All experiments were performed according to the guidelines of the German animal welfare act (TierSchG.) (AZ: 55.2-1-54-2532.Vet_02-80-2015).
1. Animal housing
<ol>
	<li>For experiments, use male C57BL/6 and BALB/c mice weighing 20-25 g with C57BL/6 mice as the recipient animals and BALB/c mice as the donor animals.</li>
	<li>Purchase the animals and house in a barrier pathogen-free facility, in accordance with the FELASA guidelines for health monitoring12.</li>
	<li>Keep the mice in standard Makrolon cages with enrichment nesting material. Provide ad libitum access to water and pelleted food at a day/night cycle of 12 h.</li>
	<li>Maintain the room temperature at 22 &#177; 2&#176;C and the relative humidity at 55 &#177; 5%.</li>
</ol>
2. Recipient preparation
<ol>
	<li>First, anesthetize the recipient animal (C57BL/6) with an intraperitoneal injection of midazolam (5 mg/kg; 5 mg/mL), medetomidin (0.5 mg/kg; 1 mg/mL) and fentanyl (0.05 mg/kg; 0.05 mg/mL). 
	NOTE: The correct depth of the anesthesia should be reached in 5-10 min.
	<ol>
		<li>Pinch the hind feet with forceps to check for reflexes to confirm the depth of anesthesia.</li>
	</ol>
	</li>
	<li>Clip all the hair of the cervical lateral region with an electric hair clipper for small animals and apply ophthalmic ointment with cotton swabs to prevent the eyes from drying out during the procedure.</li>
	<li>Place the animal in a supine position on a heating pad under the microscope and gently tape its legs to the operating table with skin sensitive plaster strips.</li>
	<li>Tilt its head back and scrub the operative field several times with alcohol.</li>
	<li>Make a skin incision from the jugular incision to the right lower mandible with small scissors.</li>
	<li>Remove the right lower lobe of the submandibular gland via bipolar cautery of the vessel pedicle and subsequent excision with microscissors.</li>
	<li>Remove the right sternocleidomastoid muscle via bipolar cautery of the upper and lower portion and subsequent excision with microscissors to gain access to the common carotid artery.</li>
	<li>Mobilize the common carotid artery as far distally and proximally as possible by pulling the surrounding connective tissue apart with fine forceps.</li>
	<li>Tie two 7-0 silk ligatures with minimal distance between each around the middle of the common carotid artery and transect the common carotid artery with fine microscissors between the ligatures.</li>
	<li>Pass the proximal, ligated end through the cuff and fix it with a small artery clamp. 
	NOTE: The cuffs that were used in this procedure were cut with microscissors from tubes of polyimide with an outer diameter of 0.610 mm and a wall thickness of 0.0254 mm. The completed cuffs had a length of ~2 mm with one half, which is used for the cuffing process, being a full cylinder and the other half, which is clamped, being a half-cylinder.</li>
	<li>Remove the ligature with fine microscissors, as close to the ligature as possible, and flush the lumen with heparinized saline (50 IU/mL) with a 30 G needle, while taking care not to damage the vessel walls.</li>
	<li>Distend the open lumen using fine vascular dilatators and evert the carotid stump over the cuff by pulling it gently over the polyimide tube.</li>
	<li>Immediately fix the everted carotid with a loosely pre-tied 7-0 silk loop. 
	NOTE: Loosely pre-tie 4 7-0 silk loops with a diameter of about 1.5 mm before the surgery to make the cuffing-procedure smoother and easier.</li>
	<li>Perform the same procedure (2.10-2.13) at the other end of the carotid artery.</li>
	<li>Set the recipient animal aside and moisturize the operative field with saline until the aortic segment is explanted.</li>
</ol>
3. Donor operation
<ol>
	<li>Anesthetize the donor mouse (BALB/c) in the same fashion as the recipient animal.
	<ol>
		<li>Pinch the hind feet with forceps to check for reflexes to confirm sufficient anesthesia.</li>
	</ol>
	</li>
	<li>Clip all hair of the abdominal and thoracic region with an electric hair clipper for small animals and apply ophthalmic ointment with cotton swabs to prevent the eyes from drying out during the procedure.</li>
	<li>Place the animal in a supine position on a heating pad under the microscope and gently tape its legs to the operating table with skin sensitive plaster strips.</li>
	<li>Scrub the operative field several times with alcohol.</li>
	<li>Perform a midline abdominal laparotomy with small scissors and push the intestines slightly upward to expose the inferior vena cava (IVC).</li>
	<li>Inject the inferior vena cava (IVC) with 1 mL of heparinized saline using a 30 G needle.</li>
	<li>Cut the abdominal aorta and IVC below the renal arteries with small scissors to exsanguinate the donor animal. Loosely place a compress into the abdomen to absorb the blood.</li>
	<li>Perform a thoracotomy at the bilateral diversion of the ribs with scissors and tilt the anterior chest wall cranially with a surgical clamp to expose the mediastinum.</li>
	<li>Cut the IVC and the esophagus directly above the diaphragm with microscissors.</li>
	<li>Remove the heart and the lungs by tilting them upward with forceps holding the cut IVC/esophagus and then excising them with microscissors from their base to gain access to the thoracic aorta in the dorsal mediastinum.</li>
	<li>Mobilize the thoracic aorta from its surrounding tissue by pulling apart the surrounding connective tissue and fat with fine forceps while being careful not to damage any intercostal arteries.</li>
	<li>Cauterize all branches from the thoracic aorta with bipolar cautery forceps and excise the aortic segment between the diaphragm and the aortic arch using microscissors.</li>
	<li>Flush the excised aortic segment with heparinized saline with a 30 G needle, while taking care not to damage the vessel walls, to remove any remaining blood or clots, and transfer the graft to the recipient animal. 
	NOTE: Directly place the aortic graft in the roughly right position in the recipient during transfer. If there are problems confusing the different ends of the graft in the recipient animal, a loose ligature around the distal end could help.</li>
</ol>
4. Implantation
<ol>
	<li>Pull the proximal end of the donor aortic segment over the proximal cuff on top of the everted carotid artery with fine forceps and immediately fix it with a loosely pre-tied 7-0 silk loop.</li>
	<li>Trim the distal, free end of the aortic graft with microscissors so that the graft length fits the distance between the two cuffs.</li>
	<li>Repeat step 4.1 on the other end of the aorta with the other cuff to complete the anastomosis.</li>
	<li>Remove the distal clamp to allow retrograde perfusion.</li>
	<li>After achieving hemostasis, remove the proximal clamp to complete the anastomosis.</li>
	<li>Finally, close the wound with 6-0 continuous suture.</li>
</ol>
5. Postoperative care
<ol>
	<li>Monitor the mouse closely in the first 6 h after the operation and then several times a day for the first 72 h after the transplantation to detect any complications instantly.</li>
	<li>For postoperative analgesia, inject the transplanted mouse with buprenorphine (0.05-0.1 mg/kg) subcutaneously directly after the transplantation and then every 12 h for 72 h to provide appropriate, long term analgesia.</li>
</ol>
6. Aortic graft explanations
<ol>
	<li>Anesthetize the transplanted animal with an intraperitoneal injection of midazolam (5 mg/kg; 5 mg/mL), medetomidin (0.5 mg/kg; 1 mg/mL) and fentanyl (0.05 mg/kg; 0.05 mg/mL) 4 weeks after transplantation.
	<ol>
		<li>Pinch the hind feet with forceps to check for reflexes to confirm sufficient anesthesia.</li>
	</ol>
	</li>
	<li>Clip all hair of the abdominal, thoracic and cervical region with an electric hair clipper for small animals.</li>
	<li>Place the animal in a supine position on a heating pad under the microscope and gently tape its legs to the operating table with skin sensitive plaster strips.</li>
	<li>Scrub the operative field several times with alcohol.</li>
	<li>Perform a midline abdominal laparotomy with small scissors and push the intestines slightly upward to expose the inferior vena cava (IVC).</li>
	<li>Inject the inferior vena cava (IVC) with 1 mL of heparinized saline using a 30 G needle.</li>
	<li>Cut the abdominal aorta and IVC below the renal arteries with small scissors to exsanguinate the donor animal. Loosely place a compress into the abdomen to absorb the blood.</li>
	<li>Make a skin incision from the jugular incision to the right lower mandible with small scissors corresponding to the skin incision made during the transplant procedure.</li>
	<li>Identify the transplanted aortic graft together with the distal and proximal cuff and blunt remove any surrounding tissue with forceps.</li>
	<li>Using microscissors, cut through the common carotid artery distal and proximal to the aortic graft with the cuffs in order to explant the aortic graft together with the two cuff ends.</li>
	<li>Cut the aortic segment in half and preserve the specimens for further analyses (frozen sections, paraffin embedded sections, snap frozen material)13,14.</li>
</ol>

<ol>
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R., Kaplan, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lack+of+improvement+in+renal+allograft+survival+despite+a+marked+decrease+in+acute+rejection+rates+over+the+most+recent+era.">Lack of improvement in renal allograft survival despite a marked decrease in acute rejection rates over the most recent era.</a> American Journal of Transplantation. 4 (3), 378-383 (2004).</li><li>Bagnasco, S. M., Kraus, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intimal+arteritis+in+renal+allografts:+new+takes+on+an+old+lesion.">Intimal arteritis in renal allografts: new takes on an old lesion.</a> Current Opinion in Organ Transplantation. 20 (3), 343-347 (2015).</li><li>Hollis, I. B., Reed, B. N., Moranville, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Medication+management+of+cardiac+allograft+vasculopathy+after+heart+transplantation.">Medication management of cardiac allograft vasculopathy after heart transplantation.</a> Pharmacotherapy. 35 (5), 489-501 (2015).</li><li>Verleden, G. M., Raghu, G., Meyer, K. C., Glanville, A. R., Corris, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+classification+system+for+chronic+lung+allograft+dysfunction.">A new classification system for chronic lung allograft dysfunction.</a> The Journal of Heart and Lung Transplantation. 33 (2), 127-133 (2014).</li><li>Ramzy, D., 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=Cardiac+allograft+vasculopathy:+a+review.">Cardiac allograft vasculopathy: a review.</a> Canadian Journal of Surgery. 48 (4), 319-327 (2005).</li><li>Skoric, 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=Cardiac+allograft+vasculopathy:+diagnosis,+therapy,+and+prognosis.">Cardiac allograft vasculopathy: diagnosis, therapy, and prognosis.</a> Croatian Medical Journal. 55 (6), 562-576 (2014).</li><li>Koulack, J., 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=Development+of+a+mouse+aortic+transplant+model+of+chronic+rejection.">Development of a mouse aortic transplant model of chronic rejection.</a> Microsurgery. 16 (2), 110-113 (1995).</li><li>Rowinska, Z., 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=Using+the+Sleeve+Technique+in+a+Mouse+Model+of+Aortic+Transplantation+-+An+Instructional+Video.">Using the Sleeve Technique in a Mouse Model of Aortic Transplantation - An Instructional Video.</a> Journal of Visualized Experiments. (128), (2017).</li><li>Dietrich, H., 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=Mouse+model+of+transplant+arteriosclerosis:+role+of+intercellular+adhesion+molecule-1.">Mouse model of transplant arteriosclerosis: role of intercellular adhesion molecule-1.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 20 (2), 343-352 (2000).</li><li>M&#228;hler Convenor, 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=FELASA+recommendations+for+the+health+monitoring+of+mouse,+rat,+hamster,+guinea+pig+and+rabbit+colonies+in+breeding+and+experimental+units.">FELASA recommendations for the health monitoring of mouse, rat, hamster, guinea pig and rabbit colonies in breeding and experimental units.</a> Laboratory Animals. 48 (3), 178-192 (2014).</li><li>Ollinger, R., 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=Blockade+of+p38+MAPK+inhibits+chronic+allograft+vasculopathy.">Blockade of p38 MAPK inhibits chronic allograft vasculopathy.</a> Transplantation. 85 (2), 293-297 (2008).</li><li>Thomas, M. N., 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=SDF-1/CXCR4/CXCR7+is+pivotal+for+vascular+smooth+muscle+cell+proliferation+and+chronic+allograft+vasculopathy.">SDF-1/CXCR4/CXCR7 is pivotal for vascular smooth muscle cell proliferation and chronic allograft vasculopathy.</a> Transplant International. 28 (12), 1426-1435 (2015).</li><li>Ollinger, R., 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=Bilirubin:+a+natural+inhibitor+of+vascular+smooth+muscle+cell+proliferation.">Bilirubin: a natural inhibitor of vascular smooth muscle cell proliferation.</a> Circulation. 112 (7), 1030-1039 (2005).</li><li>Segura, A. M., Buja, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+allograft+vasculopathy:+a+complex+multifactorial+sequela+of+heart+transplantation.">Cardiac allograft vasculopathy: a complex multifactorial sequela of heart transplantation.</a> Texas Heart Institute Journal. 40 (4), 400-402 (2013).</li><li>McDaid, J., Scott, C. J., Kissenpfennig, A., Chen, H., Martins, P. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+utility+of+animal+models+in+developing+immunosuppressive+agents.">The utility of animal models in developing immunosuppressive agents.</a> European Journal of Pharmacology. 759, 295-302 (2015).</li><li>Shi, C., Russell, M. E., Bianchi, C., Newell, J. B., Haber, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+model+of+accelerated+transplant+arteriosclerosis.">Murine model of accelerated transplant arteriosclerosis.</a> Circulation Research. 75 (2), 199-207 (1994).</li><li>Koulack, J., 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=Importance+of+minor+histocompatibility+antigens+in+the+development+of+allograft+arteriosclerosis.">Importance of minor histocompatibility antigens in the development of allograft arteriosclerosis.</a> Clinical Immunology and Immunopathology. 80 (3 Pt 1), 273-277 (1996).</li><li>Maglione, 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=A+novel+technique+for+heterotopic+vascularized+pancreas+transplantation+in+mice+to+assess+ischemia+reperfusion+injury+and+graft+pancreatitis.">A novel technique for heterotopic vascularized pancreas transplantation in mice to assess ischemia reperfusion injury and graft pancreatitis.</a> Surgery. 141 (5), 682-689 (2007).</li><li>Oberhuber, R., 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=Murine+cervical+heart+transplantation+model+using+a+modified+cuff+technique.">Murine cervical heart transplantation model using a modified cuff technique.</a> Journal of Visualized Experiments. (92), e50753 (2014).</li><li>Nakao, A., Ogino, Y., Tahara, K., Uchida, H., Kobayashi, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orthotopic+intestinal+transplantation+using+the+cuff+method+in+rats:+a+histopathological+evaluation+of+the+anastomosis.">Orthotopic intestinal transplantation using the cuff method in rats: a histopathological evaluation of the anastomosis.</a> Microsurgery. 21 (1), 12-15 (2001).</li></ol>

The authors declare that they have no competing financial interests.

Chronic allograft vasculopathy is the major cause of late graft loss after solid organ transplantation of the heart and likely renal and lung allografts8. Thus far, no causal therapeutic regimen could be developed in order to prevent the formation of CAV.
The pathophysiology of CAV is multifactorial and involves immunological and non-immunological aspects16. The use of rodent models in transplantation have been essential in understanding the unde...

Over the past six decades, solid organ transplantation has evolved from an experimental procedure to a standard of care for the treatment of end-stage organ failure1. Due to the improvement of antimicrobial agents, surgical techniques and advancement in immunosuppressive regiments, the early success rate of solid organ transplantation have significantly increased over the past decades2.
However, long-term graft survival rates have not significantly improved in the same manner3. The development of CAV is the major factor limiting long-term survival4,5,6. This pathology is characterized by the formation of a concentric neointimal layer consisting of smooth muscle cells, leading to progressive narrowing of the vessel and consecutive malperfusion of the transplanted solid organ. In heart transplant recipients, CAV lesions can be diagnosed in up to 75% of patients 3 years after transplantation7.
The pathophysiology of CAV is not fully understood yet. It seems to be related to numerous immunological and non-immunological factors, leading to endothelial damage with subsequent endothelial activation and dysfunction8. Thus far, no causal treatment option exists for the prevention of CAV, emphasizing the need for a reproducible small animal model in order to study the formation and potential therapy of CAV.
With the use of murine aortic transplantation models, CAV like lesions can be seen 4 weeks after transplantation. Those lesions consist mainly of vascular smooth muscle cells, thereby, resembling the human pathology. Because of a wide variety of transgenic and knock out mice, the use of mouse models in transplant associated pathologies offers a unique opportunity to identify new therapeutic options and understand their development. Due to the small diameter of the transplanted vessels however, the use of mouse models is commonly associated with long learning curves and an initial high complication rate9. With the introduction of the non-suture cuff technique, this most challenging part of the operation can be facilitated and the diameter of the anastomosis is kept constant10,11.

<table>
	<tbody>
		<tr>
			<td>Balb-c Mice (H2-d)</td>
			<td>Charles River</td>
			<td>Strain# 028</td>
			<td>Donor animal</td>
		</tr>
		<tr>
			<td>Bipolar cautery system</td>
			<td>ERBE</td>
			<td>ICC 50 / 20195-023</td>
			<td>Bipolar cautery</td>
		</tr>
		<tr>
			<td>C57BL/6J (H-2b)</td>
			<td>Charles River</td>
			<td>Strain# 027</td>
			<td>Recipient animal</td>
		</tr>
		<tr>
			<td>Halsey Needle Holders</td>
			<td>FST</td>
			<td>12501-12</td>
			<td>Needle Holder</td>
		</tr>
		<tr>
			<td>Halsted-Mosquito Forceps</td>
			<td>AESCULAP</td>
			<td>BH111R</td>
			<td>Curved Clamp</td>
		</tr>
		<tr>
			<td>Medical Polyimide Tubing</td>
			<td>Nordson MEDICAL</td>
			<td>141-0031</td>
			<td>Cuff-Material</td>
		</tr>
		<tr>
			<td>Micro Serrefines</td>
			<td>FST</td>
			<td>18055-04</td>
			<td>Micro Vessel Clip</td>
		</tr>
		<tr>
			<td>Micro-Adson Forceps (serrated)</td>
			<td>FST</td>
			<td>11018-12</td>
			<td>Standard Forceps</td>
		</tr>
		<tr>
			<td>Micro-Serrefine Clamp Applying Forceps</td>
			<td>FST</td>
			<td>18057-14</td>
			<td>Clipapplicator</td>
		</tr>
		<tr>
			<td>S&#38;T Forceps - SuperGrip Tips (Angled 45&#176;)</td>
			<td>S&#38;T</td>
			<td>00649-11</td>
			<td>Fine Forceps</td>
		</tr>
		<tr>
			<td>S&#38;T Vessel Dilating Forceps - Angled 10&#176; (Tip diameter 0.2 mm)</td>
			<td>S&#38;T</td>
			<td>00125-11</td>
			<td>Vesseldilatator</td>
		</tr>
		<tr>
			<td>Schott VisiLED Set</td>
			<td>Schott</td>
			<td>MC 1500 / S80-55</td>
			<td>Light</td>
		</tr>
		<tr>
			<td>Stereoscopic microscope</td>
			<td>ZEISS</td>
			<td>SteREO Discovery.V8</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>Student Fine Scissors / Surgical Scissors - Sharp-Blunt</td>
			<td>FST</td>
			<td>91460-11 / 14001-12</td>
			<td>Standard Sissors</td>
		</tr>
		<tr>
			<td>Vannas-T&#252;bingen Spring Scissors (curved, 8.5 cm)</td>
			<td>FST</td>
			<td>15004-08</td>
			<td>Microsissors (curved)</td>
		</tr>
		<tr>
			<td>Vannas-T&#252;bingen Spring Scissors (straight, 8.5 cm)</td>
			<td>FST</td>
			<td>15003-08</td>
			<td>Microsissors (straight)</td>
		</tr>
	</tbody>
</table>

murine cervical aortic transplantation model using a modified non-suture cuff technique

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. However, only minor improvement in the long-term results of transplanted solid organs could be observed over the past decades. In this context, chronic allograft vasculopathy (CAV) still represents the leading cause of late organ failure in cardiac, renal and pulmonary transplantation.
Thus far, the underlying pathogenesis of CAV development remains unclear, explaining why effective treatment strategies are presently missing and emphasizing a need for relevant experimental models in order to study the underlying pathophysiology leading to CAV formation. The following protocol describes a murine heterotopic cervical aortic transplantation model using a modified non-suture cuff technique. In this technique, a segment of the thoracic aorta is interpositioned in the right common carotid artery. With the use of the non-suture cuff technique, an easy to learn and reproducible model can be established, minimizing the possible heterogeneity of sutured vascular micro anastomoses.

In the fully MHC-mismatch transplantation model, a concentric neointimal layer can be seen 4 weeks after transplantation (Figure 2). This layer consists primarily of vascular smooth muscle cells as immunohistological staining for SM22 (a selective marker for mature vascular smooth muscle cells) revealed. As stated before, these vascular smooth muscle cells are pathognomonic for lesions seen in chronic allograft vasculopathy. For further analyses, aortic segments should be sectioned and stain...

Watch this Scientific Journal Video about Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique at JoVE.com

Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. ...

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6.2K Views.  Ludwig Maximilian University of Munich. Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

Video: Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant vasculopathy (TVP).
The commonly used animal model for cardiovascular chronic rejection studies is the heterotopic heart transplant model performed in laboratory rodents. This model is used widely in experiments since Ono and Lindsey (3) published their technique. To analyze the findings in the blood vessels, the heart has to be sectioned and all vessels have to be measured.
Another method to investigate chronic rejection in cardiovascular questionings is the aortic transplant model (1, 2). In the orthotopic aortic transplant model, the aorta can easily be histologically evaluated (2). The PVG-to-ACI model is especially useful for CAV studies, since acute vascular rejection is not a major confounding factor and Cyclosporin A (CsA) treatment does not prevent the development of CAV, similar to what we find in the clinical setting (4). A7-day period of CsA is required in this model to prevent acute rejection and to achieve long-term survival with the development of TVP.
This model can also be used to investigate acute cellular rejection and media necrosis in xenogeneic models (5).

This video demonstrates the orthotopic aortic transplant model as a simple model to study the development of transplant vasculopathy (TVP) in rats.

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant ...

A New Murine Model of Endovascular Aortic Aneurysm Repair

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of Abdominal aortic aneurysm (AAA) pathophysiology. There is no abdominal aortic aneurysm endograft exclusion model cheap and reproducible, which would allow deep investigations of AAA before and after treatment. We hereby describe how to induce, and then to exclude with a covered coronary stentgraft an abdominal aortic aneurysm in a rat. The well known elastase induced AAA model was first reported in 19901 in a rat, then described in mice2. Elastin degradation leads to dilation of the aorta with inflammatory infiltration of the abdominal wall and intra luminal thrombus, matching with human AAA. Endovascular exclusion with small covered stentgraft is then performed, excluding any interactions between circulating blood and the aneurysm thrombus. Appropriate exclusion and stentgraft patency is confirmed before euthanasia by an angiography thought the left carotid artery. Partial control of elastase diffusion makes aneurysm shape different for each animal. It is difficult to create an aneurysm, which will allow an appropriate length of aorta below the aneurysm for an easy stentgraft introduction, and with adequate proximal and distal neck to prevent endoleaks. Lots of failure can result to stentgraft introduction which sometimes lead to aorta tear with pain and troubles to stitch it, and endothelial damage with post op aorta thrombosis. Giving aspirin to rats before stentgraft implantation decreases failure rate without major hemorrhage. Clamping time activates neutrophils, endothelium and platelets, and may interfere with biological analysis.

Histological and biochemical modifications after aneurysm endograft exclusion are still unclear. We describe a new model of endograft implantation on a murine aneurysm. Through thrombus analysis with and without persistant circulating blood stream interface, and new endovascular biomaterials evaluation, this model will help to better understand endovascular aneurysm exclusion pathobiology.

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of ...

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are available to perform mechanistic in vivo studies. While heart transplantation models using a suture technique were first successfully developed in rats, the translation into an equally widespread used murine equivalent was never achieved due the technical complexity of the microsurgical procedure. In contrast, non-suture cuff techniques, also developed initially in rats, were successfully adapted for use in mice1-3. This technique for revascularization involves two major steps I) everting the recipient vessel over a polyethylene cuff; II) pulling the donor vessel over the formerly everted recipient vessel and holding it in place with a circumferential tie. This ensures a continuity of the endothelial layer, short operating time and very high patency rates4.
Using this technique for vascular anastomosis we performed more than 1,000 cervical heart transplants with an overall success rate of 95%. For arterial inflow the common carotid artery and the proximal aortic arch were anastomosed resulting in a retrograde perfusion of the transplanted heart. For venous drainage the pulmonary artery of the graft was anastomosed with the external jugular vein of the recipient5.
Herein, we provide additional details of this technique to supplement the video.

The murine cervical heart transplantation model is well suited for immunological as well as ischemia reperfusion injury studies. We modified the procedure using a non-suture cuff technique and performed more than 1,000 successful transplants with this approach.
Herein, we provide additional details of this technique to supplement the video.

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are ...

A Murine Model of Cervical Spinal Cord Injury to Study Post-lesional Respiratory Neuroplasticity

A cervical spinal cord injury induces permanent paralysis, and often leads to respiratory distress. To date, no efficient therapeutics have been developed to improve/ameliorate the respiratory failure following high cervical spinal cord injury (SCI). Here we propose a murine pre-clinical model of high SCI at the cervical 2 (C2) metameric level to study diverse post-lesional respiratory neuroplasticity. The technique consists of a surgical partial injury at the C2 level, which will induce a hemiparalysis of the diaphragm due to a deafferentation of the phrenic motoneurons from the respiratory centers located in the brainstem. The contralateral side of the injury remains intact and allows the animal recovery. Unlike other SCIs which affect the locomotor function (at the thoracic and lumbar level), the respiratory function does not require animal motivation and the quantification of the deficit/recovery can be easily performed (diaphragm and phrenic nerve recordings, whole body ventilation). This pre-clinical C2 SCI model is a powerful, useful, and reliable pre-clinical model to study various respiratory and non-respiratory neuroplasticity events at different levels (molecular to physiology) and to test diverse putative therapeutic strategies which might improve the respiration in SCI patients.

Respiratory failure is the leading cause of death following a cervical spinal cord injury. Having a reproducible, quantifiable, and reliable pre-clinical animal model of respiratory failure induced by a partial cervical injury will help to understand the subsequent respiratory and non-respiratory neuroplasticity and allow testing putative repair strategies.

A cervical spinal cord injury induces permanent paralysis, and often leads to respiratory distress. To date, no efficient therapeutics have been developed ...

Murine Heterotopic Heart Transplant Technique

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become proficient in and utilize this technique, it is now widely used by many laboratories around the world. A significant refinement to the original technique was developed and reported in 2001 by Niimi. Described here are the techniques that have evolved over more than a decade in the hands of three surgeons (Plenter, Grazia, Pietra) in our center. These techniques are now being passed on to a younger generation of surgeons and researchers.
Based largely on the Niimi experience, the procedures used have evolved in the fine details - details which we will endeavor to relate here in such a way that others may be able to use this very useful model. Like Niimi, we have found that a video aid to learning is a priceless resource for the beginner.

The manuscript describes the steps required to perform the heterotopic heart transplant in the mouse.

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become ...

Human Skeletal Muscle Biopsy Procedures Using the Modified Bergstr&#246;m Technique

The percutaneous biopsy technique enables researchers and clinicians to collect skeletal muscle tissue samples. The technique is safe and highly effective. This video describes the percutaneous biopsy technique using a modified Bergstr&#246;m needle to obtain skeletal muscle tissue samples from the vastus lateralis of human subjects. The Bergstr&#246;m needle consists of an outer cannula with a small opening (&#8216;window&#8217;) at the side of the tip and an inner trocar with a cutting blade at the distal end. Under local anesthesia and aseptic conditions, the needle is advanced into the skeletal muscle through an incision in the skin, subcutaneous tissue, and fascia. Next, suction is applied to the inner trocar, the outer trocar is pulled back, skeletal muscle tissue is drawn into the window of the outer cannula by the suction, and the inner trocar is rapidly closed, thus cutting or clipping the skeletal muscle tissue sample. The needle is rotated 90&#176;&#160;and another cut is made. This process may be repeated three more times. This multiple cutting technique typically produces a sample of 100-200 mg or more in healthy subjects and can be done immediately before, during, and after a bout of exercise or other intervention. Following post-biopsy dressing of the incision site, subjects typically resume their activities of daily living right away and can fully participate in vigorous physical activity within 48-72 hr. Subjects should avoid heavy resistance exercise for 48 hr to reduce the risk of herniation of the muscle through the incision in the fascia.

Human Skeletal Muscle Biopsy Procedures Using the Modified Bergstr&amp;#246;m Technique

The purpose of this video is to present the modified Bergstr&#246;m skeletal muscle biopsy technique on human subjects.

The percutaneous biopsy technique enables researchers and clinicians to collect skeletal muscle tissue samples. The technique is safe and highly ...

Murine Corneal Transplantation: A Model to Study the Most Common Form of Solid Organ Transplantation

Corneal transplantation is the most common form of organ transplantation in the United States with between 45,000 and 55,000 procedures performed each year. While several animal models exist for this procedure and mice are the species that is most commonly used. The reasons for using mice are the relative cost of using this species, the existence of many genetically defined strains that allow for the study of immune responses, and the existence of an extensive array of reagents that can be used to further define responses in this species. This model has been used to define factors in the cornea that are responsible for the relative immune privilege status of this tissue that enables corneal allografts to survive acute rejection in the absence of immunosuppressive therapy. It has also been used to define those factors that are most important in rejection of such allografts. Consequently, much of what we know concerning mechanisms of both corneal allograft acceptance and rejection are due to studies using a murine model of corneal transplantation. In addition to describing a model for acute corneal allograft rejection, we also present for the first time a model of late-term corneal allograft rejection.

Mice have been used as a model for studying many forms of transplantation, including corneal transplantation. We describe in this report a murine model for both acute and late-term corneal transplantation.

Corneal transplantation is the most common form of organ transplantation in the United States with between 45,000 and 55,000 procedures performed each ...

Murine Kidney Transplant Technique

The first mouse kidney transplant technique was published in 19731 by the Russell laboratory. Although it took some years for other labs to become proficient in and utilize this technique, it is now widely used by many laboratories around the world. A significant refinement to the original technique using the donor aorta to form the arterial anastomosis instead of the renal artery was developed and reported in 1993 by Kalina and Mottram 2 with a further advancement coming from the same laboratory in 1999 3. While one can become proficient in this model, a search of the literature reveals that many labs still experience a high proportion of graft loss due to arterial thrombosis. We describe here a technique that was devised in our laboratory that vastly reduces the arterial thrombus reported by others 4,5. This is achieved by forming a heel-and-toe cuff of the donor infra-renal aorta that facilitates a larger anastomosis and straighter blood flow into the kidney.

The goal of this manuscript is to describe the steps required to perform a kidney transplant in a mouse, paying particular attention to the details of the arterial anastomosis.

The first mouse kidney transplant technique was published in 19731 by the Russell laboratory. Although it took some years for other labs to become ...

Heterotopic Cervical Heart Transplantation in Mice

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies this model by avoiding challenging suture anastomoses of small vessels thereby reducing warm ischemia time. In comparison to abdominal graft implantation the cervical model is less invasive and the implanted graft is easily accessible for further follow-up examinations. Anastomoses are performed by pulling the ascending aorta of the graft over the cuff with the recipient&#8217;s common carotid artery and by pulling the main pulmonary artery over the cuff with the external jugular vein. Selection of appropriate cuff size and complete mobilization of the vessels are important for successful revascularization. Ischemia-reperfusion (I/R) injury can be minimized by perfusing the graft with a cardioplegic solution and by hypothermia. In this article, we provide technical details for a simplified and improved cuff technique, which should allow surgeons with basic microsurgical skills to perform the procedure with a high success rate.

The cuff technique shortens and facilitates the model of heterotopic cervical heart transplantation in mice by avoiding technically challenging suture anastomoses of small vessels. The technical details shown in this video paper should allow researches to establish this model in their laboratories.

The heterotopic cervical heart transplantation in mice is a valuable tool in transplant and cardiovascular research. The cuff technique greatly simplifies ...

Using the Sleeve Technique in a Mouse Model of Aortic Transplantation - An Instructional Video

Orthotopic aortic transplantation using the sleeve technique reduces injury to the aorta with failure rate of only 10-20%. The time to anastomose the aorta in mice using the sleeve method was short and easy averaging 20 min, permitting studies of iso/allo grafts. The following article describes the aortic transplantation procedure used in our laboratory. The mice were anesthetized with a mixture of 1.5% volume isoflurane and 100% oxygen through a face mask. At this point, the segment of the aorta between the renal arteries and its bifurcation was separated from the vena cava, freely prepared and clampedat the proximal and distal segments with a single silk suture. Prior to the removal of the aorta, a saline solution containing heparin was injected into the inferior vena cava. Then the aorta was cut between the clamps and a saline heparin solution was used to flush the lumen. The sleeve technique with monofilament sutures was used in order to transplant the abdominal aorta in the orthotopic position.

We present an orthotopic aortic transplantation model using the sleeve technique in mice. It is a very rapid anastomosis method, which can be employed in studies of vascular disease.

Orthotopic aortic transplantation using the sleeve technique reduces injury to the aorta with failure rate of only 10-20%. The time to anastomose the ...