test the feasibility and acceptability of the training program. Three patients with hand dysfunction caused by stroke were then recruited to confirm the feasibility and acceptability of the training program, which was conducted on 3 consecutive days. On each day, the patient was monitored during 5 min of movement in a passive range of motion, 5 min of robot-assisted bimanual movement, and task-oriented training using the five objects. The results showed that both healthy subjects and subjects who had suffered a stroke in conjunction with the robotic hand could successfully grasp the objects. Both healthy subjects and those who had suffered a stroke performed well with the robot-assisted task-oriented training program in terms of feasibility and acceptability.

This study reports the development of a novel robot-assisted task-oriented program for hand rehabilitation. The developmental process consists of experiments using both healthy subjects and subjects who have had a stroke and suffered from subsequent motor control dysfunction.

A robot-assisted hand is used for the rehabilitation of patients with impaired upper limb function, particularly for stroke patients with a loss of motor ...

Hickman Catheter for Long-Term Vascular Access in Swine

Hickman Catheter Use for Long-Term Vascular Access in a Preclinical Swine Model

Central venous catheters (CVCs) are invaluable devices in large animal research as they facilitate a wide range of medical applications, including blood monitoring and reliable intravenous fluid and drug administration. Specifically, the tunneled multi-lumen Hickman catheter (HC) is commonly used in swine models due to its lower extrication and complication rates. Despite fewer complications relative to other CVCs, HC-related morbidity presents a significant challenge, as it can significantly delay or otherwise negatively impact ongoing studies. The proper insertion and maintenance of HCs is paramount in preventing these complications, but there is no consensus on best practices. The purpose of this protocol is to comprehensively describe an approach for the insertion and maintenance of a tunneled HC in swine that mitigates HC-related complications and morbidity. The use of these techniques in &gt;100 swine has resulted in complication-free patent lines up to 8 months and no catheter-related mortality or infection of the ventral surgical site. This protocol offers a method to optimize the lifespan of the HC and guidance for approaching issues during use.

Author Spotlight: Hickman Catheter Use for Long-Term Vascular Access in a Preclinical Swine Model  

A reliable and reproducible approach for the insertion and maintenance of a tunneled Hickman catheter for long-term vascular access in swine is described. Placement of a central venous catheter allows for convenient daily sampling of whole blood from awake animals and intravenous administration of medication and fluids.

Central venous catheters (CVCs) are invaluable devices in large animal research as they facilitate a wide range of medical applications, including blood ...

Rat Model of Normothermic Ex-Situ Perfused Heterotopic Heart Transplantation

Heart transplantation is the most effective therapy for end-stage heart failure. Despite the improvements in therapeutic approaches and interventions, the number of heart failure patients waiting for transplantation is still increasing. The normothermic ex situ preservation technique has been established as a comparable method to the conventional static cold storage technique. The main advantage of this technique is that donor hearts can be preserved for up to 12 h in a physiologic condition. Moreover, this technique allows resuscitation of the donor hearts after circulatory death and applies required pharmacologic interventions to improve donor function after implantation. Numerous animal models have been established to improve normothermic ex situ preservation techniques and eliminate preservation-related complications. Although large animal models are easy to handle compared to small animal models, it is costly and challenging. We present a rat model of normothermic ex situ donor heart preservation followed by heterotopic abdominal transplantation. This model is relatively cheap and can be accomplished by a single experimenter.

Here, we present an assessment protocol of a heterotopically implanted heart after normothermic ex situ preservation in the rat model.

Heart transplantation is the most effective therapy for end-stage heart failure. Despite the improvements in therapeutic approaches and interventions, the ...

Modified Heterotopic Abdominal Heart Transplantation and a Novel Aortic Regurgitation Model in Rats

Over the past 50 years, many researchers have reported heterotopic abdominal heart transplantation in mice and rats, with some variations in the surgical technique. Modifying the transplantation procedure to strengthen the myocardial protection could prolong the ischemia time while preserving the donor&#39;s cardiac function. This technique&#39;s key points are as follows: transecting the donor&#39;s abdominal aorta before harvesting to unload the donor&#39;s heart; perfusing the donor&#39;s coronary arteries with a cold cardioplegic solution; and topical cooling of the donor&#39;s heart during the anastomosis procedure. Consequently, since this procedure prolongs the acceptable ischemia time, beginners can easily perform it and achieve a high success rate.
Moreover, a new aortic regurgitation (AR) model was established in this work using a technique different from the existing one,&#160;which is created by inserting a catheter from the right carotid artery and puncturing the native aortic valve under continuous echocardiographic guidance. A heterotopic abdominal heart transplantation was performed using the novel AR model. In the protocol, after the donor&#39;s heart is harvested, a stiff guidewire is inserted into the donor&#39;s brachiocephalic artery and advanced toward the aortic root. The aortic valve is punctured by pushing the guidewire further even after the resistance is felt, thus inducing AR. It is easier to damage the aortic valve using this method than with the procedure described in the conventional AR model. Additionally, this novel AR model does not contribute to the recipient&#39;s circulation; therefore, this method is expected to produce a more severe AR model than the conventional procedure.

This study demonstrates a reproducible heterotopic abdominal heart transplantation technique in rats that beginners can learn and perform. Additionally, a novel aortic regurgitation model in rats is generated by performing heterotopic abdominal heart transplantation and damaging the donor's aortic valve using a guidewire after harvesting.

Over the past 50 years, many researchers have reported heterotopic abdominal heart transplantation in mice and rats, with some variations in the surgical ...