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>

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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. 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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&amp;amp;T Forceps - SuperGrip Tips (Angled 45&amp;deg;)</td><td>S&amp;amp;T</td><td>00649-11</td><td>Fine Forceps</td></tr><tr><td>S&amp;amp;T Vessel Dilating Forceps - Angled 10&amp;deg; (Tip diameter 0.2 mm)</td><td>S&amp;amp;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&amp;uuml;bingen Spring Scissors (curved, 8.5 cm)</td><td>FST</td><td>15004-08</td><td>Microsissors (curved)</td></tr><tr><td>Vannas-T&amp;uuml;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. ...

murine-cervical-aortic-transplantation-model-using-modified-non

Research

JoVE Journal

Medicine

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

Generation and Grafting of Tissue-engineered Vessels in a Mouse Model

The construction of vascular conduits is a fundamental strategy for surgical repair of damaged and injured vessels resulting from cardiovascular diseases. The current protocol presents an efficient and reproducible strategy in which functional tissue engineered vessel grafts can be generated using partially induced pluripotent stem cell (PiPSC) from human fibroblasts. We designed a decellularized vessel scaffold bioreactor, which closely mimics the matrix protein structure and blood flow that exists within a native vessel, for seeding of PiPSC-endothelial cells or smooth muscle cells prior to grafting into mice. This approach was demonstrated to be advantageous because immune-deficient mice engrafted with the PiPSC-derived grafts presented with markedly increased survival rate 3 weeks after surgery. This protocol represents a valuable tool for regenerative medicine, tissue engineering and potentially patient-specific cell-therapy in the near future.

Here, we present a protocol to generate tissue engineered vessel grafts that are functional for grafting into mice by double seeding partially induced pluripotent stem cell (PiPSC) - derived smooth muscle cells and PiPSC - derived endothelial cells on a decellularized vessel scaffold bioreactor.

The construction of vascular conduits is a fundamental strategy for surgical repair of damaged and injured vessels resulting from cardiovascular diseases. ...

Bioengineering

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 ...

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 ...

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 ...

Optimization of the Cuff Technique for Murine Heart Transplantation

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to improve surgical efficiency. In our lab, we have optimized the cuff technique with two major steps. First, we used the inner tube technique to insert a temporary inner tube into the external jugular vein and carotid artery blood vessel to facilitate eversion of the vessel over the cuff. Second, we performed complete heterotopic cardiac transplantation through the collaboration of two experienced surgeons. These modifications effectively reduced the operation time to 25 minutes, with a success rate of 95%. In this report, we describe these procedures in detail and provide a supplemental video. We believe that this report on the improved cuff technique will offer practical guidance for murine heterotopic heart transplantation and will enhance the utility of this mouse model for basic research.

We introduce an inner tube approach to the cuff technique for mouse cervical heterotopic heart transplantation to help evert the vessel over the cuff. We found that cooperation between two experienced surgeons markedly shortens the operation time.

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to ...

Immunology and Infection

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after transplantation, and the impact of immunomodulatory therapies on graft rejection. Heterotopic cervical heart transplantation in mice was first described in 1991 using sutured anastomoses and subsequently modified to include cuff techniques. This modification allowed for improved success rates, and since then, there have been multiple reports that have proposed further technical improvements. However, translation into more widespread utilization remains limited due to the technical difficulty associated with graft anastomoses, which requires precision to achieve adequate length and caliber of the cuffs to avoid vascular anastomotic twisting or excessive tension, which can result in damage to the graft. The present protocol describes a modified technique for performing heterotopic cervical cardiac transplantation in mice which involves cuff placement on the recipient's common carotid artery and the donor's pulmonary artery in alignment with the direction of the blood flow.

Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after ...

Implantation of Tissue-Engineered Vascular Graft in Mouse Carotid Artery via Cuff Technique

The development of small-diameter vascular grafts has been a global endeavor, with numerous research groups contributing to this field. Animal experimentation plays a pivotal role in assessing the efficacy and safety of vascular grafts, particularly in the absence of clinical applications. Compared to alternative animal models, the mouse implantation model offers several advantages, including a well-defined genetic background, a mature method for disease model construction, and a straightforward surgical procedure. Based on these advantages, the present study devised a simple cuff technique for the implantation of tissue-engineered vascular grafts in the mouse carotid artery. This technique began with the fabrication of polycaprolactone (PCL) small-diameter vascular grafts via electrostatic spinning, followed by the seeding of macrophages onto the grafts through perfusion adsorption. Subsequently,&#160;the cellularized tissue-engineered vascular grafts were transplanted into the mouse carotid artery using the cuff technique to evaluate patency and regenerative capability. After 30 days of in vivo implantation, vascular patency was found to be satisfactory, with evidence of neo-tissue regeneration and the formation of an endothelial layer within the lumen of the grafts. All data were analyzed using statistical and graphing software. This study successfully established a mouse carotid artery implantation model that can be used to explore the cellular sources of vascular regeneration and the mechanisms of action of active substances. Furthermore, it provides theoretical support for the development of novel small-diameter vascular grafts.

Here, a protocol is presented for the implantation of a tissue-engineered vascular graft into the mouse carotid artery using the cuff technique, providing a suitable animal model for investigating vascular tissue regeneration mechanisms.

The development of small-diameter vascular grafts has been a global endeavor, with numerous research groups contributing to this field. Animal ...

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for investigating the mechanism of cardiac allograft rejection in preclinical studies because of their high homology with human genes. This understanding would help develop unique approaches to improving patients&#39; long-term survival treated with cardiac allografts. In a mouse model, abdominal donor heart implantation is commonly performed with an end-to-end&#160;anastomosis to the recipient&#39;s aorta and inferior vena cava using stitches. In this model, the donor&#39;s heart is implanted by end-to-end anastomosis to the recipient&#39;s carotid artery and jugular vein by the modified-Cuff technique. The transplantation surgery is performed without stitching and thus may increase the survival of the recipient since there is no interference with the blood supply and venous reflux of the lower body. This mouse model would help investigate the mechanisms underlying the immunological and pathological (acute/chronic) rejection of cardiac allografts.

In the present protocol, a mouse heart transplantation model is used for investigating the mechanism of cardiac allograft rejection. In this heterotopic heart transplantation model, operation efficiency is improved, and the survival of cardiac grafts is ensured by a cervical end-to-end anastomosis of heart implantation using a modified Cuff technique.

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for ...

A Simple Cuff Technique for Murine Left Lung Transplantation

Over the past decade, our laboratory has made significant progress in developing and refining vascularized mouse lung transplantation models using an efficient and highly reliable "cuff technique" of transplantation. This article describes a sophisticated and comprehensive method for orthotopic lung transplantation in a vascularized orthotopic lung model, representing the most physiologic and clinically relevant model of mouse lung transplantation to date. The transplantation process consists of two distinct stages: donor harvest and subsequent implantation into the recipient. The method has been successfully mastered, and with several months of sufficient training, a skilled practitioner can perform the procedure in approximately 90 min from skin-to-skin. Surprisingly, once individuals overcome the initial learning curve, the survival rate during the perioperative period approaches nearly 100%. The mouse model allows for the use of multiple commercially available transgenic and mutant strains of mice, enabling the study of tolerance and rejection. Additionally, the unique features of this model make it a valuable tool for investigating tumor biology and immunology.

The present protocol describes a cuff technique for a mouse left lung transplantation model. This technique has been developed over several years and has performed well, serving effectively in immunological research.

Over the past decade, our laboratory has made significant progress in developing and refining vascularized mouse lung transplantation models using an ...