We extend our heartfelt appreciation to all who contributed to this work, both past and present members. This work was supported by grants from the German Federal Ministry for Economic Affairs and Energy, EXIST - Transfer of Research (03EFIBE103). Yimeng Hao is supported by the China Scholarship Council (CSC: 202008450028).

This preclinical study approved by the legal and ethical committee of the Regional Office for Health and Social Affairs, Berlin (LAGeSo). All animals&#160;(Ovis aries) received humane care in compliance with the guidelines of the European and German Laboratory Animal Science Societies (FELASA, GV-SOLAS). The procedure is illustrated by performing autologous pulmonary valve replacement in a 3-year-old, 47 kg, female sheep J.
1. Preoperative management
<ol>
	<li>House all experimental sheep in the same room containing straw for 1 week from the day of arrival to the pericardiectomy day to maintain social companionship (Figure 1A).</li>
	<li>Deprive the sheep of food but not water for 12 h prior to the pericardiectomy and implantation.</li>
	<li>Pre-medicate the sheep with an intramuscular injection of midazolam (0.4 mg/kg), butorphanol (0.4 mg/kg), and glycopyrrolate (0.011 mg/kg or 200 mcg) 20 min before intubation.</li>
</ol>
2. Induction of general anesthesia
<ol>
	<li>Aseptically place an 18 G safety intravenous (IV) catheter, an injection port, and a T port in the cephalic vein (Figure 1B).</li>
	<li>Induce anesthesia by intravenous injection of propofol (20 mg/mL, 1&#8211;2.5 mg/kg) and fentanyl (0.01 mg/kg) to effect.</li>
	<li>Indications of an adequate level of sedation include jaw relaxation, loss of swallowing, and papillary reflex. After sedation, intubate the sheep with an appropriately sized endotracheal tube (Figure 1C). Shave the sheep and then transfer it to the operating room (OR).</li>
</ol>
3. Intra-operative anesthesia management for pericardiectomy and implantation
<ol>
	<li>Use a pressure-cycled mechanical ventilator to initiate intermittent positive pressure ventilation (IPPV) with 100% oxygen in the OR.</li>
	<li>Connect the sheep to the anesthetic device platform and ventilate the sheep throughout the anesthesia under pressure mode (tidal volume (TV) = 8-12 mL/kg, respiratory frequency (RF) = 12-14 breaths/min). Adjust the TV and RF to keep the end-tidal carbon dioxide (EtCO2) between 35-45 mmHg and the arterial partial pressure of CO2 (PaCO2) below 50 mmHg.</li>
	<li>Maintain anesthesia combined with isoflurane (to effect, suggested maintenance concentration 1.5%-2.5%) in oxygen with a flow rate of 1 L/min (inspired fraction of oxygen (FiO2) = 75%), combined with a continuous rate infusion (CRI) of fentanyl (5-15 mcg/kg/h) and midazolam (0.2-0.5 mg/kg/h).</li>
	<li>Place an 18 G safety IV catheter in the auricular artery for the measurement of invasive blood pressure (IBP).</li>
	<li>Connect the sheep to the multi-function anesthesia platform for hemodynamic monitoring, which displays the direct measurement of invasive blood pressure (IBP) in the auricular artery (zeroed at the level of the heart), body temperature with a rectal probe, a lead-IV electrocardiogram, plethysmographic oxygen saturation (SpO2), TV, RF, EtCO2, heart rate (HR), and FiO2.</li>
	<li>Position a gastric tube to evacuate excess gas and fluids from the reticulorumen in preparation for the pericardiectomy. Equip the gastric tube with a marker guide wire as a reference for the implantation.</li>
	<li>Place a foley urinary catheter via the urethra inside the bladder connected to a urine bag. Distend the foley balloon with a minimum of 5 mL of saline solution (0.9% NaCl).</li>
	<li>Carry out an activated coagulation test (ACT: 240-300 s) 30 min before implantation to confirm sufficient heparinization before and antagonization after the implantation. Perform arterial blood gas analysis (ABGs) to analyze the internal environment 30 min prior to pericardiectomy and implantation and every hour during the two procedures.</li>
	<li>Administer the following antibiotics, namely, sulbactam/ampicillin (20mg/kg) 30 min via intravenous drip prior to pericardiectomy and implantation. Ensure a continuous infusion of crystalloids (5 mL/kg/h, isotonic balanced electrolyte solution) and hydroxyethyl starch (HES, 30 mL/h) throughout the pericardiectomy and implantation.</li>
</ol>
4. Pericardiectomy
<ol>
	<li>Preparation for pericardiectomy
	<ol>
		<li>Place the sheep on the operating table in the right lateral recumbent position with 30&#176; elevation on the left side, and then secure her limbs with harnesses and straps.</li>
		<li>Sterilize the surgical site (pericardiectomy: superiorly to the left clavicle, anteriorly to the sternum, inferiorly to the level of the diaphragm, and posteriorly to the left midclavicular line) with chlorhexidine-alcohol before performing the minithoracotomy. Cover the remaining areas with sterile draping (Figure 2A).</li>
		<li>Make a 5 cm skin incision at the fourth intercostal parasternal position using a #10 surgical blade under general anesthesia.</li>
		<li>Dissect the pectoralis major- pectoralis minor- anterior serratus-intercostal muscle via the left lateral minithoracotomy (m-LLT) into 5 cm incisions in length consecutively and separately in the third and fourth intercostal space for ideal exposure (Figure 2B).</li>
		<li>Make the incision at least 2 cm offset from the sternum to prevent injury to the left internal thoracic artery and veins. Cease the ventilator for 10 s to prevent lung injury before opening the thorax.</li>
		<li>Use several sterile gauzes to compress the left lung for better exposure of the surgical field after placing a rib spreader (Figure 2C). Visualize the pericardium and thymus in the surgical field (Figure 2D).</li>
	</ol>
	</li>
	<li>Start the pericardiectomy at the attachment point of the pericardium and diaphragm and harvest the pericardial tissue between the two phrenic nerves, up to the innominate veins, down to the diaphragm.
	<ol>
		<li>Compress the left lung as mentioned in step 4.1.5 to expose the attachment of the diaphragm-pericardium-mediastinal pleura. Cut open the left mediastinal pleura at the attachment of the diaphragm-pericardium-mediastinal pleura by making a 1 cm incision in length using a surgical scissor. Extend the incision upward into the innominate veins along the line which is 1 cm offset from the left phrenic nerve (Figure 2E).</li>
		<li>Repeat the procedure for the right part of the pericardium by elevating the apex to the left using fingers. Dissect the thymic and pericardial fat from the sternum.</li>
		<li>Meet the two incisions of the pericardium in front of the aorta. Cross clamp the intersection of pericardium and thymus from the two pericardial incisions in front of the aorta by holding them firmly in place and tying six surgical knots manually using a 4-0 non-resorbable suture.</li>
		<li>Avoid injury of the phrenic nerve and the underlying vascular structures, when harvesting the pericardium. Dissect adipose tissue including the thymus from the surface of the pericardium during pericardiectomy. Use a cautery tool (i.e., electrotome, Bovie) for hemostasis.</li>
	</ol>
	</li>
	<li>Place the harvested pericardium onto the sterile plate with a centimeter scale to remove the extra adipose tissue, and then wash it twice in 0.9% NaCl (Figure 2F). Double-check all the surgical areas for hemostasis.</li>
	<li>Suture the opened right mediastinal pleura to the residual right pericardial edge with 3-0 polydioxanone in a running fashion twice. Inflate the right lung to the largest volume manually using a breathing bag and hold for 10 s prior to closing the right thorax. Suture the opened left mediastinal pleura to the residual left pericardial edge with 3-0 polydioxanone in a running fashion twice.</li>
	<li>Close the left thoracic incisions in four layers as described below.
	<ol>
		<li>Suture the intercostal muscles and anterior serratus with 2- 0 polydioxanone in a simple interrupted or cruciate fashion, pectoralis major-pectoralis minor with 3-0 polydioxanone in a running fashion, the subcutis with 3-0 polydioxanone in a cruciate fashion, and the skin with 3-0 nylon in a simple interrupted fashion. Place all the sutures at 1 cm intervals.</li>
		<li>Inflate the left lung to the largest volume manually using a breathing balloon and hold for 10 s prior to closing the intercostal muscles.</li>
	</ol>
	</li>
	<li>Cover the incision with sterile gauze and compress it manually for 5 min to prevent hemorrhage after heparinization for the new heart valve implantation. Then bandage the surgical site.</li>
	<li>Stop the intravenous anesthetics and isoflurane when performing the skin suture to reduce the depth of sedation.</li>
	<li>Remove the gastric tube and urinary catheter after the spontaneous respiration returns. Then transfer the sheep with pulse oximetry to the recovery room on the stretcher.</li>
	<li>Remove the endotracheal tube when the swallowing reflex, the papillary reflex, and normal spontaneous breathing recover. Administer 0.5 mg/kg meloxicam subcutaneously once a day before the implantation.</li>
	<li>Once the anesthesia is completely reversed (i.e., when the sheep is able to stand independently), the sheep can be given access to food and water.</li>
</ol>
5. Preparation of the three-dimensional autologous heart valve
<ol>
	<li>Trim the pericardium by removing the adipose tissue (Figure 3A,B,C), and then place it onto the 3D shaping heart valve mold. (Due to a pending patent application, figures cannot be provided in this step.)</li>
	<li>Put the pericardium and the 3D shaping heart valve model into an incubator with a non-toxic crosslinker (30 mL) for 2 days and 21 h (Figure 3D; due to the pending&#160;patent application, figures and detailed information of non-toxic crosslinker cannot be provided in this step).</li>
</ol>
6. Preparation of the APV
<ol>
	<li>Wash the crosslinked heart valve in 0.9% NaCl twice and suture it into a Nitinol stent (30 mm in diameter, 29.4 mm in height, 48 rhombic cells) in a discontinuous fashion after 2 days and 21 h. Use 5-0 polypropylene to suture the heart valve in place using six to eight knots to align the attachment points between the heart valve and stent. (Due to a patent application, figures cannot be provided in this step.)</li>
	<li>Cut the three free edges of the autologous pulmonary valve open with a no. 15 surgical blade (Figure 4A,B). Hold the stented pulmonary valve with a surgical tweezer, lift and leave the APV in 0.9% NaCl to test its opening and closing and to evaluate whether the three commissures need further cutting to achieve a larger opening of the orifice.</li>
	<li>Incubate the APV in an incubator for 30 min for sterilization in 47.6 mL of PBS with 0.8% amphotericin B (0.4 mL) and 4.0% penicillin/streptomycin (2 mL). Crimp the stented heart valve into the head of a delivery system (DS) using a commercial crimper for two-fold testing (Figure 4C-D) and fit it into the delivery system (Figure 4E).</li>
</ol>
7. Transcatheter autologous pulmonary valve implantation via the left jugular vein
<ol>
	<li>Anesthetize the sheep for APV implantation as illustrated in steps 1 to 3.</li>
	<li>Vessel access: Shave the sheep and sterilize the surgical field, which includes superiorly to the inferior border of the mandible, anteriorly to the anterior median line, inferiorly to the superior border of left clavicle, and posteriorly to the posterior median line using a povidone-iodine antiseptic before performing the implantation. Cover the remaining unshaved and unsterilized areas with sterile draping.
	<ol>
		<li>Mark the left jugular vein on the neck and using the Seldinger technique place the guidewire into the left jugular vein. Enlarge the puncture point with a no. 10 blade, place a 11 F sheath into the left jugular vein for the ICE probe and delivery system (Figure 5A,B). Place a purse-string suture around the sheath introducer with a 4-0 non-absorbable suture.</li>
	</ol>
	</li>
	<li>Intracardiac echocardiography (ICE)17
	<ol>
		<li>Perform ICE before and immediately after the implantation using a 10 Fr ultrasound catheter (Figure 5C). Asses the parameters including the dimensions and functions of the NPV, APV, and tricuspid valve by 2D, color, pulsed wave, and continuous Doppler in the short and long axis.</li>
		<li>Evaluate the degree of valvular regurgitation in the vena contracta by semi-quantitative assessment18&#160;via ICE (Figure 6).</li>
	</ol>
	</li>
	<li>Angiography19: Perform angiography using a portable C-arm and a functional screen to guide the implantation by measuring the diameters of the RVOT, NPV, pulmonary bulb, and supravalvular pulmonary artery, as well as to evaluate the APV after implantation (Figure 7A-D).</li>
	<li>Hemodynamics20: Measure and record the right ventricular and pulmonary artery pressure before and after the implantation using a 5.2 F 145&#176; pigtail catheter. Measure the systemic arterial pressure via the auricular artery.</li>
	<li>Implantation
	<ol>
		<li>Establishment of the TPVR tract: Place a 0.035-inch angled guidewire to the right pulmonary artery under the guidance of fluoroscopy. Then, place a 5.2 Fr pigtail catheter into the left jugular vein and advance it into the right pulmonary artery with the guidance of the previously placed guidewire under fluoroscopy.</li>
		<li>Retrieve the angled guidewire out of the left jugular vein. Place a 5 Fr Berman angiographic balloon catheter into the left jugular vein and advance it into the right pulmonary artery using the guidance of the guidewire.</li>
		<li>Pre-shape the 0.035-inch ultra-stiff guidewire into a circle of about 8-10 cm in length with a diameter equaling the distance from the central point of the tricuspid valve to the central point of the pulmonary valve according to the fluoroscopy measurement and advance it into the right pulmonary artery under the guidance of the balloon catheter (Figure 8A). Ensure the wire does not interfere with the tricuspid valve chordae.</li>
		<li>Dilate the skin with a no. 11 blade and dilate the left jugular vein using commercial dilators from 16 Fr to 22 Fr sequentially (Figure 8B). Close the incision with a 3-0 polydioxanone purse-string suture after dilatation (Figure 8C). Perform angiography to ensure the desired position of the stent-bearing part of the DS as described in19.</li>
		<li>Mark the sinotubular junction of the pulmonary valve at the end-systolic and end-diastolic cardiac phases during pulmonary angiography as the distal border of the landing zone and the basal plane of the pulmonary valve as the proximal border of the landing zone.</li>
		<li>Reopen and inspect the stented autologous valve for crimping-induced damage. Re-crimp the APV and fit it into the head of the DS (Figure 8D). Advance the loaded DS via the pre-shaped guidewire through the right ventricular inflow tract (RVIT) and the RVOT to the NPV position (Figure 8E,F, and Figure 9A).</li>
		<li>Retract the cover tube of the DS and deploy the APV slowly and directly over the NPV in the landing zone at the end of the diastolic phase under fluoroscopic guidance (Figure 9A-C). Exercise caution when loaded DS is crossing the junction between the RVIT and the RVOT in order to prevent myocardial injury and ventricular fibrillation. The optimal position for the APV is when the middle part of the stent is placed onto the NPV.</li>
		<li>Retract the tip of the DS carefully into the cover tube after deployment and retrieve the DS from the sheep (Figure 9D). Repeat ICE (Figure 6D-F), angiography (Figure 7C-D), and hemodynamic measurements for post-examination of the dimensions and functions of the implanted APV. Close the incision on the left side of the neck with the pre-placed purse-string suture and compress it manually.</li>
	</ol>
	</li>
</ol>
8. Peri-implantation medication
<ol>
	<li>Prior to implantation, administer the sheep with heparin at a dose of 5000 IU to maintain an activated clotting time (ACT) of 240-300 s. Use ACT tests throughout the procedure. Repeat ACT tests every 30 min after the start of the procedure to confirm both sufficient heparinization before and antagonization after the implantation.</li>
	<li>Before the APV implantation, administer 10% magnesium at a dose of 0.02 mol/L and amiodarone at a dose of 3-5 mg/kg to prevent cardiac arrhythmias.</li>
	<li>Administer sulbactam/ampicillin (20 mg/kg) intravenously to prevent infection and endocarditis at the start of the pericardiectomy and implantation procedure.</li>
</ol>
9. Postoperative management
<ol>
	<li>Perform a daily postoperative follow-up for 5 days, checking the general condition of the sheep in terms of heart rate and rhythm, breathing depth, breathing rhythm, and breath sound (for checking postoperative pneumonia), signs of pain, and other abnormalities. Check the wound for postoperative swelling, inflammation, redness, bleeding, and secretion.</li>
	<li>Continue anticoagulation for 5 days with dalteparin 5000 IU or another low-molecular-weight heparin administered subcutaneously once daily. Administer 1 mg/kg meloxicam by subcutaneous injection for postoperative analgesia for 5 days.</li>
	<li>Perform a laboratory blood test, including hematology, liver function, renal function, and serum chemistry to evaluate the sheep&#39;s physical condition.</li>
</ol>
10. Follow-up
<ol>
	<li>Perform ICE, cardiac magnetic resonance imaging (cMRI), angiography, and record hemodynamics every 3-6 months after implantation for up to 21 months. Perform ICE and angiography as illustrated above.</li>
	<li>Perform cMRI to evaluate the regurgitation fraction (RF) on a 3.0 T MRI scanner using a standard electrocardiogram-gated cine-MRI method21. Perform final cardiac computed tomography (CT) to evaluate the stent position and the deformation of the right heart throughout the entire cardiac cycle as illustrated in our previous study22.</li>
</ol>

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(179), e63367 (2022).</li><li>Knirsch, W., 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=Establishing+a+pre-clinical+growing+animal+model+to+test+a+tissue+engineered+valved+pulmonary+conduit.">Establishing a pre-clinical growing animal model to test a tissue engineered valved pulmonary conduit.</a> Journal of Thoracic Disease. 12 (3), 1070-1078 (2020).</li><li>Zhang, X., 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=Tissue+engineered+transcatheter+pulmonary+valved+stent+implantation:+current+state+and+future+prospect.">Tissue engineered transcatheter pulmonary valved stent implantation: current state and future prospect.</a> International Journal of Molecular Sciences. 23 (2), 723 (2022).</li><li>Al Hussein, 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=Challenges+in+perioperative+animal+care+for+orthotopic+implantation+of+tissue-engineered+pulmonary+valves+in+the+ovine+model.">Challenges in perioperative animal care for orthotopic implantation of tissue-engineered pulmonary valves in the ovine model.</a> Tissue Engineering and Regenerative Medicine. 17 (6), 847-862 (2020).</li><li>Emmert, M. Y., 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=Computational+modeling+guides+tissue-engineered+heart+valve+design+for+long-term+in+vivo+performance+in+a+translational+sheep+model.">Computational modeling guides tissue-engineered heart valve design for long-term in vivo performance in a translational sheep model.</a> Science Translational Medicine. 10 (440), (2018).</li><li>Schmidt, 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=Minimally-invasive+implantation+of+living+tissue+engineered+heart+valves:+.+a+comprehensive+approach+from+autologous+vascular+cells+to+stem+cells.">Minimally-invasive implantation of living tissue engineered heart valves: . a comprehensive approach from autologous vascular cells to stem cells.</a> Journal of the American College of Cardiology. 56 (6), 510-520 (2010).</li></ol>

The authors have no financial conflicts of interest to disclose.

This study represents an important step forward in developing a living pulmonary valve for TPVR. In an adult sheep model, the method was able to show that an APV derived from the sheep&#39;s own pericardium can be implanted with a self-expandable Nitinol stent via jugular vein catheterization. In&#160;sheep J, the stented autologous pulmonary valve was successfully implanted in the correct pulmonary position using a self-designed universal delivery system. After implantation, the heart valve of sheep J showed go...

Bonhoeffer et al.1 marked the beginning of transcatheter pulmonary valve replacement (TPVR) in 2000 as a rapid innovation with significant progress toward minimizing complications and providing an alternative therapeutic approach. Since then, the use of TPVR for treating the right ventricular outflow tract (RVOT) or bioprosthetic valve dysfunction has increased rapidly2,3. To date, the TPVR devices currently available on the market have provided satisfying long-term and short-term results for patients with RVOT dysfunction4,5,6. Furthermore, various types of TPVR valves including decellularized heart valves and stem cell-driven heart valves are being developed and evaluated, and their feasibility has been demonstrated in preclinical large animal models7,8. Aortic valve reconstruction using an autologous pericardium was first reported by Dr. Duran, for which three consecutive bulges of different sizes were used as templates to guide the shaping of the pericardium according to the dimensions of the aortic annulus, with the survival rate of 84.53% at the follow-up of 60 months9. The Ozaki procedure, which is considered a valve repair procedure rather than a valve replacement procedure, involves replacing aortic valve leaflets with the glutaraldehyde-treated autologous pericardium; however, when compared to Dr. Duran&#39;s procedure, it improved significantly in measuring the diseased valve with a template to cut fixed pericardium10 and satisfactory results were not only achieved from the adult cases but also pediatric cases11. Currently, only the Ross procedure can provide a living valve substitute for the patient who has a diseased aortic valve with obvious advantages in terms of avoiding long-term anticoagulation, growth potential, and low risk of endocarditis12. But re-interventions may be required for the pulmonary autograft and right ventricle to pulmonary artery conduit after such a complex surgical procedure.
The current bioprosthetic valves that are available for clinical use inevitably degrade over time due to graft-versus-host reactions to the xenogeneic porcine or bovine tissues13. Valve-related calcification, degradation, and insufficiency could necessitate repeated interventions after several years, especially in young patients who would need to undergo multiple pulmonary valve replacements in their lifetime due to the lack of growth of the valves, a property inherent to current bioprosthetic materials14. Furthermore, the currently available, essentially non-regenerative, TPVR valves have major limitations such as thromboembolic and bleeding complications, as well as limited durability due to adverse tissue remodeling which could lead to leaflet retraction and universal valvular dysfunction15,16.
It is hypothesized that developing a native-like autologous pulmonary valve (APV) mounted onto a self-expandable Nitinol stent for TPVR with the characteristics of self-repair, regeneration, and growth capacity would ensure physiological performance and long-term functionality. And the non-toxic crosslinker treated autologous pericardium can awake from the harvesting and manufacturing procedures. To this end, this preclinical trial was conducted to implant a stented autologous pulmonary valve in an adult sheep model with the aim of developing ideal interventional valvular substitutes and a low-risk procedural methodology to improve the transcatheter therapy of RVOT dysfunction. In this paper, sheep J was selected to illustrate the comprehensive TPVR procedure including pericardiectomy and trans jugular vein implantation of an autologous heart valve.

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			<td>10 % Magnesium</td>
			<td>Inresa Arzneimittel GmbH</td>
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			<td>Siemens Healthcare GmbH</td>
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			<td>Sanofi-Aventis Deutschland GmbH</td>
			<td>PZN: 4599382</td>
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			<td>Amplatz ultra-stiff guidewire</td>
			<td>COOK MEDICAL LLC, USA</td>
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			<td>Anesthetic device platform</td>
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			<td>ARROW Berman Angiographic Balloon Catheter</td>
			<td>Teleflex Medical Europe Ltd</td>
			<td>LOT: 16F16M0070</td>
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			<td>Richter Pharma AG</td>
			<td>Vnr531943</td>
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			<td>BV Pulsera, Philips Heathcare, Eindhoven, The Netherlands</td>
			<td>CAN/CSA-C22.2 NO.601.1-M90</td>
			<td>Medical electral wquipment</td>
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			<td>Edwards Lifesciences, Irvine, CA, USA</td>
			<td>9600CR</td>
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			<td>CT</td>
			<td>Siemens Healthcare GmbH</td>
			<td>&#8722;</td>
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			<td>Dilator</td>
			<td>Edwards Lifesciences, Irvine, CA, USA</td>
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			<td>Ethicon Suture</td>
			<td>Ethicon</td>
			<td>LOT:MKH259</td>
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			<td>Ethicon</td>
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			<td>ST. JUDE MEDICAL Minnetonka MN</td>
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			<td>Janssen-Cilag Pharma GmbH</td>
			<td>DE/H/1047/001-002</td>
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			<td>Pfizer Pharma GmbH, Berlin, Germany</td>
			<td>PZN: 5746520</td>
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			<td>Functional screen</td>
			<td>BV Pulsera, Philips Heathcare, Eindhoven, The Netherlands</td>
			<td>System ID: 44350921</td>
			<td>Medical electral wquipment</td>
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			<td>Hemochron Celite ACT</td>
			<td>International Technidyne Corporation, Edison, USA</td>
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			<td>ACT</td>
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			<td>Merckle GmbH</td>
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			<td>Hydroxyethyl starch (Haes-steril 10 %)</td>
			<td>Fresenius Kabi Deutschland GmbH</td>
			<td>ATC Code: B05A</td>
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			<td>Imeron 400 MCT</td>
			<td>Bracco Imaging</td>
			<td>PZN00229978</td>
			<td>2.0&#8211;2.5 ml/kg, Contrast agent</td>
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			<td>Isoflurane</td>
			<td>CP-Pharma Handelsges. GmbH</td>
			<td>ATCvet Code: QN01AB06</td>
			<td>250 ml, MAC: 1 %</td>
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			<td>Jonosteril Infusionsl&#246;sung</td>
			<td>Fresenius Kabi Deutschland GmbH</td>
			<td>PZN: 541612</td>
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			<td>Ketamine</td>
			<td>Actavis Group PTC EHF</td>
			<td>ART.-Nr. 799-762</td>
			<td>2&#8211;5 mg/kg/h</td>
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			<td>Boehringer Ingelheim Vetmedica GmbH</td>
			<td>M21020A-09</td>
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			<td>Hameln pharma plus GMBH</td>
			<td>MIDAZ50100</td>
			<td>0.4mg/kg</td>
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			<td>Philips Healthcare</td>
			<td>&#8722;</td>
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			<td>Cordis, Miami Lakes, FL, USA</td>
			<td>REF: 533-534A</td>
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			<td>B. Braun Melsungen AG</td>
			<td>PZN 11164495</td>
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			<td>Propofol</td>
			<td>B. Braun Melsungen AG</td>
			<td>PZN 11164443</td>
			<td>10mg/ml, 2.5&#8211;8.0 mg/kg/h</td>
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			<td>Safety IV Catheter with Injection port</td>
			<td>B. Braun Melsungen AG</td>
			<td>LOT: 20D03G8346</td>
			<td>18 G Catheter with Injection port</td>
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			<td>Pfizer Pharma GmbH, Berlin, Germany</td>
			<td>PZN: 4843132</td>
			<td>3 g, 2.000 mg/ 1.000 mg</td>
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			<td>Sulbactam/ ampicillin</td>
			<td>Instituto Biochimico Italiano G Lorenzini S.p.A. &#8211; Via Fossignano 2, Aprilia (LT) &#8211; Italien</td>
			<td>ATC Code: J01CR01</td>
			<td>20 mg/kg, 2 g/1 g</td>
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			<td>Surgical Blade</td>
			<td>Brinkmann Medical ein Unternehmen der Dr. Junghans Medical GmbH</td>
			<td>PZN: 354844</td>
			<td>15 #</td>
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			<td>Surgical Blade</td>
			<td>Brinkmann Medical ein Unternehmen der Dr. Junghans Medical GmbH</td>
			<td>PZN: 354844</td>
			<td>11 #</td>
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			<td>Suture</td>
			<td>Johnson &#38; Johnson</td>
			<td>Hersteller Artikel Nr. EH7284H</td>
			<td>5-0 polypropylene</td>
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transcatheter pulmonary valve replacement from autologous pericardium with a self-expandable nitinol stent in an adult sheep model

Transcatheter pulmonary valve replacement has been established as a viable alternative approach for patients suffering from right ventricular outflow tract or bioprosthetic valve dysfunction, with excellent early and late clinical outcomes. However, clinical challenges such as stented heart valve deterioration, coronary occlusion, endocarditis, and other complications must be addressed for lifetime application, particularly in pediatric patients. To facilitate the development of a lifelong solution for patients, transcatheter autologous pulmonary valve replacement was performed in an adult sheep model. The autologous pericardium was harvested from the sheep via left anterolateral minithoracotomy under general anesthesia with ventilation. The pericardium was placed on a 3D shaping heart valve model for non-toxic cross-linking for 2 days and 21 h. Intracardiac echocardiography (ICE) and angiography were performed to assess the position, morphology, function, and dimensions of the native pulmonary valve (NPV). After trimming, the crosslinked pericardium was sewn onto a self-expandable Nitinol stent and crimped into a self-designed delivery system. The autologous pulmonary valve (APV) was implanted at the NPV position via left jugular vein catheterization. ICE and angiography were repeated to evaluate the position, morphology, function, and dimensions of the APV. An APV was successfully implanted in&#160;sheep J. In this paper, sheep J was selected to obtain representative results. A 30 mm APV with a Nitinol stent was accurately implanted at the NPV position without any significant hemodynamic change. There was no paravalvular leak, no new pulmonary valve insufficiency, or stented pulmonary valve migration. This study demonstrated the feasibility and safety, in a long-time follow-up, of developing an APV for implantation at the NPV position with a self-expandable Nitinol stent via jugular vein catheterization in an adult sheep model.

In sheep J, the APV (30 mm in diameter) were successfully implanted in the &#34;landing zone&#34; of the RVOT.
In sheep J, the hemodynamics remained stable throughout the left anterolateral minithoracotomy under general anesthesia with ventilation, as well as in the follow-up MRI and ICE (Table 1, Table 2, and Table 3). Autologous pericardium measuring 9 cm x 9 cm was harvested and trimmed by removing extra tissue (Figure ...

Watch this Scientific Journal Video about Transcatheter Pulmonary Valve Replacement from Autologous Pericardium with a Self-Expandable Nitinol Stent in an Adult Sheep Model at JoVE.com

Transcatheter Pulmonary Valve Replacement from Autologous Pericardium with a Self-Expandable Nitinol Stent in an Adult Sheep Model

This study demonstrates the feasibility and safety of developing an autologous pulmonary valve for implantation at the native pulmonary valve position by using a self-expandable Nitinol stent in an adult sheep model. This is a step toward developing transcatheter pulmonary valve replacement for patients with right ventricular outflow tract dysfunction.

Transcatheter pulmonary valve replacement has been established as a viable alternative approach for patients suffering from right ventricular outflow ...

transcatheter-pulmonary-valve-replacement-from-autologous-pericardium

Research

JoVE Journal

Medicine

2.7K Views.  Charité University Medicine Berlin. This study demonstrates the feasibility and safety of developing an autologous pulmonary valve for implantation at the native pulmonary valve position by using a self-expandable Nitinol stent in an adult sheep model. This is a step toward developing transcatheter pulmonary valve replacement for patients with right ventricular outflow tract dysfunction.

Video: Transcatheter Pulmonary Valve Replacement from Autologous Pericardium with a Self-Expandable Nitinol Stent in an Adult Sheep Model

Human Internal Mammary Artery (IMA) Transplantation and Stenting: A Human Model to Study the Development of In-Stent Restenosis

Preclinical in vivo research models to investigate pathobiological and pathophysiological processes in the development of intimal hyperplasia after vessel stenting are crucial for translational approaches1,2.
The commonly used animal models include mice, rats, rabbits, and pigs3-5. However, the translation of these models into clinical settings remains difficult, since those biological processes are already studied in animal vessels but never performed before in human research models6,7. In this video we demonstrate a new humanized model to overcome this translational gap. The shown procedure is reproducible, easy, and fast to perform and is suitable to study the development of intimal hyperplasia and the applicability of diverse stents.
This video shows how to perform the stent technique in human vessels followed by transplantation into immunodeficient rats, and identifies the origin of proliferating cells as human.

Human Internal Mammary Artery IMA Transplantation and Stenting: A Human Model to Study the Development of In-Stent Restenosis

This video shows a model to study the development of intimal hyperplasia after stent deployment using a human vessel (IMA) in an immunodeficient rat model.

Preclinical in vivo research models to investigate pathobiological and pathophysiological processes in the development of intimal hyperplasia after vessel ...

A Murine Model of Stent Implantation in the Carotid Artery for the Study of Restenosis

Despite the considerable progress made in the stent development in the last decades, cardiovascular diseases remain the main cause of death in western countries. Beside the benefits offered by the development of different drug-eluting stents, the coronary revascularization bears also the life-threatening risks of in-stent thrombosis and restenosis. Research on new therapeutic strategies is impaired by the lack of appropriate methods to study stent implantation and restenosis processes. Here, we describe a rapid and accessible procedure of stent implantation in mouse carotid artery, which offers the possibility to study in a convenient way the molecular mechanisms of vessel remodeling and the effects of different drug coatings.

A model of stent implantation in mouse carotid artery is described. Compared to other similar methods, this procedure is very rapid, simple and accessible, offering the possibility to study in a convenient way the vascular wall reaction to different drug-eluting stents and the molecular mechanisms of restenosis.

Despite  the considerable progress made in the stent development in the last decades,  cardiovascular diseases remain the main cause of death in western ...

Transplantation of Pulmonary Valve Using a Mouse Model of Heterotopic Heart Transplantation

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed results. However, the cellular and molecular mechanisms of the neotissue development, valve thickening, and stenosis development are not researched extensively. To answer the above questions, we developed a murine heterotopic heart valve transplantation model. A heart valve was harvested from a valve donor mouse and transplanted to a heart donor mouse. The heart with a new valve was transplanted heterotopically to a recipient mouse. The transplanted heart showed its own heartbeat, independent of the recipient&#8217;s heartbeat. The blood flow was quantified using a high frequency ultrasound system with a pulsed wave Doppler. The flow through the implanted pulmonary valve showed forward flow with minimal regurgitation and the peak flow was close to 100 mm/sec. This murine model of heart valve transplantation is highly versatile, so it can be modified and adapted to provide different hemodynamic environments and/or can be used with various transgenic mice to study neotissue development in a tissue engineered heart valve.

In order to understand the cellular and molecular mechanisms underlying neotissue formation and stenosis development in tissue engineered heart valves, a murine model of heterotopic heart valve transplantation was developed. A pulmonary heart valve was transplanted to recipient using the heterotopic heart transplantation technique.

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed ...

An Improved Method for Rapid Intubation of the Trachea in Mice

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity is a commonly used model to study both acute lung injury and fibrosis, and multiple methods have been developed for administering bleomycin (and other toxic agents) into the lungs. However, many of these approaches, such as transtracheal instillation, have inherent drawbacks, including the need for strong anesthetics and survival surgery. This paper reports a quick, reproducible method of intratracheal intubation that involves mild inhaled anesthesia, visualization of the trachea, and the use of a surrogate spirometer to confirm exposure. As a proof of concept, 8-12 week old C57BL/6 mice were administered either 2.0 U/kg of bleomycin or an equivalent volume of PBS, and both damage and fibrotic endpoints were measured post-exposure. This procedure allows researchers to treat a large cohort of mice in a relatively short period with little expense and minimal post-procedure care.

This article presents a rapid and simple method for administering bleomycin directly into the mouse trachea via intubation. Key advantages of this method are that it is highly reproducible, easy to master, and does not require specialized equipment or lengthy recovery times.

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity ...

Full-root Aortic Valve Replacement by Stentless Aortic Xenografts in Patients with Small Aortic Roots

In patients with small aortic roots who need an aortic valve replacement with biological valve substitutes, the implantation of the stented pericardial valve might not meet the functional needs. The implantation of a too-small stented pericardial valve, leading to an effective orifice area indexed to a body surface area less than 0.85 cm2/m2, is regarded as prosthesis-patient mismatch (PPM). A PPM negatively affects the regression of left ventricular hypertrophy and thus the normalization of left ventricular function and the alleviation of symptoms. Persistent left ventricular hypertrophy is associated with an increased risk of arrhythmias and sudden cardiac death. In the case of predictable PPM, there are three options: 1) accept the PPM resulting from the implantation of a stented pericardial valve when comorbidities of the patient forbid the more technically demanding operative technique of implanting a larger prosthesis, 2) enlarge the aortic root to accommodate a larger stented valve substitute, or 3) implant a stentless biological valve or a homograft. Compared to classical aortic valve replacement with stented pericardial valves, the full-root implantation of stentless aortic xenografts offers the possibility of implanting a 3-4 mm larger valve in a given patient, thus allowing significant reduction in transvalvular gradients. However, a number of cardiac surgeons are reluctant to transform a classical aortic valve replacement with stented pericardial valves into the more technically challenging full-root implantation of stentless aortic xenografts. Given the potential hemodynamic advantages of stentless aortic xenografts, we have adopted full-root implantation to avoid PPM in patients with small aortic roots necessitating an aortic valve replacement. Here, we describe in detail a technique for the full-root implantation of stentless aortic xenografts, with emphasis on the management of the proximal suture line and coronary anastomoses. Limitations of this technique and alternative options are discussed.

Full-root aortic valve replacement by stentless aortic xenograft is a viable option in patients with small aortic roots. We describe, a technique for the full-root implantation of stentless aortic xenografts, with emphasis on the management of the proximal suture line and coronary anastomoses, and discuss its limitations and alternative options.

In patients with small aortic roots who need an aortic valve replacement with biological valve substitutes, the implantation of the stented pericardial ...

Standardized Technique of Aortic Valve Re-implantation for Valve-sparing Aortic Root Replacement

Despite the obvious advantages of the preservation of a normal aortic valve during aortic root replacement, the complexity of valve sparing procedures prevents a number of cardiac surgeons from incorporating them into their practice. The aim of this protocol is to describe a simplified and user-friendly technique of an aortic valve-sparing root replacement (VSRR) procedure by re-implantation of the aortic valve. Proper selection of patients and limitations of the technique are discussed.
In 54 consecutive patients, normal appearing aortic valves were re-implanted in a commercially available polyester prosthesis with pre-shaped sinuses by a simplified and standardized technique. Placement of the first row of the proximal suture line, choice of the prosthesis size, and adjustment of the height of the commissures of the patient to the fixed height of the sinus portion of the prosthesis were slightly modified from the reference techniques with the aim of increasing its feasibility for use by other cardiac surgeons. Early mortality and morbidity as well as 5-year survival, freedom from aortic valve reoperation, and freedom from recurrent moderate regurgitation were collected in all patients.
Thirty-day mortality, re-sternotomy for bleeding, re-sternotomy for mediastinitis, and the incidence of stroke were very low, 1.8% for each (1 of 54). No patient required permanent pace-maker implantation. At 5 years, survival, freedom from aortic valve reoperation, and freedom from recurrent moderate regurgitation were 97.5%, 95.2%, and 91.6%, respectively.
Mid-term results of our standardized technique of re-implantation of the aortic valve for valve-sparing aortic root replacement are very good and compare with more complex techniques reported by experienced surgeons. By following the present protocol of the standardized re-implantation technique, a greater number of cardiac surgeons can perform this procedure with comparable good results.

Valve-sparing aortic root replacement has the advantage of preserving the patient&#39;s own aortic valve. The complexity of the reported techniques to date restricts their use to a limited number of cardiac surgeons. This protocol describes step-by-step a standardized technique reproducible by a greater number of cardiac surgeons.

Despite the obvious advantages of the preservation of a normal aortic valve during aortic root replacement, the complexity of valve sparing procedures ...

Improved Registration of 3D CT Angiography with X-ray Fluoroscopy for Image Fusion During Transcatheter Aortic Valve Implantation

The fusion of 3D anatomical models derived from high-fidelity pre-interventional computed tomography angiography (CTA), and x-ray (XR) fluoroscopy to facilitate anatomical guidance is of huge interest for complex cardiac interventions like TAVI procedures with cerebral protection. Co-registration of CTA and XR has been introduced either based on additional intraoperative non-/contrast-enhanced cone-beam computed tomography (CBCT) or two separate aortograms. With the related increase of radiation exposure and/or contrast agent (CA) dose, a potential additional risk for the patient is introduced. Here, we propose a modified co-registration approach making use of arteriograms of the iliofemoral arteries, routinely performed during the femoral puncture and sheath introduction. On-the-fly refinement of the co-registration during the on-going procedure enables accurate co-registration without any additional angiograms, thus reducing CA, XR dose and procedure time, while simultaneously improving operator confidence and procedure safety.

The aim of this study was to improve the co-registration for image fusion (IF) of pre-interventional CT data with real-time x-ray (XR) fluoroscopy during transfemoral transcatheter aortic valve implantation (TAVI).

The fusion of 3D anatomical models derived from high-fidelity pre-interventional computed tomography angiography (CTA), and x-ray (XR) fluoroscopy to ...

An Image Guided Transapical Mitral Valve Leaflet Puncture Model of Controlled Volume Overload from Mitral Regurgitation in the Rat

Mitral regurgitation (MR) is a widely prevalent heart valve lesion, which causes cardiac remodeling and leads to congestive heart failure. Though the risks of uncorrected MR and its poor prognosis are known, the longitudinal changes in cardiac function, structure and remodeling are incompletely understood. This knowledge gap has limited our understanding of the optimal timing for MR correction, and the benefit that early versus late MR correction may have on the left ventricle. To investigate the molecular mechanisms that underlie left ventricular remodeling in the setting of MR, animal models are necessary. Traditionally, the aorto-caval fistula model has been used to induce volume overload, which differs from clinically relevant lesions such as MR. MR represents a low-pressure volume overload hemodynamic stressor, which requires animal models that mimic this condition. Herein, we describe a rodent model of severe MR in which the anterior leaflet of the rat mitral valve is perforated with a 23G needle, in a beating heart, with echocardiographic image guidance. The severity of MR is assessed and confirmed with echocardiography, and the reproducibility of the model is reported.&#160;&#160;

A rodent model of left heart volume overload from mitral regurgitation is reported. Mitral regurgitation of controlled severity is induced by advancing a needle of defined dimensions into the anterior leaflet of the mitral valve, in a beating heart, with ultrasound guidance.&#160;

Mitral regurgitation (MR) is a widely prevalent heart valve lesion, which causes cardiac remodeling and leads to congestive heart failure. Though the ...

An Intact Pericardium Ischemic Rodent Model

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to induce myocardial injury). The majority of pre-clinical myocardial infarction models require the disruption of pericardial integrity with loss of the homeostatic cellular milieu. However, recently a methodology has been developed by us to induce myocardial infarction, which minimizes pericardial damage and retains the heart&#39;s resident immune cell population. An improved cardiac functional recovery in mice with an intact pericardial space following coronary ligation has been observed. This method provides an opportunity to study inflammatory responses in the pericardial space following myocardial infarction. Further development of the labeling techniques can be combined with this model to understand the fate and function of pericardial immune cells in regulating the inflammatory mechanisms that drive remodeling in the heart, including fibrosis.

This protocol outlines the steps for inducing myocardial infarction in mice while preserving the pericardium and its contents.

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to ...

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.