This work was supported by grants from the National Heart and Lung Institute (R01 HL086582) and the New York Stem Cell Science Fund (NYSTEM, Contract#&#160;C024316) to S.T.A. and D.D.S. Illustrations used in JoVE video were completed by John Nyquist; Medical Illustrator from State University of New York at Buffalo.

NOTE: This protocol has been approved by the Animal Care and Use Committee at the State University of New York at Buffalo.
1. Pre-surgical Preparation
<ol>
	<li>Use sheep (Dorset cross, female, approximately 1-3 years old with a weight of 40-60 kg) for the following study. Administer cyclosporine A (200 mg/day), aspirin (975 mg/day), and coumadin (20-30 mg/day) by mouth, starting 3 days before surgery and continue for the duration of all studies.</li>
	<li>Ensure the sheep has fasted 12 hr before surgery (normal feed = 1.8 kg hay and 0.4 kg grain).</li>
	<li>Sterilize all surgical supplies using a steam autoclave at 121 &#176;C and 15 psi for 30 min.</li>
	<li>Place the following items in a paraformaldehyde desiccator 48 hr prior to surgery: 4 mm Doppler flow probes, 1 mm ultrasonic crystals, indwelling arterial catheters, extension tubing, and 42&#8221; Tygon Tubing. 
	NOTE: A table of relevant medical equipment needed for the surgery and for sterilization is listed in Table 1. Pre-placement of the instrumentation into the Tygon tubing at this step will help save time during surgery.
	<ol>
		<li>Label each end of the probes, catheters and crystals as left and right, if applicable.</li>
	</ol>
	</li>
</ol>
2. Surgical Operation
<ol>
	<li>Induce sheep for anesthesia with diazepam (0.5 mg/kg) and ketamine (4 mg/kg) intravenously (IV). Alternatively, use Telazol (4 mg/kg) IV.
	<ol>
		<li>Perform orotracheal intubation with an 8.5-10.0 mm internal diameter cuffed endotracheal tube7.</li>
		<li>Administer inhalant&#160;anesthesia through a rebreathing circuit with a tidal volume ventilator (7-10 ml/kg) or a pressure-regulated ventilator (15-20 cm H2O). Use a precision vaporizer to administer isoflurane or sevoflurane at an appropriate amount (3%-4% initially) to reach a medium to deep surgical anesthetic stage. The minimum alveolar concentration for sheep is 1.4% or 1.9% respectively8.</li>
		<li>Assess anesthetic depth by observing motor response to stimuli, palpebral reflex, eye position, and heart rate. Monitor blood oxygen saturation (95%-100%) using pulse oximetry, CO2 concentration (45-55 mmHg) in expiratory gases using capnography and maintain body temperature during the procedure (38.5-39.5 &#176;C) using an auto regulated warming blanket.</li>
	</ol>
	</li>
	<li>Shave wool from the entire neck of the sheep, and over one cephalic vein, using a #40 blade on standard clippers. Prepare the skin of both sites for surgery using 70% isopropyl alcohol and 7.5% betadine scrub saturated gauze. Begin with alcohol gauze to facilitate removal of skin oils. Alternate between betadine gauze and alcohol gauze three times.</li>
	<li>Place sheep onto the operation table in dorsal recumbency on top of a warming blanket. Pass a medium sized orogastric tube to allow for passive expulsion of stomach contents. Extend the sheep&#8217;s neck and use supportive cushioning as needed to maintain placement.
	<ol>
		<li>Perform a final aseptic scrub using 7.5% betadine soaked gauze and allow to sit for 5 min prior to surgery.</li>
	</ol>
	</li>
	<li>Administer IV fluids (Lactated Ringer&#8217;s Solution or 0.9% Saline) at 10 ml/kg/hr through an angiocath placed in the cephalic vein. Administer intraoperative antibiotics and analgesia: Penicillin G Procaine 6,600 U/kg intramuscularly (IM), Gentamicin 1.6 mg/kg IM, and Buprenorphine 0.005-0.01 mg IV or IM.</li>
	<li>Make a ~12 cm incision lengthwise over the ventral midline neck using electro cautery. Isolate left and right carotid arteries (~6 cm) by removing connective tissue using a blunt dissection technique. Tie off and cauterize micro-vessels branching from the carotid arteries to minimize bleeding.</li>
	<li>Maintain sterility by utilizing (a non-sterile) surgical nurse to assist with burrowing all wiring and tubing (flow probe, ultrasonic crystal wires and catheter tubing) in the subcutaneous layer of the skin. Use a blunted trocar, which exits through the surgically prepped incision at the dorsolateral neck.
	<ol>
		<li>Reach under the sterile drape and turn the head of the sheep so that the side of the neck can be visualized under the drape.</li>
		<li>Use an 8 cm curved hemostat to tunnel through the subcutaneous space between the ventral midline neck incision and the side of the neck. Open and close the hemostat to bluntly dissect a space for the tubing ~1.5 cm wide. The tips of the hemostat should reside halfway between the head and shoulders, approximately 10 cm caudal to the right or left ear. Turn the hemostat so that the tips are pointing toward the superficial skin.</li>
		<li>Reach under the sterile drape and make a 1.5 cm incision through the skin, over the tips of the hemostat with a sterile #11 blade. Visualize the tips of the hemostat to confirm a clear exit through the skin.</li>
		<li>Pass the Tygon tube containing all wiring and tubing through the subcutaneous tunnel. Hold the wires and tubing above the sterile drape.</li>
		<li>Reach under the drape to remove the outer Tygon tubing from the neck, exposing the implanted wiring as it leaves the neck of the sheep. Pull individual lines out to minimize any slack in the subcutaneous space. Leave enough distance to properly attach the instrumentation to the artery.</li>
	</ol>
	</li>
	<li>Place 4 mm Doppler flow probes on both carotid arteries and attain an initial reading (Figure 1). Administer 100 U/kg of heparin IV 30 min before clamping the artery.</li>
	<li>Continue heparin administration at 100 U/kg/hr until the end of surgery. Clamp the carotid artery using non-crushing vascular clamps and excise a portion (approximately 4 cm in length). The contralateral carotid flow rate will increase 50%-100% to maintain blood flow to the brain. 
	NOTE: It is possible to limit longitudinal stretch by recoil of native vessel by removing shorter segments than being replaced and/or stretching the native artery with vascular clamps to shorten gap until full anastomotic procedure is completed. This will help limit the tension on individual holding sutures and the implanted graft.</li>
	<li>Suture the TEV in place using simple interrupted stitches with 7-0 proline ethalloy double armed monofilament suture. If necessary, apply vascular smooth muscle relaxants such as Papaverine (15 mg/ml) or Nicardipine (1.25 mg/ml) topically to the native vasculature in order to prevent vasoconstriction which would hamper anastomotic suturing. 
	NOTE: Begin placing sutures with approximately 1 mm spacing. This can greatly vary from case to case. The composition and TEV thickness will affect the effective distance between sutures. As the thickness of the native tissue or TEV decreases, it may be necessary to place the sutures closer together.
	<ol>
		<li>First anchor four points of the TEV to the native artery by placing two opposing stitches on both proximal and distal ends (Figure 2-Di). Hold each anchor taught using hemostats. 
		NOTE: Proximal and distal descriptions are in reference to direction of blood flow throughout the paper.</li>
		<li>Add 5-6 more sutures on the superficial side of both the proximal and distal ends to begin the anastomosis. (Figure 2-Pr). Simultaneously rotate the vascular clamps 180 degrees.</li>
		<li>Re-establish tension on the anchoring sutures. Add additional (5 to 6) interrupted sutures to proximal and distal ends on rotated side of the TEV.</li>
	</ol>
	</li>
	<li>Once the TEV is securely sutured in place, rotate it back to the original position and remove the vascular clamps one at a time, distal clamp first. Slight bleeding at the anastomosis sites is common. This may naturally resolve after several minutes of clamp release and reclamping or require the placement of additional sutures. Place the Doppler flow probe (Figure 3-Fl) back on native artery proximal to blood flow entering the TEV and monitor flow rate. 
	NOTE: Expect the flow rates of the left and right carotid to equilibrate after approximately 15 min. If the flow rate on the carotid artery with implanted TEV steadily drops, it is possible the TEV is clotting. Other possible abnormalities regarding flow can be attributed to constriction of the native arteries proximal or distal to the TEV. If this occurs, the use of additional vascular smooth muscle relaxant may be applied, and the native vessel should return to a basal tone after 30-60 min following the closure of tissue over the graft site.
	<ol>
		<li>If desired, excise the contralateral carotid artery and suture it back into place as a &#8220;Sham&#8221; control. This is more clinically relevant than leaving the right carotid artery alone and only attaching the flow probe, ultrasonic crystals, and catheter. If a sham control is wanted, perform this before continuing to step 2.11.</li>
	</ol>
	</li>
	<li>Suture 1 mm ultrasonic crystals (Figure 3-cr1 cr2) to opposing sides of the TEV using 7-0 Proline. Thread suture through the ultrasonic crystal head and stitch only to the superficial layer of the TEV.</li>
	<li>Catheterize the artery using a modified 18 G catheter with a Teflon woven placket (Figure 3-Ca &#38; Figure 4A). Place the catheter distal to the TEV on native arterial tissue.
	<ol>
		<li>Suture the placket to the arterial wall with 5/0 Ethibond to control any bleeding. Use cyclohexanone to adhere the microbore tubing to the catheter that has been flushed with saline. Use the tubing as an extension line.</li>
		<li>Use a 20 G Luer Stub Adapter with a Surflo Injection Plug to seal the exteriorized end of tubing (Figure 4B). To maintain patency of the catheter, obtain the priming volume of the line and flush it with 10 ml of saline and then 5,000 U/ml of heparin sodium injection every 2-3 days.</li>
	</ol>
	</li>
	<li>Record the distance between the flow probe and the ultrasonic crystals, as well as the distance between the flow probe and the catheter. This will enable pulse wave velocity to be calculated in conjunction with software. If such calculations are not needed, do not implant a catheter.</li>
	<li>Obtain an intraoperative reading if desired to ensure all implanted hardware is functional (see section 3).</li>
	<li>Secure implanted lines and wires to nearby musculature using 2/0 Silk and a taper needle (Figure 3).
	<ol>
		<li>Position the vascular flow probe wire parallel to the vessel, with the probe caudal and the wire extending cranially, and then making a &#8220;U-Turn&#8221; toward the lateral musculature. Secure the wire to adjacent musculature, using 2-0 silk on a taper needle at two locations, so that the wire or flow probe is not able to place any strain on the vessel. Be sure sutures are snug but do not over tighten and strangulate musculature (Figure 3).</li>
		<li>Suture the crystal wires and arterial catheter line to the lateral musculature, allowing for ~1.5 cm of slack, similar to previous steps for securing flow probe (Figure 3).</li>
		<li>Group all wires and lines together and anchor them to musculature just before exiting out through subcutaneous tunnel, similar to previous steps.</li>
	</ol>
	</li>
	<li>Close the surgical site with a 2-0 Vicryl suture in layers using a running suture pattern on facia and subdermal, running mattress stitch on the skin (facia, non-cutting needle; skin, cutting needle). Close the 1.5 cm incision at the dorsal neck around the exteriorized wires and lines using 2/0 Vicryl and a cutting needle.</li>
	<li>Place flow probe wires, catheter lines, and ultrasonic crystal wires into a pouch (10 cm x 10 cm) that is securely sutured to the skin of the sheep (Figure 5 &#8212; after recovery).</li>
	<li>Gradually wean the sheep off anesthesia and the tidal volume ventilator then extubate the sheep when spontaneous breathing is resumed. Remove the angiocath inserted into the cephalic vein and bandage if needed.</li>
	<li>Bandage the neck using triple antibiotic ointment on the incisions, a Telfa pad, stretch roll gauze, and elasticon.</li>
	<li>Administer post-operative analgesia: flunixin meglumine 2.2 mg/kg IM once during recovery, then 1.1 mg/kg IM once a day for two days, buprenorphine 0.005-0.01 mg IV or&#160;IM twice a day for one day.</li>
</ol>
3. In Vivo Monitoring
<ol>
	<li>Place sheep into mobile cart to ensure proper restraint. This allows the sheep to remain calm and conscious without compromising the hardware. May need to acclimate sheep to cart 2 or 3 times for 30 min prior to obtaining instrumentation recordings.</li>
	<li>Remove all wires and lines from pouch and connect to the monitoring devices. Connect flow probe to a Flowmeter, 1 mm ultrasonic crystals connected to TRB-USB box, and catheter lines to pressure transducers. A flow chart of this setup is provided (Figure 6).</li>
	<li>Calibrate flow probes and pressure transducers prior to data acquisition. 
	NOTE: Due to potential variability between software versions and differences in equipment used, calibrations and settings will vary from case to case.</li>
	<li>Utilize an oscilloscope to fine tune the Sonometrics crystal measurement, according to manufacturer&#8217;s protocol.</li>
	<li>Record data using computer software (Figure 7). Traces in the top half of Figure 7 in white color corresponds to the implanted TEV, while traces in the bottom half in red color corresponds to the Sham/Native.&#160;For both the TEV and Sham the flow rate (ml/min), arterial blood pressure (mm Hg) and diameter (mm) are recorded live.</li>
	<li>Record for at least 1 min with no disturbances. Export this data for more detailed analysis. After the recording, disconnect all wires and place back into pouch sutured on the neck of the sheep.</li>
</ol>

<ol>
	<li>Goldman, 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=Long-term+patency+of+saphenous+vein+and+left+internal+mammary+artery+grafts+after+coronary+artery+bypass+surgery:+Results+from+a+Department+of+Veterans+Affairs+Cooperative+Study.">Long-term patency of saphenous vein and left internal mammary artery grafts after coronary artery bypass surgery: Results from a Department of Veterans Affairs Cooperative Study.</a> Journal of the American College of Cardiology. 44, 2149-2156 (2004).</li><li>Achouh, P., 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=Long-term+(5-+to+20-year)+patency+of+the+radial+artery+for+coronary+bypass+grafting.">Long-term (5- to 20-year) patency of the radial artery for coronary bypass grafting.</a> The Journal of Thoracic And Cardiovascular Surgery. 140, 73-79 (2010).</li><li>Conklin, B. S., Richter, E. R., Kreutziger, K. L., Zhong, D. S., Chen, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+evaluation+of+a+novel+decellularized+vascular+xenograft.">Development and evaluation of a novel decellularized vascular xenograft.</a> Medical Engineering &amp; Physics. 24, 173-183 (2002).</li><li>Zhu, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+anti-atherosclerotic+tissue-engineered+blood+vessel+by+A20-regulated+endothelial+progenitor+cells+seeding+decellularized+vascular+matrix.">Development of anti-atherosclerotic tissue-engineered blood vessel by A20-regulated endothelial progenitor cells seeding decellularized vascular matrix.</a> Biomaterials. 29, 2628-2636 (2008).</li><li>Quint, C., 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=Decellularized+tissue-engineered+blood+vessel+as+an+arterial+conduit.">Decellularized tissue-engineered blood vessel as an arterial conduit.</a> Proceedings of the National Academy of Sciences. 108, 9214-9219 (2011).</li><li>Kaushal, 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=Functional+small-diameter+neovessels+created+using+endothelial+progenitor+cells+expanded+ex+vivo.">Functional small-diameter neovessels created using endothelial progenitor cells expanded ex vivo.</a> Nat Med. 7, 1035-1040 (2001).</li><li>Galatos, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anesthesia+and+Analgesia+in+Sheep+and+Goats.">Anesthesia and Analgesia in Sheep and Goats.</a> Veterinary Clinics of North America: Food Animal Practice. 27, 47-59 (2011).</li><li>Okutomi, T., Whittington, R. A., Stein, D. J., Morishima, H. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+the+effects+of+sevoflurane+and+isoflurane+anesthesia+on+the+maternal-fetal+unit+in+sheep.">Comparison of the effects of sevoflurane and isoflurane anesthesia on the maternal-fetal unit in sheep.</a> J Anesth. 23, 392-398 (2009).</li><li>Swartz, D. D., Russell, J. A., Andreadis, S. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineering+of+fibrin-based+functional+and+implantable+small-diameter+blood+vessels.">Engineering of fibrin-based functional and implantable small-diameter blood vessels.</a> American Journal of Physiology - Heart and Circulatory Physiology. 288, H1451-H1460 (2005).</li><li>Niklason, L. 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=Functional+arteries+grown+in+vitro.">Functional arteries grown in vitro.</a> Science. 284, 489-493 (1999).</li><li>Dahl, S. L. 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=Readily+Available+Tissue-Engineered+Vascular+Grafts.">Readily Available Tissue-Engineered Vascular Grafts.</a> Science Translational Medicine. 3, 68ra69 (2011).</li><li>Wu, W., Allen, R. A., Wang, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast-degrading+elastomer+enables+rapid+remodeling+of+a+cell-free+synthetic+graft+into+a+neoartery.">Fast-degrading elastomer enables rapid remodeling of a cell-free synthetic graft into a neoartery.</a> Nature Medicine. 18, 1148-1153 (2012).</li><li>Saami, K. Y., Bryan, W. T., Joel, L. B., Shay, S., Randolph, L. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+fate+of+an+endothelium+layer+after+preconditioning.">The fate of an endothelium layer after preconditioning.</a> Journal of vascular surgery : Official Publication, the Society for Vascular Surgery [and] International Society for Cardiovascular Surgery, North American Chapter. 51, 174-183 (2010).</li><li>Ueberrueck, T., 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=Comparison+of+the+ovine+and+porcine+animal+models+for+biocompatibility+testing+of+vascular+prostheses.">Comparison of the ovine and porcine animal models for biocompatibility testing of vascular prostheses.</a> Journal of Surgical Research. 124, 305-311 (2005).</li><li>Labb&#233;, R., Germain, L., Auger, 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=A+completely+biological+tissue-engineered+human+blood+vessel.">A completely biological tissue-engineered human blood vessel.</a> The FASEB Journal. 12, 47-56 (1998).</li><li>Samaha, F. J., Oliva, A., Buncke, G. M., Buncke, H. J., Siko, P. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+clinical+study+of+end-to-end+versus+end-to-side+techniques+for+microvascular+anastomosis.">A clinical study of end-to-end versus end-to-side techniques for microvascular anastomosis.</a> Plastic and Reconstructive Surgery. 99, 1109-1111 (1997).</li><li>Huang, 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=A+novel+end-to-side+anastomosis+technique+for+hepatic+rearterialization+in+rat+orthotopic+liver+transplantation+to+accommodate+size+mismatches+between+vessels.">A novel end-to-side anastomosis technique for hepatic rearterialization in rat orthotopic liver transplantation to accommodate size mismatches between vessels.</a> European Surgical Research. 47, 53-62 (2011).</li><li>Peng, H., Schlaich, E. M., Row, S., Andreadis, S. T., Swartz, D. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Novel+Ovine+ex+vivo+Arteriovenous+Shunt+Model+to+Test+Vascular+Implantability.">A Novel Ovine ex vivo Arteriovenous Shunt Model to Test Vascular Implantability.</a> Cells, Tissues, Organs. 195, 108 (2011).</li><li>Zilla, P., Bezuidenhout, D., Human, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prosthetic+vascular+grafts:+Wrong+models,+wrong+questions+and+no+healing.">Prosthetic vascular grafts: Wrong models, wrong questions and no healing.</a> Biomaterials. 28, 5009-5027 (2007).</li><li>Berger, K., Sauvage, L. R., Rao, A. M., Wood, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Healing+of+Arterial+Prostheses+in+Man:+Its+Incompleteness.">Healing of Arterial Prostheses in Man: Its Incompleteness.</a> Annals of Surgery. 175, 118-127 (1972).</li><li>Byrom, M. J., Bannon, P. G., White, G. H., Ng, M. K. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+for+the+assessment+of+novel+vascular+conduits.">Animal models for the assessment of novel vascular conduits.</a> Journal of Vascular Surgery : Official Publication, the Society for Vascular Surgery [and] International Society for Cardiovascular Surgery, North American Chapter. 52, 176-195 (2010).</li><li>Swartz, D. D., Andreadis, S. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+for+vascular+tissue-engineering.">Animal models for vascular tissue-engineering.</a> Current Opinion in Biotechnology. 24, 916-925 (2013).</li><li>Liang, M. -. S., Andreadis, S. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineering+fibrin-binding+TGF-&#946;1+for+sustained+signaling+and+contractile+function+of+MSC+based+vascular+constructs.">Engineering fibrin-binding TGF-&#946;1 for sustained signaling and contractile function of MSC based vascular constructs.</a> Biomaterials. 32, 8684-8693 (2011).</li><li>Han, J., Liu, J. Y., Swartz, D. D., Andreadis, S. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+and+functional+effects+of+organismal+ageing+on+smooth+muscle+cells+derived+from+bone+marrow+mesenchymal+stem+cells.">Molecular and functional effects of organismal ageing on smooth muscle cells derived from bone marrow mesenchymal stem cells.</a> Cardiovascular Research. 87, 147-155 (2010).</li></ol>

There are no disclosures to report.

The purpose of this report is to provide a reliable and reproducible procedure to implant TEVs of interest in the ovine carotid artery. The native carotid arteries of the animals used in this model were 0.5-0.75 mm in thickness and 4.5-5 mm in outside diameter. The surgical technique described here has been successful for implanting TEVs of varying geometries measuring 0.25-1 mm in thickness, 4-5 mm outside diameter and 4 cm in length with great success proving effective up to 3 months duration, the intended end point. T...

The primary focus for the development of TEVs has been to provide a substitute for autologous graft replacement when autologous vessels are not available and to limit donor sight morbidity. For example, the number of coronary artery bypass surgeries per year has exceeded 350,000 in the USA, and the ideal source of suitable grafts remains the left internal mammary artery, left anterior descending coronary artery and saphenous vein1. Since many individuals who suffer from vascular diseases may not have suitable arteries and veins for autologous graft replacement, the development of TEVs has thus become an intense field of research for decades1-6. While the engineering and optimization of novel TEVs have undergone many advancements, reporting on the surgical techniques employed to implant the TEVs themselves has not been a topic of such intense discussion. Rather, protocols regarding the implantation of TEVs into animal models are largely left up to research investigators.
The following manuscript demonstrates how to implant TEVs by utilizing an end-to-end anastomosis approach. This procedure was optimized by using a specific anastomotic suturing pattern, stabilizing suture technique, optimizing longitudinal tension and the addition of in vivo monitoring instrumentation. This method is contrasted with some of the many variations that have been previously used. Furthermore, this procedure describes how to acquire parameters such as arterial blood pressure, TEV diameter/compliance and flow rate through the TEV after surgery up until explantation. This data collection provides an indispensable analysis of the TEV while it is in the process of remodeling.

<table><tbody><tr><td>Pressure Transducer</td><td>Becton Dickinson</td><td>P23XL-1</td><td>Quantity: 1+ (1 for each artery).&lt;br/&gt;Used with water-filled diaphragm domes.</td></tr><tr><td>Amplifier and transducer box</td><td>Gould</td><td>5900 Signal Conditioner Cage</td><td>Quantity: 1.&lt;br/&gt;Two transducers and amplifiers should be included in cage. While this specific unit may be discontinued, other commercially available pressure transducers with a BNC/analog output will communicate with the Sonometrics equipment.</td></tr><tr><td>T403 Console with TS420 perivascular flowmeter module (x2)</td><td>Transonic Systems</td><td>T403 module and TS420 (x2)</td><td>Quantity: 1.&lt;br/&gt;Flow probes measuring flow through each of the carotid arteries will connect to each of the TS420 units.</td></tr><tr><td>Digital ultrasonic measurement unit</td><td>Sonometrics</td><td>TR-USB</td><td>Quantity: 1</td></tr><tr><td>Flow Probe Precision S-Series 4&amp;nbsp;mm</td><td>Transonic Systems Inc.</td><td>MC4PSS-LS-WC100-CM4B-GA</td><td>Quantity: 2</td></tr><tr><td>1&amp;nbsp;mm Sonometrics Crystals</td><td>Sonometrics Systems</td><td>1R-38S-20-NC-SH</td><td>Quantity: 2-4 (2 for each artery)</td></tr><tr><td>Catheter for implantation</td><td>BD (Becton Dickinson)</td><td>&amp;nbsp;381447</td><td>Quantity: 1+ (1 for each artery).&lt;br/&gt;Catheter is cut and secured to microbore tubing, stylette is utilized for insertion.</td></tr><tr><td>Tygon Microbore Tubing</td><td>Norton Performance Plastics</td><td>(AAQ04127) Formulation S-54-HL</td><td>Cut to length for an extension set</td></tr><tr><td>Luer Stub Adapter</td><td>BD (Becton Dickinson)</td><td>427564 (20 gauge)</td><td>Quantity: 1+ (1 for each arterial catheter)</td></tr><tr><td>Surflo Injection Plug</td><td>Terumo</td><td>SR-IP2</td><td>Quantity: 1+ (1 for each arterial catheter)</td></tr><tr><td>Meadox</td><td>PTFE (Teflon) Felt</td><td>19306</td><td>Cut to size.&lt;br/&gt;The PTFE felt used in our studies was discontinued. However, comparable companies such as &amp;ldquo;Surgical Mesh&amp;rdquo; offer products which are equivalent.</td></tr></tbody></table>

surgical technique for the implantation of tissue engineered vascular grafts and subsequent in vivo monitoring

The development of Tissue Engineered Vessels (TEVs) is advanced by the ability to routinely and effectively implant TEVs (4-5 mm in diameter) into a large animal model. A step by-step protocol for inter-positional placement of the TEV and real-time digital assessment of the TEV and native carotid arteries is described here. In vivo monitoring is made possible by the implantation of flow probes, catheters and ultrasonic crystals (capable of recording dynamic diameter changes of implanted TEVs and native carotid arteries) at the time of surgery. Once implanted, researchers can calculate arterial blood flow patterns, invasive blood pressure and artery diameter yielding parameters such as pulse wave velocity, augmentation index, pulse pressures and compliance. Data acquisition is accomplished using a single computer program for analysis throughout the duration of the experiment. Such invaluable data provides insight into TEV matrix remodeling, its resemblance to native/sham controls and overall TEV performance in vivo.

More than 30 sheep have undergone the surgical technique described in this report for the implantation of TEVs (in press) 9. A table summarizing the most recent sheep operations after protocol optimization are shown in Table 2. All sheep recovered after TEV implantation with no life threatening complications. In some animals, fibrosis was observed in native artery near the tip of the indwelling arterial catheter. A significant increase of inflammation with the presence of added instrumentation...

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Surgical Technique for the Implantation of Tissue Engineered Vascular Grafts and Subsequent In Vivo Monitoring

A step-by-step protocol for the inter-positional placement of Tissue Engineered Vessels (TEVs) into the carotid artery of a sheep using end-to-end anastomosis and real-time digital assessment in vivo until animal sacrifice.

Surgical Technique for the Implantation of Tissue Engineered Vascular Grafts and Subsequent In Vivo Monitoring

The development of Tissue Engineered Vessels (TEVs) is advanced by the ability to routinely and effectively implant TEVs (4-5 mm in diameter) into a large ...

surgical-technique-for-implantation-tissue-engineered-vascular-grafts

Research

JoVE Journal

Bioengineering

11.7K Views.  State University of New York Buffalo School of Medicine. A step-by-step protocol for the inter-positional placement of Tissue Engineered Vessels (TEVs) into the carotid artery of a sheep using end-to-end anastomosis and real-time digital assessment in vivo until animal sacrifice.

Video: Surgical Technique for the Implantation of Tissue Engineered Vascular Grafts and Subsequent In Vivo Monitoring

Engineering Biological-Based Vascular Grafts Using a Pulsatile Bioreactor

Much effort has been devoted to develop and advance the methodology to regenerate functional small-diameter arterial bypasses. In the physiological environment, both mechanical and chemical stimulation are required to maintain the proper development and functionality of arterial vessels1,2.
Bioreactor culture systems developed by our group are designed to support vessel regeneration within a precisely controlled chemo-mechanical environment mimicking that of native vessels. Our bioreactor assembly and maintenance procedures are fairly simple and highly repeatable3,4. Smooth muscle cells (SMCs) are seeded onto a tubular polyglycolic acid (PGA) mesh that is threaded over compliant silicone tubing and cultured in the bioreactor with or without pulsatile stimulation for up to 12 weeks. There are four main attributes that distinguish our bioreactor from some predecessors. 1) Unlike other culture systems that simulate only the biochemical surrounding of native blood vessels, our bioreactor also creates a physiological pulsatile environment by applying cyclic radial strain to the vessels in culture. 2) Multiple engineered vessels can be cultured simultaneously under different mechanical conditions within a controlled chemical environment. 3) The bioreactor allows a mono layer of endothelial cells (EC) to be easily coated onto the luminal side of engineered vessels for animal implantation models. 4) Our bioreactor can also culture engineered vessels with different diameter size ranged from 1 mm to 3 mm, saving the effort to tailor each individual bioreactor to fit a specific diameter size. 
The engineered vessels cultured in our bioreactor resemble native blood vessels histologically to some degree. Cells in the vessel walls express mature SMC contractile markers such as smooth muscle myosin heavy chain (SMMHC)3. A substantial amount of collagen is deposited within the extracellular matrix, which is responsible for ultimate mechanical strength of the engineered vessels5. Biochemical analysis also indicates that collagen content of engineered vessels is comparable to that of native arteries6. Importantly, the pulsatile bioreactor has consistently regenerated vessels that exhibit mechanical properties that permit successful implantation experiments in animal models3,7. Additionally, this bioreactor can be further modified to allow real-time assessment and tracking of collagen remodeling over time, non-invasively, using a non-linear optical microscopy (NLOM)8. To conclude, this bioreactor should serve as an excellent platform to study the fundamental mechanisms that regulate the regeneration of functional small-diameter vascular grafts.

Our group has developed a bioreactor culture system that mimics the physiological pulsatile stresses of the cardiovascular system to regenerate implantable small-diameter vascular grafts.

Much effort has been devoted to develop and advance the methodology to regenerate functional small-diameter arterial bypasses. In the physiological ...

A Full Skin Defect Model to Evaluate Vascularization of Biomaterials In Vivo

Insufficient vascularization is considered to be one of the main factors limiting the clinical success of tissue-engineered constructs. In order to evaluate new strategies that aim at improving vascularization, reliable methods are required to make the in-growth of new blood vessels into bio-artificial scaffolds visible and quantify the results. Over the past couple of years, our group has introduced a full skin defect model that enables the direct visualization of blood vessels by transillumination and provides the possibility of quantification through digital segmentation. In this model, one surgically creates full skin defects in the back of mice and replaces them with the material tested. Molecules or cells of interest can also be incorporated in such materials to study their potential effect. After an observation time of one&#8217;s own choice, materials are explanted for evaluation. Bilateral wounds provide the possibility of making internal comparisons that minimize artifacts among individuals as well as of decreasing the number of animals needed for the study. In comparison to other approaches, our method offers a simple, reliable and cost effective analysis. We have implemented this model as a routine tool to perform high-resolution screening when testing vascularization of different biomaterials and bio-activation approaches.

A Full Skin Defect Model to Evaluate Vascularization of Biomaterials In Vivo

Vascularization is key to approaches in successful tissue engineering. Therefore, reliable technologies are required to evaluate the development of vascular networks in tissue-constructs. Here we present a simple and cost-effective method to visualize and quantify vascularization in vivo.

Insufficient vascularization is considered to be one of the main factors limiting the clinical success of tissue-engineered constructs. In order to ...

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

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

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

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

Engineered Vascularized Muscle Flap

One of the main factors limiting the thickness of a tissue construct and its consequential viability and applicability in vivo, is the control of oxygen supply to the cell microenvironment, as passive diffusion is limited to a very thin layer. Although various materials have been described to restore the integrity of full-thickness defects of the abdominal wall, no material has yet proved to be optimal, due to low graft vascularization, tissue rejection, infection, or inadequate mechanical properties. This protocol describes a means of engineering a fully vascularized flap, with a thickness relevant for muscle tissue reconstruction. Cell-embedded poly L-lactic acid/poly lactic-co-glycolic acid constructs are implanted around the mouse femoral artery and vein and maintained in vivo for a period of one or two weeks. The vascularized graft is then transferred as a flap towards a full thickness defect made in the abdomen. This technique replaces the need for autologous tissue sacrifications and may enable the use of in vitro engineered vascularized flaps in many surgical applications.

To date, thick tissue defects are typically reconstructed by applying autologous tissue flaps or engineered tissues. In this protocol, we present a new method for engineering vascularized tissue flap bearing an autologous pedicle, to serve as a substitute to autologous flaps.

One of the main factors limiting the thickness of a tissue construct and its consequential viability and applicability in vivo, is the control of oxygen ...

Computed Tomography and Optical Imaging of Osteogenesis-angiogenesis Coupling to Assess Integration of Cranial Bone Autografts and Allografts

A major parameter determining the success of a bone-grafting procedure is vascularization of the area surrounding the graft. We hypothesized that implantation of a bone autograft would induce greater bone regeneration by abundant blood vessel formation. To investigate the effect of the graft on neovascularization at the defect site, we developed a micro&#8211;computed tomography (&#181;CT) approach to characterize newly forming blood vessels, which involves systemic perfusion of the animal with a polymerizing contrast agent. This method enables detailed vascular analysis of an organ in its entirety. Additionally, blood perfusion was assessed using fluorescence imaging (FLI) of a blood-borne fluorescent agent. Bone formation was quantified by FLI using a hydroxyapatite-targeted probe and &#181;CT analysis. Stem cell recruitment was monitored by bioluminescence imaging (BLI) of transgenic mice that express luciferase under the control of the osteocalcin promoter. Here we describe and demonstrate preparation of the allograft, calvarial defect surgery, &#181;CT scanning protocols for the neovascularization study and bone formation analysis (including the in vivo perfusion of contrast agent), and the protocol for data analysis.
The 3D high-resolution analysis of vasculature demonstrated significantly greater angiogenesis in animals with implanted autografts, especially with respect to arteriole formation. Accordingly, blood perfusion was significantly higher in the autograft group by the 7th day after surgery. We observed superior bone mineralization and measured greater bone formation in animals that received autografts. Autograft implantation induced resident stem cell recruitment to the graft-host bone suture, where the cells differentiated into bone-forming cells between the 7th and 10th postoperative day. This finding means that enhanced bone formation may be attributed to the augmented vascular feeding that characterizes autograft implantation. The methods depicted may serve as an optimal tool to study bone regeneration in terms of tightly bounded bone formation and neovascularization.

Implantation of autologous and allogeneic bone grafts constitute accepted approaches to treat major craniofacial bone loss. Yet the effect of graft composition on the interplay among neovascularization, cell differentiation and bone formation is unclear. We present a multimodal imaging protocol aimed to elucidate the angiogenesis-osteogenesis interdependence at the graft proximity.

A major parameter determining the success of a bone-grafting procedure is vascularization of the area surrounding the graft. We hypothesized that ...

Implantation of Inferior Vena Cava Interposition Graft in Mouse Model

Biodegradable scaffolds seeded with bone marrow mononuclear cells (BMCs) are often used for reconstructive surgery to treat congenital cardiac anomalies. The long-term clinical results showed excellent patency rates, however, with significant incidence of stenosis. To investigate the cellular and molecular mechanisms of vascular neotissue formation and prevent stenosis development in tissue engineered vascular grafts (TEVGs), we developed a mouse model of the graft with approximately 1 mm internal diameter. First, the TEVGs were assembled from biodegradable tubular scaffolds fabricated from a polyglycolic acid nonwoven felt mesh coated with &#949;-caprolactone and L-lactide copolymer. The scaffolds were then placed in a lyophilizer, vacuumed for 24 hr, and stored in a desiccator until cell seeding. Second, bone marrow was collected from donor mice and mononuclear cells were isolated by density gradient centrifugation. Third, approximately one million cells were seeded on a scaffold and incubated O/N. Finally, the seeded scaffolds were then implanted as infrarenal vena cava interposition grafts in C57BL/6 mice. The implanted grafts demonstrated excellent patency (&#62;90%) without evidence of thromboembolic complications or aneurysmal formation. This murine model will aid us in understanding and quantifying the cellular and molecular mechanisms of neotissue formation in the TEVG.

To improve our knowledge of cellular and molecular neotissue formation, a murine model of the TEVG was recently developed. The grafts were implanted as infrarenal vena cava interposition grafts in C57BL/6 mice. This model achieves similar results to those achieved in our clinical investigation, but over a far shortened time-course.

Biodegradable scaffolds seeded with bone marrow mononuclear cells (BMCs) are often used for reconstructive surgery to treat congenital cardiac anomalies. ...

Medicine

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and trade one defect for another. Tissue engineering holds the promise to address this increasing demand. However, most tissue engineering strategies fail to generate stable and functional tissue substitutes because of poor vascularization. This paper focuses on an in vivo tissue engineering chamber model of intrinsic vascularization where a perfused artery and a vein either as an arteriovenous loop or a flow-through pedicle configuration is directed inside a protected hollow chamber. In this chamber-based system angiogenic sprouting occurs from the arteriovenous vessels and this system attracts ischemic and inflammatory driven endogenous cell migration which gradually fills the chamber space with fibro-vascular tissue. Exogenous cell/matrix implantation at the time of chamber construction enhances cell survival and determines specificity of the engineered tissues which develop. Our studies have shown that this chamber model can successfully generate different tissues such as fat, cardiac muscle, liver and others. However, modifications and refinements are required to ensure target tissue formation is consistent and reproducible. This article describes a standardized protocol for the fabrication of two different vascularized tissue engineering chamber models in vivo.

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

This is a guideline for constructing in vivo vascularized tissue using a microsurgical arteriovenous loop or a flow-through pedicle configuration inside a tissue engineering chamber. The vascularized tissues generated can be employed for organ regeneration and replacement of tissue defects, as well as for drug testing and disease modeling.

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and ...

The Arteriovenous (AV) Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

A functional blood vessel network is a prerequisite for the survival and growth of almost all tissues and organs in the human body. Moreover, in pathological situations such as cancer, vascularization plays a leading role in disease progression. Consequently, there is a strong need for a standardized and well-characterized in vivo model in order to elucidate the mechanisms of neovascularization and develop different vascularization approaches for tissue engineering and regenerative medicine.
We describe a microsurgical approach for a small animal model for induction of a vascular axis consisting of a vein and artery that are anastomosed to an arteriovenous (AV) loop. The AV loop is transferred to an enclosed implantation chamber to create an isolated microenvironment in vivo, which is connected to the living organism only by means of the vascular axis. Using 3D imaging (MRI, micro-CT) and immunohistology, the growing vasculature can be visualized over time. By implanting different cells, growth factors and matrices, their function in blood vessel network formation can be analyzed without any disturbing influences from the surroundings in a well controllable environment. 
In addition to angiogenesis and antiangiogenesis studies, the AV loop model is also perfectly suited for engineering vascularized tissues. After a certain prevascularization time, the generated tissues can be transplanted into the defect site and microsurgically connected to the local vessels, thereby ensuring immediate blood supply and integration of the engineered tissue. By varying the matrices, cells, growth factors and chamber architecture, it is possible to generate various tissues, which can then be tailored to the individual patient's needs.

The Arteriovenous AV Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

We describe a microsurgical approach for the generation of an arteriovenous (AV) loop as a model for analyzing vascularization in vivo in an isolated and well-characterized environment. This model is not only useful for the investigation of angiogenesis, but is also optimally suited for engineering axially vascularized and transplantable tissues.

A functional blood vessel network is a prerequisite for the survival and growth of almost all tissues and organs in the human body. Moreover, in ...

Scaling of Engineered Vascular Grafts Using 3D Printed Guides and the Ring Stacking Method

Coronary artery disease remains a leading cause of death, affecting millions of Americans. With the lack of autologous vascular grafts available, engineered grafts offer great potential for patient treatment. However, engineered vascular grafts are generally not easily scalable, requiring manufacture of custom molds or polymer tubes in order to customize to different sizes, constituting a time-consuming and costly practice. Human arteries range in lumen diameter from about 2.0-38 mm and in wall thickness from about 0.5-2.5 mm. We have created a method, termed the "Ring Stacking Method," in which variable size rings of tissue of the desired cell type, demonstrated here with vascular smooth muscle cells (SMCs), can be created using guides of center posts to control lumen diameter and outer shells to dictate vessel wall thickness. These tissue rings are then stacked to create a tubular construct, mimicking the natural form of a blood vessel. The vessel length can be tailored by simply stacking the number of rings required to constitute the length needed. With our technique, tissues of tubular forms, similar to a blood vessel, can be readily manufactured in a variety of dimensions and lengths to meet the needs of the clinic and patient.

Scalable engineered blood vessels would improve clinical applicability. Using easily sizable 3D-printed guides, rings of vascular smooth muscle were created and stacked into a tubular form, forming a vascular graft. Grafts can be sized to meet the range of human coronary artery dimensions by simply changing the 3D-printed guide size.

Coronary artery disease remains a leading cause of death, affecting millions of Americans. With the lack of autologous vascular grafts available, ...

Nondestructive Monitoring of Degradable Scaffold-Based Tissue-Engineered Blood Vessel Development Using Optical Coherence Tomography

Engineered vascular grafts with structural and mechanical properties similar to natural blood vessels are expected to meet the growing demand for arterial bypass. Characterization of the growth dynamics and remodeling process of degradable polymer scaffold-based tissue-engineered blood vessels (TEBVs) with pulsatile stimulation is crucial for vascular tissue engineering. Optical imaging techniques stand out as powerful tools for monitoring vascularization of engineered tissue enabling high-resolution imaging in real-time culture. This paper demonstrates a nondestructive and fast real-time imaging strategy to monitor the growth and remodeling of TEBVs in long-term culture by using optical coherence tomography (OCT). Geometric morphology is evaluated, including vascular remodeling process, wall thickness, and comparison of TEBV thickness in different culture time points and presence of pulsatile stimulation. Finally, OCT provides practical possibilities for real-time observation of the degradation of polymer in the reconstructing tissues under pulsatile stimulation or not and in each vessel segment, by compared with the assessment of polymer degradation using scanning electron microscopic(SEM) and polarized microscope.

A step by step protocol for nondestructive and long-period monitoring the process of vascular remodeling and scaffold degradation in real-time culture of biodegradable polymeric scaffold-based tissue-engineered blood vessels with pulsatile stimulation using optical coherence tomography is described here.

Engineered vascular grafts with structural and mechanical properties similar to natural blood vessels are expected to meet the growing demand for arterial ...

Surgical Technique for the Implantation of Tissue Engineered Vascular Grafts and Subsequent In Vivo Monitoring

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Engineering Biological-Based Vascular Grafts Using a Pulsatile Bioreactor

A Full Skin Defect Model to Evaluate Vascularization of Biomaterials In Vivo

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

Engineered Vascularized Muscle Flap

Computed Tomography and Optical Imaging of Osteogenesis-angiogenesis Coupling to Assess Integration of Cranial Bone Autografts and Allografts

Implantation of Inferior Vena Cava Interposition Graft in Mouse Model

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

The Arteriovenous (AV) Loop in a Small Animal Model to Study Angiogenesis and Vascularized Tissue Engineering

Scaling of Engineered Vascular Grafts Using 3D Printed Guides and the Ring Stacking Method

Nondestructive Monitoring of Degradable Scaffold-Based Tissue-Engineered Blood Vessel Development Using Optical Coherence Tomography