The study was funded by the integrated budget for interdepartmental research (BIRD) 2019.

All procedures have been approved by the University of Padova Animal Care Committee (OPBA, protocol number n&#176; 55/2017) and authorized by the Italian Ministry of Health (Authorization n&#176; 700/2018-PR), in compliance with the European Union Directive 2010/63/UE and the Italian Law 26/2014 for the Care and Use of Laboratory Animals.
1. Animal care and experimental model
<ol>
	<li>Ensure all Lewis rats are obtained from a single company (Table of materials). Maintain the rats in conventional facilities with free access to food and water.</li>
	<li>Ensure that the weight of the rats ranges from 320-400 g for the recipient group and 200-250 g for the donor group.</li>
</ol>
2. Preoperative protocol
NOTE: All operations must be performed under clean conditions. Use male and female adult Lewis rats as recipients and donors as well in order to perform a syngeneic transplant.
<ol>
	<li>Perform an intraperitoneal injection of tramadol (5 mg/kg) 15 min before surgery.</li>
	<li>Administer a single dose of intramuscular Gentamicin (5 mg/kg) immediately before surgery.</li>
	<li>For anesthesia induction, supply 4% sevoflurane in 1 L/min of oxygen to a poly(methyl methacrylate) chamber where the animal is placed. For anesthesia maintenance, use 2.0-2.5% sevoflurane in 1 L/min of oxygen throughout the procedure.</li>
	<li>Shave the animal along the midline for 2 cm width from the sternum to 1 cm above the genital area with a razor. Then, sterilize the skin with iodine solution.</li>
	<li>In order to prevent the animal from getting wet and to prevent heat dispersion during the surgery, cover the animal with a transparent plastic film.</li>
	<li>Evaluate the level of anesthesia before performing the procedure by assessing the absence of response to a noxious stimulus.</li>
</ol>
3. Donor operation
<ol>
	<li>Animal and heart preparation:
	<ol>
		<li>Place the anesthetized animal on a cork tray with the caudal side facing the surgeon. Perform a xipho-pubic incision of about 5-6 cm, and retract the two musculocutaneous flaps laterally.</li>
		<li>Administer a volume of 1 mL of saline solution at 4 &#176;C containing 500 IU of heparin through the abdominal vena cava.</li>
		<li>After 1 min, cut the diaphragm from left to right and perform an anterior thoracotomy to expose the heart.</li>
		<li>Cool the beating heart by dripping saline solution at 4 &#176;C.</li>
		<li>Perform a pericardiectomy and a thymectomy in order to obtain a complete view of the aortic arch. Remove the remaining fatty tissues surrounding the aorta.</li>
		<li>Cut at the arch, just above the origin of the innominate artery; sever the latter one, too.</li>
		<li>Cut the thoracic inferior vena cava (IVC) and insert a 22 G cannula to infuse the heart with 20-25 mL of saline solution at 4 &#176;C, exerting light pressure. Discontinue the perfusion when the heart stops beating and flow from the aorta became clear.</li>
	</ol>
	</li>
	<li>PAG explant: 
	NOTE: An accurate harvesting and delicate handling of the PAG is mandatory for achieving optimal implantation in the recipient. Do not touch it directly with instruments, instead use cotton swabs.
	<ol>
		<li>Perform an ultrasound study to assess the PA diameter in its native position.</li>
		<li>Insert a micro-plier under the posterior wall of the vessel and cut the latter using a micro-scissor as close as possible to its bifurcation to maximize the length of the PAG.</li>
		<li>Gently hold the PA with the ring-tipped micro-forceps and separate it from the right ventricle with the micro-spring scissors. Harvest the PAG, including some right ventricular muscle.</li>
	</ol>
	</li>
	<li>PAG preparation:
	<ol>
		<li>Place the PAG on a gauze moistened with cold saline on the operating table and inspect the vessel under the operating microscope.</li>
		<li>Cut any abundant surrounding tissue, leaving only 1 mm of ventricular muscle. Set the length of the vessel at 5 mm.</li>
	</ol>
	</li>
</ol>
4. Pulmonary artery graft (PAG) implantation
<ol>
	<li>Preparation of the recipient animal:
	<ol>
		<li>Place the anesthetized animal on a cork tray with the caudal side facing the surgeon.</li>
		<li>Perform a median longitudinal incision and use two mini retractors to keep the abdomen open.</li>
		<li>Extract the intestines with two cotton swabs and cover them with a gauze soaked with 39 &#176;C saline allowing the visualization of the retroperitoneal area with exposure of the infra-renal abdominal aorta (AA). 
		NOTE: During the surgery, it is important to occasionally moisten the intestines using a syringe containing 39 &#176;C saline to prevent hypothermia, a critical condition common in rodents.</li>
		<li>Strip the posterior parietal peritoneum between the two renal arteries and the iliac bifurcation using two cotton swabs and remove the fatty tissue around the infrarenal AA. Leave only a small portion of fat above the AA, to facilitate handling on the vessel.</li>
		<li>Separate the AA from the IVC. To perform this procedure, first, pass a curved forceps behind the posterior aortic wall and use it to open a passage between the AA and IVC. Then, use a 2-0 silk suture to create a loop around the AA, in order to lift the vessel and separate the AA from IVC. Ligate any lumbar artery arising from the infrarenal AA with 6/0 silk suture and divide it.</li>
		<li>Rotate the animal 90&#176; counterclockwise, placing the head on the operator&#39;s left side. The AA now lay horizontally in the microscopic field.</li>
		<li>Use two Yasargil clips to clamp the infrarenal AA and place them at a distance of 1.5 cm from each other. Transect the AA at the midpoint between the two clips.</li>
		<li>Irrigate the two ends of the vessels with heparin (1 UI/mL) in saline solution to remove any clots. Remove any adventitial debris from the vessels.</li>
	</ol>
	</li>
	<li>PAG implantation:
	<ol>
		<li>Place the PAG between the two ends, with the ventricular end towards the cranial portion of the animal.</li>
		<li>Use a 10-0 polypropylene suture to perform two landmark single stitches connecting the PG to the AA. Perform the procedure on both ends of the PAG by placing the suture on opposite sides of the vessel circumference.</li>
		<li>Perform an end-to-end anastomosis between PAG and AA, beginning with the distal end. Use one of the two ends of the distal landmark suture for the posterior anastomosis using a recipient-to-graft out-in/in-out sequence to perform a running suture of about six stitches.</li>
		<li>Once the suture reaches the proximal landmark, perform a double half hitch completed by a square knot using the suture and one of the two ends of the proximal landmark suture. Apply rubber-shod mosquito forceps to the sutures to provide traction.</li>
		<li>Perform the same anastomosis on the anterior wall. Carry on the entire procedure on the proximal end of the PAG. Take particular attention when performing the proximal anastomosis to avoid including any leaflet in the suture line.</li>
		<li>Release the distal clip first to let the PAG be filled with retrograde blood (low-pressure flow) in order to check the anastomosis. Repair any blood leakage with a single suture. Once the distal anastomosis is evaluated, perform the same procedure on the proximal end.</li>
	</ol>
	</li>
	<li>Final stages of the operation on the recipient:
	<ol>
		<li>Assess the patency of the PAG and apply two strips of gelatin sponge over the suture lines on both sides of the PAG (if necessary). Exert gentle pressure for a few seconds with two cotton swabs to help hemostasis.</li>
		<li>Relocate the intestines into the abdominal cavity and close the walls with a 4/0 polypropylene running suture.</li>
	</ol>
	</li>
</ol>
5. Sham-operated procedure
<ol>
	<li>Perform an identical preparation of the animal as previously illustrated for recipient rats.</li>
	<li>Cut the infra-renal AA, midway between the renal and the iliac arteries&#39; origin.</li>
	<li>Reapproximate the two ends of the AA using an end-to-end anastomosis, as previously described. Remove the two clips and perform an accurate hemostasis procedure.</li>
	<li>Reposition the intestines and close the abdominal wall in layers, as for the recipient animals.</li>
</ol>
6. Postoperative care and follow-up
<ol>
	<li>Administer warm saline solution (5 mL) into the subcutaneous tissue of the animal&#39;s back for hydration. Place the rat under a heating lamp and visually monitor it until awakening, which usually takes up to 5 min after anesthesia is stopped. Place the animal in a cage at a room temperature of 22-24 &#176;C, with immediate and unrestricted access to food and water.</li>
	<li>Administer intramuscular tramadol (5 mg/kg) for postoperative analgesia twice daily for the first 48 h after surgery. Thereafter, monitor the recipient&#39;s health status and body weight daily, on a regular basis.</li>
	<li>Follow-up: During the follow-up, perform seriate ultrasound studies at one week, one month, and two months to evaluate PAG function. During these studies, measure the vessel diameter, the peak systolic velocity (PSV), and the end-diastolic velocity. Measure these parameters inside the PAG and at the level of proximal and distal AA.</li>
	<li>Euthanize the animals after two months of follow-up by application of CO2 for a few minutes, and then explant the PAG, which will undergo histopathological analysis.</li>
</ol>

<ol>
	<li>Botto, L. D., Correa, A., Erickson, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Racial+and+temporal+variations+in+the+prevalence+of+heart+defects.">Racial and temporal variations in the prevalence of heart defects.</a> Pediatrics. 107 (3), 32 (2001).</li><li>Vergnat, 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=Aortic+stenosis+of+the+neonate:+A+single-center+experience.">Aortic stenosis of the neonate: A single-center experience.</a> The Journal of Thoracic and Cardiovascular Surgery. 157 (1), 318-326 (2019).</li><li>Hra&#353;ka, V., 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=The+long-term+outcome+of+open+valvotomy+for+critical+aortic+stenosis+in+neonates.">The long-term outcome of open valvotomy for critical aortic stenosis in neonates.</a> The Annals of Thoracic Surgery. 94 (5), 1519-1526 (2012).</li><li>Kaza, A. K., Pigula, F. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Are+bioprosthetic+valves+appropriate+for+aortic+valve+replacement+in+young+patients.">Are bioprosthetic valves appropriate for aortic valve replacement in young patients.</a> Seminars in Thoracic and Cardiovascular Surgery: Pediatric Cardiac Surgery Annual. 19 (1), 63-67 (2016).</li><li>Myers, P. O., 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=Outcomes+after+mechanical+aortic+valve+replacement+in+children+and+young+adults+with+congenital+heart+disease.">Outcomes after mechanical aortic valve replacement in children and young adults with congenital heart disease.</a> The Journal of Thoracic and Cardiovascular Surgery. 157 (1), 329-340 (2019).</li><li>Donald, J. S., 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=Ross+operation+in+children:+23-year+experience+from+a+single+institution.">Ross operation in children: 23-year experience from a single institution.</a> The Annals of thoracic surgery. 109 (4), 1251-1259 (2020).</li><li>Khwaja, S., Nigro, J. J., Starnes, V. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Ross+procedure+is+an+ideal+aortic+valve+replacement+operation+for+the+teen+patient.">The Ross procedure is an ideal aortic valve replacement operation for the teen patient.</a> Seminars in Thoracic and Cardiovascular Surgery: Pediatric Cardiac Surgery Annual. , 173-175 (2005).</li><li>Elkins, R. C., Lane, M. M., McCue, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ross+operation+in+children:+late+results.">Ross operation in children: late results.</a> The Journal of Heart Valve Disease. 10 (6), 736-741 (2001).</li><li>Chambers, J. C., Somerville, J., Stone, S., Ross, D. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pulmonary+autograft+procedure+for+aortic+valve+disease:+long-term+results+of+the+pioneer+series.">Pulmonary autograft procedure for aortic valve disease: long-term results of the pioneer series.</a> Circulation. 96 (7), 2206-2214 (1997).</li><li>Mazine, A., 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=Ross+procedure+in+adults+for+cardiologists+and+cardiac+surgeons:+JACC+state-of-the-art+review.">Ross procedure in adults for cardiologists and cardiac surgeons: JACC state-of-the-art review.</a> Journal of the American College of Cardiology. 72 (22), 2761-2777 (2018).</li><li>Sengupta, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+laboratory+rat:+Relating+its+age+with+humans.">The laboratory rat: Relating its age with humans.</a> International Journal of Preventive Medicine. 4 (6), 624-630 (2013).</li><li>Ashfaq, A., Leeds, H., Shen, I., Muralidaran, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reinforced+ross+operation+and+intermediate+to+long+term+follow+up.">Reinforced ross operation and intermediate to long term follow up.</a> Journal of Thoracic Disease. 12 (3), 1219-1223 (2020).</li><li>Vida, V. 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=Age+is+a+risk+factor+for+maladaptive+changes+of+the+pulmonary+root+in+rats+exposed+to+increased+pressure+loading.">Age is a risk factor for maladaptive changes of the pulmonary root in rats exposed to increased pressure loading.</a> Cardiovascular Pathology: The Official Journal of the Society for Cardiovascular Pathology. 21 (3), 199-205 (2012).</li><li>Nappi, F., 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=An+experimental+model+of+the+Ross+operation:+Development+of+resorbable+reinforcements+for+pulmonary+autografts.">An experimental model of the Ross operation: Development of resorbable reinforcements for pulmonary autografts.</a> The Journal of Thoracic and Cardiovascular Surgery. 149 (4), 1134-1142 (2015).</li><li>Vanderveken, E., 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=Mechano-biological+adaptation+of+the+pulmonary+artery+exposed+to+systemic+conditions.">Mechano-biological adaptation of the pulmonary artery exposed to systemic conditions.</a> Scientific Reports. 10 (1), 2724 (2020).</li></ol>

The authors have nothing to disclose.

Aortic valve replacement with the autologous pulmonary root (Ross operation) represents an attractive option for congenital aortic valve stenosis repair due to the favorable profile and potential growth of the autograft10. The major limitation to this procedure is the potential dilatation of the aortic neo-valve, which predisposes to the development of long-term regurgitation. The possibility of characterizing the modifications on the pulmonary artery after exposure to systemic pressures could rep...

Congenital aortic valve stenosis is a subgroup of congenital heart disease characterized by an obstruction of the left ventricular tract in which the lesion is located at the valvular level. The malformation affects approximately 0.04-0.38 per 1000 live births1.
The available options for the correction are many, each with its own advantages and disadvantages. For patients suitable for a biventricular correction2, the approach may be aimed at valve repair (percutaneous or surgical valvulotomy) or its replacement3. The latter is preferred when the aortic valve is considered unsalvageable; however, the available options are limited for pediatric patients. Indeed, bioprosthetic valves are not indicated for aortic replacement in the young population due to their early calcification4. On the other hand, degeneration in mechanical valves is considerably slower, but these require lifelong anticoagulant therapy5. In addition, the major limitation of these prostheses is represented by the lack of growth potential, which predisposes the patients to additional reinterventions.
An interesting therapeutic option in the pediatric population is the transfer of the pulmonary autograft to the aortic position named &#34;Ross operation&#34;. In this case, the pulmonary valve is then replaced with a homograft (Figure 1)6. This procedure can possibly represent the best surgical choice for children because the pulmonary autograft preserves its growth potential and does not carry the risks of lifelong anticoagulant therapy. Furthermore, the Ross procedure can be of great value also in young adults to avoid a mechanical or biological valve, having the potential to become the best surgical solution.
Results after aortic valve replacement with pulmonary autograft are excellent, with survival greater than 98% and good long-term outcomes7. Literature studies report 93% and 90% freedom from replacing the pulmonary homograft at 4 and 12 years, respectively8.
The major limitation of this procedure is the tendency of the autograft to dilate in the long term, especially when employed as a freestanding root replacement. This can cause valvular incompetence which may require a reintervention. Indeed, the longest follow-up study performed so far reports freedom from reoperation for autograft replacement of 88% at 10 years and 75% at 20 years9.
The possibility of recreating a Ross operation in an experimental setting represents a fundamental prerequisite to investigate the underlying mechanism of the pulmonary autograft adaption to systemic pressures. Several models have been proposed in the past. However, these are usually limited to ex-vivo experiments or in-vivo animal models with a relatively expensive large animals. In this study, we sought to establish a rodent model of pulmonary artery graft (PAG) implantation in a systemic position, as freestanding root.

<table><tbody><tr><td>0.9% Sodium Chloride</td><td>Monico SpA</td><td>AIC 030805105</td><td>Two bottles of 100 mL. The cold one (4&amp;deg;C) for flushing the harvesting organ; the warm one (39&amp;deg;C) for moistening, and rehydration of the recipient</td></tr><tr><td>7.5% Povidone-Iodine</td><td>B Braun</td><td>AIC 032151211</td><td></td></tr><tr><td>Barraquer</td><td>Aesculap</td><td>FD 232R</td><td>Straight micro needle holder for the vascular anastomoses</td></tr><tr><td>Castroviejo needle holder</td><td>Not available</td><td>J 4065</td><td>To close the animal</td></tr><tr><td>Clip applying forceps</td><td>Rudolf Medical</td><td>RU 3994-05</td><td>For clip application</td></tr><tr><td>Cotton swabs</td><td>Johnson &amp;amp; Johnson Medical SpA</td><td>N/A</td><td>Supermarket product. Sterilized</td></tr><tr><td>Curved micro jeweller forceps</td><td>Rudolf Medical</td><td>RU 4240-06</td><td>Used to pass sutures underneath the vases.</td></tr><tr><td>Depilatory cream</td><td>RB healthcare</td><td>N/A</td><td>Supermarket product</td></tr><tr><td>Electrocautery machine</td><td>LED SpA</td><td>Surton 200</td><td></td></tr><tr><td>Fine scissors</td><td>Rudolf Medical</td><td>RU 2422-11</td><td>For opening the abdomen (recipient)</td></tr><tr><td>Fine-tip curved Vannas micro scissors</td><td>Aesculap</td><td>OC 497R</td><td>Only for preparing the pulmonary root, cut the lumbar vases and the 10/0 Prolene</td></tr><tr><td>Fluovac Isoflurane/Halotane Scavanger unit</td><td>Harvard Apparatus Ltd</td><td>K 017041</td><td>Complete of anesthesia machine, anesthesia tubing, induction chamber and scavenger unit with absorbable filter</td></tr><tr><td>Gentamycin</td><td>MSD Italia Srl</td><td>AIC 020891014</td><td>Antibiotic. Single dose, 5 mg/kg intramuscular, administered during surgery</td></tr><tr><td>Heparin</td><td>Pharmatex Italia Srl</td><td>AIC 034692044</td><td>500 IU into the recipient abdominal vena cava</td></tr><tr><td>I.V. Catheter</td><td>Smiths Medical Ltd</td><td>4036</td><td>20G</td></tr><tr><td>Insulin Syringe, 1 mL</td><td>Fisher Scientific</td><td>14-841-33</td><td>To inject heparin in the harvesting animal and to flush the sectioned aorta in the recipient</td></tr><tr><td>Jeweler bipolar forceps</td><td>GIMA SpA</td><td>30665</td><td>0.25 mm tip. For electrocautery of very small vases</td></tr><tr><td>Lewis rats (LEW/HanHsd)</td><td>Envigo RMS SRL, San Pietro al Natisone, Udine, Italy</td><td>86104M</td><td>Male or female, weighing 200-250 g (pulmonary root harvesting animals) and 320-400 g (recipients)</td></tr><tr><td>Micro-Mosquito</td><td>Rudolf Medical</td><td>RU 3121-10</td><td>In number of four, with tips covered with silicon tubing. To keep in traction the Prolene suture during anastomosis</td></tr><tr><td>Operating microscope</td><td>Leica Microsystems</td><td>M 400-E</td><td>Used with 6x, 10x and 16x in-procedure interchangeable magnifications</td></tr><tr><td>Perma-Hand silk 2-0</td><td>Johnson &amp;amp; Johnson Medical SpA</td><td>C026D</td><td>To lift the aorta</td></tr><tr><td>Petrolatum ophthalmic ointment</td><td>Dechra</td><td>NDC 17033-211-38</td><td></td></tr><tr><td>Prolene 10-0</td><td>Johnson &amp;amp; Johnson Medical SpA</td><td>W2790</td><td>Very fine non-absorbable suture, with a BV75-3 round bodied needle, for the vascular anastomoses</td></tr><tr><td>Retractors</td><td>Not any</td><td>N/A</td><td>Two home-made retractors</td></tr><tr><td>Ring tip micro forceps</td><td>Rudolf Medical</td><td>RU 4079-14</td><td>For delicate manipulation</td></tr><tr><td>Sevoflurane</td><td>AbbVie Srl</td><td>AIC 031841036</td><td>Mixed with oxygen, for inhalatory anesthesia</td></tr><tr><td>Spring type micro scissors</td><td>Rudolf Medical</td><td>RU 2380-14</td><td>Straight; 14 cm long</td></tr><tr><td>Standard aneurysm clips</td><td>Rudolf Medical</td><td>RU 3980-12</td><td>Two clips (7.5 mm; 180 g; 1.77 N) to close the aorta</td></tr><tr><td>Sterile gauze of non-woven fabric material</td><td>Luigi Salvadori SpA</td><td>26161V</td><td>7.5x7.5 cm, four layers</td></tr><tr><td>Straight Doyen scissors</td><td>Rudolf Medical</td><td>RU/1428-16</td><td>For use to the donor</td></tr><tr><td>Straight micro jeweller forceps</td><td>Rudolf Medical</td><td>RU 4240-04</td><td>10.5 cm long. Used throughout the anastomosis</td></tr><tr><td>Syringes</td><td>Artsana SpA</td><td>N/A</td><td>20 mL (for the harvesting animal) and 5 mL (for the recipient). For saline flushing and dipping</td></tr><tr><td>TiCron 4-0</td><td>Covidien</td><td>CV-331</td><td>For closing muscles and skin</td></tr><tr><td>Tissue forceps V. Mueller</td><td>McKesson</td><td>CH 6950-009</td><td>Used for skin and muscles</td></tr><tr><td>Tramadol</td><td>SALF SpA</td><td>AIC 044718029</td><td>Analgesic. Single dose, 5 mg/kg intramuscular</td></tr><tr><td>Virgin silk 8-0</td><td>Johnson &amp;amp; Johnson Medical SpA</td><td>W818</td><td>For arterial branch ligation</td></tr></tbody></table>

a rodent model of the ross operation: syngeneic pulmonary artery graft implantation in a systemic position

The Ross operation for aortic valve disease has regained new interest due to its outstanding long-term results. Nonetheless, when employed as freestanding root replacement, the possible dilation of the pulmonary autograft and subsequent aortic regurgitation is described. Several animal models have been proposed. However, these are usually limited to ex-vivo models or in-vivo experiments with relatively expensive large animal models. In this study, we sought to establish a rodent model of pulmonary artery graft (PAG) implantation in a systemic position. A total of 39 adult Lewis rats were included. Immediately after euthanasia, the pulmonary root was harvested from a donor animal (n=17). Syngeneic recipient (n=17) and sham-operated (n=5) rats were sedated and ventilated. In the recipient group, the PAG was implanted with an end-to-end anastomosis in infra-renal abdominal aortic position. Sham-operated rats underwent only transection and re-anastomosis of the aorta. Animals were followed with serial ultrasound studies for two months and post-mortem histological analysis. The median PAG diameter in the native position was 3.20 mm (IQR=3.18-3.23). At follow-up, the median diameter of the PAG was 4.03 mm (IQR=3.74-4.13) at 1 week, 4.07 mm (IQR=3.80-4.28) at 1 month, and 4.27 mm (IQR=3.90-4.35) at 2 months (p&lt;0.01). Peak systolic velocity was 220.07 mm/s (IQR=210.43-246.41) at 1 week, 430.88 mm/s (IQR=375.28-495.56) at 1 month, and 373.68 mm/s (IQR=305.78-429.81) at 2 months (p=0.02) and did not differ from the sham-operated group at the end of the experiment (p=0.5). Histological analysis did not show any sign of endothelial thrombosis. This study showed that rodent models may allow for the evaluation of the long-term adaptation of the pulmonary root to a high-pressure system. A systemically placed syngeneic PAG implantation represents a simple and feasible platform for the development and evaluation of novel surgical techniques and drug therapies to further improve the outcomes of the Ross operation.

A total of 39 adult Lewis rats were included in this study: 17 animals were used as PAG donors, 17 animals as recipients and 5 as sham-operated (control group) (Table 1). Male rats were 22 (56%) and female 17 (44%); the latter were used only in the donor group.
No fatal event occurred during the operation with 100% survival. During the follow-up, two animals of the transplant group had a fatal outcome, at 12 and 51 days, respectively; the survival rate at the end of the study ...

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A Rodent Model of The Ross Operation: Syngeneic Pulmonary Artery Graft Implantation in A Systemic Position

We demonstrate how to establish a murine model of pulmonary root implantation into the descending aorta to simulate the Ross procedure. This model enables the medium/long-term evaluation of pulmonary autograft remodeling in a systemic position, representing the basis of developing therapeutic strategies to promote its adaptation.

The Ross operation for aortic valve disease has regained new interest due to its outstanding long-term results. Nonetheless, when employed as freestanding ...

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3.0K Views.  University of Padua. We demonstrate how to establish a murine model of pulmonary root implantation into the descending aorta to simulate the Ross procedure. This model enables the medium/long-term evaluation of pulmonary autograft remodeling in a systemic position, representing the basis of developing therapeutic strategies to promote its adaptation.

Video: A Rodent Model of The Ross Operation: Syngeneic Pulmonary Artery Graft Implantation in A Systemic Position

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 ...

Development of Obliterative Bronchiolitis in a Murine Model of Orthotopic Lung Transplantation

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and standardized not only for the study of allo-rejection but also between syngeneic strains for examining mechanisms of ischemia-reperfusion injury after lung transplantation. Although the application of the rat and other large animal model 2 contributed significantly to the elucidation of these studies, the scope of those investigations is limited by the scarcity of knockout and transgenic rats. Due to no effective therapies for obliterative bronchiolitis, the leading cause of death in lung transplant patients, there has been an intensive search for pre-clinical models that replicate obliterative bronchiolitis. The tracheal allograft model is the most widely used and may reproduce some of the histopathologic features of obliterative bronchiolitis 3. However, the lack of an intact vasculature with no connection to the recipient's conducting airways, and incomplete pathologic features of obliterative bronchiolitis limit the utility of this model 4. Unlike transplantation of other solid organs, vascularized mouse lung transplants have only recently been reported by Okazaki and colleagues for the first time in 2007 5. Applying the basic principles of the rat lung transplant, our lab initiated the obliterative bronchiolitis model using minor histoincompatible antigen murine orthotopic single-left lung transplants which allows the further study of obliterative bronchiolitis immunopathogenesis6.

Obliterative bronchiolitis is the key impediment to the long-term survival of lung transplant recipients and the lack of a robust preclinical model precludes examining obliterative bronchiolitis immunopathogenesis. Unlike other solid organ transplants, vascularized mouse lung transplantation has only recently been developed. Here we show our independently developed obliterative bronchiolitis model after murine orthotopic single-lung transplantation.

Orthotopic lung transplantation in rats was first reported by Asimacopoulos and colleagues in 1971 1. Currently, this method is well accepted and ...

Implantation of Engineered Tissue in the Rat Heart

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in common laboratory rats or mice and these quite frequently serve as an intermediate step between bench-top experiments and large animal testing or human trials. Given that rodents provide an established, cost-effective, and physiologically-relevant model system in which to test novel combinations of scaffolding materials and cells, they are particularly well-suited for cardiovascular tissue engineering studies. Presently, we describe an open-heart surgical procedure to implant engineered tissue containing myogenic progenitor cells in the atrioventricular (AV) groove of a rat heart. These implants are intended to create an electrical conduit between the right atrium and right ventricle with the ultimate goal of providing an alternative treatment to conventional pacemaker implantation in pediatric patients with complete heart block[1]. The engineered tissue is implanted in the AV-groove by means of a thoracotomy. For our purposes, Lewis rats are anesthetized and invasively ventilated to maintain positive airway pressure during the sterile surgical procedure. The approach to the heart is performed by a right thoracotomy through an antero-lateral incision at the 5th intercostal space. The tissue construct is fixed in the AV groove using a single 7-0 Prolene suture and positioned between the right ventricle and atrium at the ventral portion of the heart. The epicardium is partially removed to allow direct contact between the recipient myocardial cells and those contained in the engineered tissue. Following implantation, the chest wall is closed in layers, any pneumothorax is evacuated, and the animal is extubated and treated with analgesic.

Here, we describe a cardiac surgical procedure to implant engineered tissue in the atrioventricular (AV)-groove of an adult Lewis rat.

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in ...

Biology

Mouse Models for Graft Arteriosclerosis

Graft arteriosclerois (GA), also called allograft vasculopathy, is a pathologic lesion that develops over months to years in transplanted organs characterized by diffuse, circumferential stenosis of the entire graft vascular tree. The most critical component of GA pathogenesis is the proliferation of smooth muscle-like cells within the intima. When a human coronary artery segment is interposed into the infra-renal aortae of immunodeficient mice, the intimas could be expand in response to adoptively transferred human T cells allogeneic to the artery donor or exogenous human IFN-&gamma; in the absence of human T cells. Interposition of a mouse aorta from one strain into another mouse strain recipient is limited as a model for chronic rejection in humans because the acute cell-mediated rejection response in this mouse model completely eliminates all donor-derived vascular cells from the graft within two-three weeks. We have recently developed two new mouse models to circumvent these problems. The first model involves interposition of a vessel segment from a male mouse into a female recipient of the same inbred strain (C57BL/6J). Graft rejection in this case is directed only against minor histocompatibility antigens encoded by the Y chromosome (present in the male but not the female) and the rejection response that ensues is sufficiently indolent to preserve donor-derived smooth muscle cells for several weeks. The second model involves interposing an artery segment from a wild type C57BL/6J mouse donor into a host mouse of the same strain and gender that lacks the receptor for IFN-&gamma; followed by administration of mouse IFN-&gamma; (delivered via infection of the mouse liver with an adenoviral vector. There is no rejection in this case as both donor and recipient mice are of the same strain and gender but donor smooth muscle cells proliferate in response to the cytokine while host-derived cells, lacking receptor for this cytokine, are unresponsive. By backcrossing additional genetic changes into the vessel donor, both models can be used to assess the effect of specific genes on GA progression. Here, we describe detailed protocols for our mouse GA models.

We describe protocols for our mouse graft arteriosclerois (GA) models which involve interposition of a mouse vessel segment into a recipient of the same inbred strain. By backcrossing additional genetic changes into the vessel donor, the model can assess the effect of specific genes on GA.

Graft arteriosclerois (GA),  also called allograft vasculopathy, is a pathologic lesion that develops over  months to years in transplanted organs ...

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 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 ...

An Immunological Model for Heterotopic Heart and Cardiac Muscle Cell Transplantation in Rats

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications have been reported since the first description in 1964. After 30 years of performing heterotopic heart transplantation in rats, we have developed a simplified surgical approach, which can be easily taught and performed without further surgical training or background.
After dissection of the ascending aorta and the pulmonary artery and ligation of superior and inferior caval and pulmonary veins, the donor heart is harvested and subsequently perfused with ice-cold saline solution supplemented with heparin. After clamping and incising the recipient abdominal vessels, the donor ascending aorta and pulmonary artery are anastomosed to the recipient abdominal aorta and inferior vena cava, respectively, using continuous running sutures.
Depending on different donor-recipient combinations, this model allows analyses of either acute or chronic rejection of allografts. The immunological significance of this model is further enhanced by a novel approach of in-ear injection of vital cardiac muscle cells and subsequent analysis of draining cervical lymphatic tissue.

We describe a model of heterotopic abdominal heart transplantation in rats, implying modifications of current strategies, which lead to a simplified surgical approach. Additionally, we describe a novel rejection model by in-ear injection of vital cardiac muscle cells, allowing further transplant immunological analyses in rats.

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications ...

Immunology and Infection

A Rat Lung Transplantation Model of Warm Ischemia/Reperfusion Injury: Optimizations to Improve Outcomes

From our experience with rat lung transplantation, we have found several areas for improvement. Information in the existing literature regarding methods for choosing appropriate cuff sizes for the pulmonary vein (PV), pulmonary artery (PA), or bronchus (Br) are varied, thus making the determination of proper cuff size during rat lung transplantation an exercise of trial and error. By standardizing the cuffing technique to use the smallest effective cuff appropriate for the size of the vessel or bronchus, one can make the transplantation procedure safer, faster, and more successful. Since diameters of the PV, PA, and Br are related to the body weight of the rat, we present a strategy to choosing an appropriate size using a weight-based guide. Since lung volume is also related to body weight, we recommend that this relationship should also be considered when choosing the proper volume of air for donor lung inflation during warm ischemia as well as for the proper volume of PBS to be instilled during bronchoalveolar lavage (BAL) fluid collection. We also describe methods for 4th intercostal space dissection, wound closure, and sample collection from both the native and transplanted lobes.

Here, we present optimizations to a rat lung transplantation model that serve to improve outcomes. We provide a size guide for cuffs based on body weight, a measurement strategy to ascertain the 4th intercostal space, and methods of wound closure and BAL (bronchoalveolar lavage) fluid and tissue collection.

From our experience with rat lung transplantation, we have found several areas for improvement. Information in the existing literature regarding methods ...

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of transplant with the potential to deliver durable heart valves capable of somatic growth with no requirement for anticoagulation. However, the immunobiology of heart valve transplants remains unexplored, highlighting the need for animal models to study this new type of transplant. Previous rat models for heterotopic aortic valve transplantation into the abdominal aorta have been described, though they are technically challenging and costly. For addressing this challenge, a renal subcapsular transplant model was developed in rodents as a practical and more straightforward method for studying heart valve transplant immunobiology. In this model, a single aortic valve leaflet is harvested and inserted into the renal subcapsular space. The kidney is easily accessible, and the transplanted tissue is securely contained in a subcapsular space that is well vascularized and can accommodate a variety of tissue sizes. Furthermore, because a single rat can provide three donor aortic leaflets and a single kidney can provide multiple sites for transplanted tissue, fewer rats are required for a given study. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

This protocol describes a simple and efficient method for the transplantation of aortic valve leaflets under the renal capsule to allow for the study of alloreactivity of heart valves.

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of ...

A Rat Orthotopic Renal Transplantation Model for Renal Allograft Rejection

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential model to investigate the mechanism of renal allograft rejection in pre-clinical studies and could aid in the development of novel approaches to improve the long-term survival of renal allografts. Donor kidney implantation in rat orthotopic renal transplantation is commonly performed by end-to-side anastomosis to recipients' aorta and inferior vena cava. In this model, the donor's kidney was implanted using end-to-end anastomosis to the recipients' renal artery and renal vein. The donor's ureter was anastomosed to the recipient's bladder in an end-to-side 'tunnel' method. This model contributes to better healing of ureter-bladder anastomosis and increases the recipients' survival by avoiding interference with blood supply and venous reflux of the lower body. This model can be used to investigate the mechanisms of acute and chronic immune and pathologic rejection of renal allografts. Here, the study describes the detailed protocols of this orthotopic renal transplantation between rats.

The rat orthotopic renal transplantation model contributes to investigating the mechanism of renal allograft rejection. The current model increases the recipients' survival without interference with blood supply and venous reflux of the lower body using an end-to-end anastomosis of kidney implantation and an end-to-side "tunnel" method of ureter-bladder anastomosis.

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential ...